Western Blotting Using In-Gel Protein Labeling as a Normalization Control: Advantages of Stain-Free Technology

Abstract

Western blotting is one of the most used techniques in research laboratories. It is popular because it is an easy way of semi-quantifying protein amounts in different samples. In Western blotting, the most commonly used method for controlling for the differences in the amount of protein loaded is to independently quantify housekeeping proteins (typically actin, GAPDH or tubulin). Another less commonly used method is total protein normalization using stains, such as Ponceau S or Coomassie Brilliant Blue, which stains all the proteins on the blots. A less commonly used but powerful total protein staining technique is stain-free normalization. The stain-free technology is able to detect total protein in a large linear dynamic range and has the advantage of allowing protein detection on the gel before transblotting. This chapter discusses the theory, advantages, and method used to do total protein quantification using stain-free gels for normalization of Western blots.

1. Introduction

Western blotting is widely used in life sciences to access specific protein levels in a sample. It relies on protein separation by size in an acrylamide matrix in the presence of an electric current, followed by the transfer of proteins from the gel to a membrane, the blocking of sites on the membrane that were not exposed to proteins transferred from the gel, the incubation of the membranes with antibodies with high affinity for a target protein, and the detection of these primary antibodies by different methods [1]. Given Western blotting relevance to describe mechanisms for metabolism, disease states, and several other cellular processes, it is important to have both robust and reproducible protocols. Since Western blotting techniques are very sensitive to variations in protein amounts loaded on gels, which can occur from pipetting errors and transfer steps, it is important to ensure the proper normalization of the amount of the target protein through a loading control.

The most common loading controls are the housekeeping proteins [2,3]. These housekeeping proteins are highly abundant proteins that are constitutively expressed by cells in many different states and conditions. Actin, glyceraldehyde 3-phosphate dehydrogenase (GAPDH), and tubulin are some of the most frequent housekeeping proteins used in the literature. However, the use of housekeeping proteins as loading controls has some limitations, and in several cases is not a reliable method to normalize Western blotting data. It has been shown that in many cell states, like proliferation, differentiation, and disease, significant changes in the amount of these housekeeping proteins occur [4–8]. In some reports, the saturation of housekeeping proteins at relatively low total protein amounts leads to misleading results for the target protein amounts, especially when the target proteins are present in low amounts [6,7]. It has been suggested by several reports that the use of total protein amounts is more appropriate to normalize Western blotting data, leading to less variation, more reproducibility, and decreased potential errors associated with different cellular states.

The most used techniques to determine total protein amounts are based on using dyes to stain all proteins on the blot membrane. This can be accessed before the membrane blocking, with Ponceau S [9], of after imaging it, like Coomassie blue and Amido-black staining [10]. These total protein staining methods have their own limitations as shown in the table below (Table 1).

Table 1.

The advantages and disadvantages of different total protein staining methods

Detection Method	Pros	Cons	Refs
Ponceau S	• Easily reversible; • Water soluble; • Staining solutions can be re-used many times.	• Additional steps of washing and shaking; • Low sensitivity (~200 ng minimum); • Recommended only for membranes; • Increased background.	[9,10,12,13]
Coomassie blue	• Good sensitivity (~50 ng minimum); • Suitable both for gels and membranes; • Staining solutions can be re-used many times.	• Toxic/harmful waste; • Additional steps of washing/shaking; • Requires the incubation of weak acid and organic alcohols; • Irreversible staining.	[10,12,14,13]
Amido black	• Good sensitivity (~50 ng minimum); • Suitable both for gels and membranes; • Staining solutions can be re-used many times.	• Toxic/harmful waste; • Additional steps of washing/shaking; • Requires the incubation of weak acid and organic alcohols on the membrane; • Irreversible staining.	[10,12,13]
Stain-free	• Greater sensitivity (~20 ng minimum); • Does not require solutions; • No additional steps of washing/shaking; • No incubation; • Suitable both for gels and membranes; • No reagent waste generated.	• Requires a gel prepared with trihalo compound; • Antibodies that bind to tryptophan residues in the target protein antibody epitope may have its efficiency affected.	[15,16,11]

Stain-free imaging can be a powerful tool to overcome some of the staining protocol limitations. The concept of stain-free imaging of gels and membranes is based the ability of aromatic amino acids like tryptophan to undergo changes under UV light stimulation in the presence of trihalo compounds. Tryptophan present on proteins covalently bind to the trihalo compounds in the gel, resulting in these labelled proteins emitting detectible fluorescence, and enabling the detection of amounts as low as 20 ng of protein [11]. This method is very reliable but requires a gel that was polymerized together with trihalo compound. In this article, we demonstrate the effectiveness of the stain-free method to accurately quantify the total amount of protein in mouse heart homogenates and to normalize the amount of the proteasome subunit β₁ by Western blotting. Hence, the stain-free technology appears to be a robust technique to ensure proper loading controls for protein analysis.

2. Materials

Prepare all solutions with ultrapure water (18 MΩ) and reagents that are analytical grade. Prepare and store all reagents at room temperature unless otherwise indicated. Do not add sodium azide to any of the solutions and follow all local waste disposal protocols for disposal of Western blotting waste.

2.1. Tissue Sample Preparation for SDS-PAGE

1. 100 mg mouse heart.

2. Homogenization buffer: 150 mM NaCl, 5 mM MgCl₂, 1 mM EDTA, 50 mM Tris-HCl, pH 7.5. To prepare 250 mL homogenization buffer, add together 7.5 mL 5 M NaCl, 1.25 mL 1 M MgCl₂, 0.5 mL 0.5 M EDTA, and 6.25 mL 2 M Tris base. Dissolve all reagents in about 200 mL of water. After adjusting the pH to 7.5 with HCl, top up the solution to 250 mL with water (see Note 1).

3. Spectrophotometer for small samples: NanoDrop 2000c.

2.2. SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE)

1. Laemmli sample buffer (4×): 131.6 mM Tris–HCl, pH 6.8, 4.2 % SDS, 52.6 % (w/v) glycerol, 0.02 % (w/v) bromophenol blue. Add β-mercaptoethanol to the sample buffer when ready to use (100 μL for each 900 μL of 4× Laemmli sample buffer) (see Note 2).

2. SDS-PAGE gels with stain-free capabilities: e.g. 4–20 % 26-well Criterion TGX Stain-Free Precast Gel.

3. Protein standards: e.g. Precision Plus Protein Dual Color Standards (see Note 3).

4. 10× SDS-PAGE running buffer: 192 mM glycine, 25 mM Tris base, 0.1 % SDS. To prepare 1 L of 10× SDS-PAGE running buffer, add 144 g glycine, 30.2 g Tris base, and 50 mL 20% SDS solution. Mix the reagents in around 900 mL of water until dissolved, and then fill to 1 L with water (see Note 4).

2.3. Protein Transfer

1. Precut filter paper: e.g. Trans-Blot Turbo Midi-size Transfer Stacks.

2. Blotting membrane: e.g. Trans-Blot Turbo Nitrocellulose membrane (see Note 5).

3. Transfer buffer: e.g. Trans-Blot Turbo 5× Transfer Buffer (Bio-Rad). Prepare 1 L of 1× transfer buffer by mixing 200 mL 5× transfer buffer with 600 mL water and 200 mL 200 proof pure ethanol

4. Blotting apparatus: Trans-Blot Turbo Transfer System.

5. Tris-buffered saline (TBS): 10 mM Tris-HCl, 150 mM NaCl, pH 7.4 (see Note 6). To prepare 1 L of 1× TBS, add 2.4 g Tris base and 8.8 g NaCl to around 800 mL water. Mix thoroughly to dissolve and adjust the pH to 7.4 with HCl. Fill to 1 L with water.

6. TBST: TBS with 0.05 % (v/v) Tween-20. Add 0.5 mL Tween-20 to 1 L TBS to make TBST (see Note 7).

7. Blocking solution: 3 % non-fat milk in TBST. To make 50 mL of blocking solution, weigh out 1.5 g of powdered milk in a 50 mL conical centrifuge tube, and fill to 50 mL with TBST. Vortex the solution until completely mixed (see Note 8).

8. Antibody diluent solution: 1 % non-fat milk in TBST. To make 15 mL of antibody diluent solution, dissolve 0.15 g of powdered milk in 15 mL TBST in a conical centrifuge tube (see Note 8).

9. Substrate for enhanced chemiluminescence (ECL): Clarity Western ECL Substrate, containing Clarity Western peroxidase reagent and Clarity Western luminol/enhancer that should be mixed at a ratio of 1:1 when ready to use (see Note 9).

10. Blotting box (see Note 10).

11. Forceps for gentle handling of membranes.

12. Roller.

13. Delicate wiper paper (see Note 11).

14. Imaging equipment with stain-free technology capabilities: ChemiDoc MP.

15. Computer with Image Lab software, version 6.01 (see Note 12).

2.4. Antibodies for Immunostaining

1. Primary antibody solution: monoclonal mouse 20S Proteasome β1 (D-9) antibody (# sc-374405, Santa Cruz Biotechnology), used at a concentration of 1:2000 in 1 % (v/v) milk/TBST.

2. Secondary antibody solution: peroxidase-conjugated rabbit anti-mouse IgG antibody (# A9044, Sigma-Aldrich,) used at a concentration of 1:10000 (v/v) in 1 % milk/TBST.

3. Methods

Perform the following procedures at room temperature unless otherwise noted.

3.1. Sample Preparation

1. Thaw the mouse heart on ice and chop approximately 50 mg with a razor blade. Use 25 strokes to homogenize the chopped heart in 0.5 mL cold homogenization buffer on ice using a dounce homogenizer (see Note 13).

2. Transfer the homogenate into 1.5 mL centrifuge tubes and centrifuge at 12,000g for 15 min to separate the debris, mitochondria and nucleus. Transfer the supernatant to a new tube and measure the concentration using a NanoDrop Spectrophotometer. Dilute the homogenate to 16 mg/mL with homogenization buffer. Store lysate at −80°C if not using immediately.

3. Prepare the samples for SDS-PAGE. Combine equal parts 16 mg/mL homogenate with 2× Laemmli sample buffer (containing β-mercaptoethanol) to create 8 mg/mL heart homogenate. 2× Laemmli sample buffer is made by diluting 4× Laemmli sample buffer with water in a 1:1 ratio. Vortex the mixture, briefly spin down, and heat at 96 °C for 5 min. Cool to room temperature.

3.2. SDS-PAGE

1. Dilute the protein standard to 1/10 volume with 1× Laemmli sample buffer (see Note 3).

2. After removal of the comb and tape from the precut stain-free gel, prepare the electrophoresis tank by placing the gel inside the tank filled with SDS-PAGE running buffer. Load 1 μL (10 fold diluted) of the diluted protein standard in lane 1. Add 2 μL of 1× Laemmli sample buffer in lane

3. In the subsequent lanes, add increasing amounts of the heart homogenate (10–80 μg), and repeat this three times (Table 2) (see Notes 14).

4. Run the electrophoresis at 120 V constant voltage until the dye front reaches the agarose layer of the gel (see Note 15). Remove the gel from the cassette and cut away the wells and the agarose with a razor blade. Put a small cut at the top left side of the gel for orientation.

5. Lay the gel in the Bio-Rad ChemiDoc MP Imaging system. Image the gel using the stain-free gel setting. For the first image, be sure to check the setting to activate the gel for 1 min. This activation setting exposes the gel to UV light. Under UV light, the tryptophan resides in the proteins will react with the trihalo compounds in the gel and become fluorescent, enabling viewing of the total protein content in the gel (see Note 16). Manually adjust the exposure time if needed to avoid overexposed bands (Fig. 1A).

Table 2.

Gel loading pattern. Various volumes of the mouse heart homogenate sample (8 μg/μL) were loaded in order to result in 10–80 μg of the sample into the wells. Each amount was loaded into three different wells. The first set is from lanes 3–10, the second set is from lanes 11–18, and the third set is from lanes 19–26.

Lane	1	2	3	4	5	6	7	8	9	10
			11	12	13	14	15	16	17	18
			19	20	21	22	23	24	25	26
Sample amount (μg)	N/A*	N/A*	10	20	30	40	50	60	70	80
Volume (μL)	1	2	1.25	2.5	3.75	5	6.25	7.5	8.75	10

^*N/A: Not applicable. Lanes 1 and 2 was not loaded with the mouse heart homogenate. Lane 1 was loaded with 1 μL of the 1/10 diluted protein standard. Lane 2 was loaded with 2 μL 1× sample buffer.

Fig. 1.

Detection of total mouse heart protein on stain-free gel. (A) Detection of 10–80 μg of mouse heart homogenate using stain-free technology. The Criterion stain-free gel was exposed to 1 min of UV transillumination using the ChemiDoc MP to activate it before imaging. (B) Quantification of the relative intensity of total protein detected on the gel versus the total protein amount loaded onto the gel (μg) (n = 3 for each data point). Data are represented as mean ± standard deviation.

3.3. Protein Transfer

1. Soak one stack of filter paper in transfer buffer and place it in the Trans-Blot Turbo Transfer System cassette. Wet the nitrocellulose membrane with transfer buffer and lay on top of the filter paper stack. Allow at least 2 min for the filter paper and membrane to soak in the transfer buffer. Place the gel on top of the membrane, followed by another stack of filter paper soaked in transfer buffer. Firmly roll out the air bubbles with a roller. Use your fingers to apply mild pressure on the whole transfer stack while closing the lid of the cassette to ensure no additional air bubbles form. Wipe the outside of the cassette with tissue paper and place it inside the Trans-Blot Turbo Transfer System machine. Run with the Turbo midi setting of 7 min to transfer the proteins from the gel to the membrane.

2. Take the membrane from the cassette and image it while still wet in the ChemiDoc MP Imaging system using the stain-free membrane setting and optimized exposure time. There is no need to activate the membrane again. Adjust the exposure time as needed to avoid overexposed images (Fig. 2A). If necessary, the membrane can be cut into smaller sizes at this stage (see Notes 17 and 18).

3. Dry the membrane on top of a piece of tissue paper. Allowing the membrane to dry before proceeding to immunostaining has been suggested to help the proteins adhere better to the membrane (see Note 19).

Fig. 2.

Quantification of total mouse heart protein detected on nitrocellulose membrane using stain-free technology. (A) Detection of 10–80 μg of mouse heart homogenate on nitrocellulose membrane using stain-free technology. (B) Quantification of the relative intensity of total protein detected on the membrane versus the total protein amount loaded onto the gel (μg) (n = 3 for each data point). Data are represented as mean ± standard deviation.

3.4. Immunostaining

1. Place the membrane in a blotting container and add enough blocking solution (3% milk/TBST) to cover the entire membrane surface. Pour solutions into the corner of the blotting container and not directly onto the membrane so that the proteins on the membranes are not disturbed. Incubate the membrane in a shaker for 1 h. Pour out the blocking solution and wash three times with TBST for 3 min each in a shaker.

2. Add the primary mouse anti-20S proteasome β1 antibody solution and incubate at 4 °C overnight in a shaker, followed by incubation with shaking for 1 h at room temperature (see Note 20). Pour out the primary antibody solution and wash three times with TBST for 3 min each on a shaker.

3. Add the anti-mouse secondary antibody solution and incubate with shaking for 1 h. (see Note 21) Pour out the secondary antibody solution and wash three times with TBST for 3 min each on a shaker.

4. In a 1.5 mL centrifuge tube, mix together equal parts of Clarity western peroxide agent and Clarity western luminol/enhancer reagent. This creates the substrate for enhanced chemiluminescence (ECL). Pipette enough ECL substrate to evenly cover the surface of the membrane where the proteins are attached (see Note 22). Incubate the ECL covered membrane in a dark condition for 2 min before removing the excess ECL reagent (see Note 23).

5. Pick up the membrane from one side with a forceps and drain away the excess TBST by gently touching the edge of the membrane with a delicate wiper paper. Arrange the membrane on the imaging surface, taking care not to create bubbles on the surface.

6. Image the blot in the Bio-Rad ChemiDoc MP Imaging system with the Chemi Hi Sensitivity setting and optimized exposure time. For a higher resolution image, the blot can be imaged with the Chemi Hi Resolution setting with the optimized exposure time setting or with the exposure time set at approximately two times the time used for the Chemi Hi Sensitivity setting. The exposure time can be adjusted as needed to obtain the best image that contain no overexposed bands (Fig. 3A). Take a multichannel image with the stain-free and either the Chemi Hi Sensitivity or Hi Resolution setting to obtain an image of the chemiluminescent protein bands of interest overlaid with the stain free prestained protein markers. Although the other proteins on the membrane will also show up with the stain free image the chemiluminescent signal shows up on a different channel. Verify the molecular weight of the band of interest based on the multichannel image.

7. Quantify the total proteins from the gel and membrane imaged with the image analysis software (Fig. 1B and 2B). Quantify the immunostained protein band and normalize the intensity of the band against the stain-free total protein (Fig. 3B and 3C). The data should be normalized against the stain-free total protein from the membrane and not the gel (see Notes 24 and 25).

Fig. 3.

Quantification and normalization of 20S proteasome β1 levels in mouse heart homogenates. (A) Western blot of mouse heart homogenate (10–80 μg) using an antibody against the β1 subunit of the 20S proteasome. (B) Quantification of 20S proteasome β1 levels (10–80 μg), not normalized and normalized to total protein detected using stain-free technology on the nitrocellulose membrane (n = 3 for each data point). Data are represented as mean ± standard deviation. (C) Quantification of 20S proteasome β1 levels (10–40 μg), not normalized and normalized to total protein detected using stain-free technology on the nitrocellulose membrane (n = 3 for each data point). Data are represented as mean ± standard deviation.

4. Notes

1. Unused homogenization buffer can be stored at 4 °C for up to 3 months.

2. Sample buffer lacking β-mercaptoethanol can be made and stored at RT after aliquoting. After β-mercaptoethanol is added the sample buffer is ready to use and unused sample buffer with β-mercaptoethanol can be stored at −20 °C for at least 3 months or at −80 °C for at least 6 months.

3. Many pre-stained protein standards are available and can be used. However, some protein standards are better for stain-free gels as some standard proteins with high tryptophan content could result in very strong signals which affect the quantification of the lanes nearby. Standards can be diluted if the signals are too strong.

4. The SDS-PAGE running buffer should not be adjusted with acid or base. The pH of the final 1× buffer should be between 8.3–8.9. We prefer to use 20% SDS stock solution instead of powdered SDS to reduce the potential for SDS particles getting in suspension into the lab environment.

5. Nitrocellulose membranes have a slightly lower background as compared to polyvinylidene difluoride (PVDF) membranes, though PVDF membranes can also be used together with stain-free gels. In particular, low fluorescence PVDF membranes are recommended when the secondary antibodies are fluorescently labeled.

6. 10× TBS can be prepared and diluted to 1× TBS when needed. Unused 10× TBS can be kept up to 6 months at room temperature, as storing it at 4 °C will cause it to crystallize.

7. To add extremely viscous solutions like Tween-20 to other solutions, cut off the tip of the pipette with clean scissors to create a larger opening and pipette slowly.

8. Blocking and diluting antibodies can be done with either non-fat milk or BSA in TBST. For better results, dilute antibodies in 1 % BSA/TBST or 1 % milk/TBST, as these will usually give a better signal than 3–5 % BSA or milk. Depending on the antibody, milk or BSA may give a stronger signal with lower background noise. In general, BSA works better with biotin- and alkaline phosphatase-labeled antibodies, as well as anti-phosphoprotein antibodies. Unused BSA can be stored for up to 1 week at 4 °C.

9. A small aliquot of ECL substrate can be stored at room temperature, as cold ECL reagent results in lower signal intensities of the target bands.

10. In order to save on antibodies, different blotting box sizes can be used depending on the size of the membrane. For Criterion gels, the most common blotting box size is 15.5 cm (l) × 10.5 cm (w) × 3.5 cm (h).

11. Wiper papers do not leave traces of fiber or lint after being used to clean surfaces such as glass.

12. The Image Lab software comes with the ChemiDoc MP imager, as well as with several other Bio-Rad imagers, and can be installed on an unlimited number of lab computers.

13. Different ratios of tissue sample and homogenization buffer can be used depending on the concentration of protein sample required.

14. You should use the same type of tip and the same micropipette for all samples to minimize the difference in loading errors between the samples. Pre-wet the tip by sucking and dispensing the maximum volume of sample that will be loaded. In addition, a gel loading tip is recommended to prevent spillage of large sample volumes into other wells, as these tips are long with thinner openings that regular tips. Dispense the sample into the bottom of the well while slowly raising the tip towards the top of the well, ensuring that air bubbles do not get pipetted into the well. Keep the pipette plunger depressed until the tip is removed from the wells to prevent re-uptake of running buffer and sample into the pipette tip.

15. When running one Criterion SDS-PAGE at 120 V, milliamps (mA’s) of less than 35 typically indicates that the SDS-PAGE running buffer is too dilute, while mA’s of more than 80 sometimes suggests that the running buffer is too concentrated. Use new SDS-PAGE buffer. If the samples are not for quantification (such as for testing antibody specificity) you can adjust as needed with small amounts of 10× running buffer (if too dilute) or water (if too concentrated).

16. Longer activation times of 2 or 5 min do not increase the linearity of the total protein detected, although they may yield higher intensity bands as a result of more tryptophan residues being modified. However, activation times of longer than 1 min are not recommended if the primary antibody being used detects tryptophan residues, as the modification of the residues may interfere with antibody binding and subsequent detection of the protein of interest. There is no need to activate the gel again.

17. Image the membrane while still wet, as some image quality is decreased if the membrane is allowed to dry before imaging.

18. Using scissors or a razor blade, cut the membrane based on the molecular weight markers to obtain the region with the desired size of the protein of interest. If using diluted protein standards, it may be difficult to accurately cut the membrane to the desired size. In this case, stain the membrane with 0.1% Ponceau S stain for 30 seconds or until the proteins become clear enough. One corner of each piece of membrane can also be cut to indicate the orientation of the membrane. After cutting, remove the Ponceau S stain by washing multiple times with water or TBST.

19. After drying, the membrane can be stored at 4 °C for up to 1 week. Wet the membrane with TBST before re-imaging, if needed.

20. The primary antibody solution may be reused and stored at 4 °C for up to 1 week. Do not use if the BSA or milk has gone bad.

21. Try using a different secondary antibody if poor results are obtained during Western blotting after multiple trials. Good quality secondary antibodies are essential for high-quality Western blots.

22. The volume of ECL substrate needed varies depending on the size of the membrane. Uncut midi membranes will need around 5–6 mL of ECL substrate, though less ECL substrate is needed if a clear plastic support (e.g. saran wrap) is placed on top of the membrane, spreading the ECL evenly.

23. If the signal is weak, the ECL reagent can be left for longer than 2 min, but when the ECL reagent is left on the membrane for longer than 5 min, the length of time that the maximum signal lasts is shorter than when the ECL reagent is left for 2 min. Therefore, we recommend removing the excess ECL reagent and leaving the ECL reagent on for no more than 5 min (however, this will depend on your specific ECL reagent used).

24. Various image analysis software have different tools and methods to quantify gels and blots. Here, all quantification and normalization was done using Image Lab software, version 6.0.1, and the following tips are for that software. The Lane Finder tool can be used to automatically find all the lanes, though these should be further manually manipulated to better fit each lane. Do not change the width of just one lane, though individual lanes may be moved or bent. The Band Finder tool can be used to detect all the bands in each land with the custom sensitivity set to 100 %. In addition, the bands should be added, deleted, and/or adjusted to maintain consistency across lanes and samples by checking the Lane Profile tool, which shows the signal intensities of the bands above the variable background level.

25. All graphs were plotted with the SigmaPlot software, version 12. The results show that the total protein for the mouse heart homogenate was linear up to 80 μg of total protein (Fig. 2B), while the antibody against the 20S proteasome β1 was only linear up to 40 μg of total protein (Fig. 3B and 3C). Automatically fitting a linear line to the data can give the false impression that the antibody is linear up to 80 μg. However, we can determine when the line is no longer linear by looking for a significant reduction in the increase in signal intensity between each additional 10 μg of sample. The values for the normalized protein start dropping off after 40 μg, and that the not normalized values begin to curve the line after this point. It is best to take a look at the raw data. In order for the normalized data to be truly linear, the relative intensity of the values should stay close to 1. If the differences between each data point varies by more than 10%, we cannot consider it to be linear. We recommend stain-free western blotting for the quantification and normalization of proteins, with the caveat that the stain-free total protein may have a higher linear detection range than the antibody used, depending on the protein samples and antibodies utilized.