Week 1: Protein Isolation and Quantification for SDS-polyacrylamide Gel Electrophoresis (SDS-PAGE)

Introduction

The procedures for protein extraction vary widely depending on the types of organisms, tissues, and cells. The effectiveness of cell disruption/lysis determines the success of protein isolation. There are gentle approaches for cell lysis, such as freeze-thaw, enzymatic, osmotic, and detergent lysis. Other harsher methods are sonication, mechanical or glass-bead homogenization, and grinding with or without liquid N₂. Although detergent-based lysis, such as with SDS, is popular due to its ease of use, low cost, and efficient protocols, it may affect downstream manipulations.

Prior to SDS-PAGE, isolated protein must be quantified to determine the volume required for loading into each well. There are different ways of protein quantification. The most straightforward approach is to measure the absorbance at 280 nm because amino acids with aromatic side chains, i.e., tyrosine, tryptophan, and phenylalanine, absorb UV light at 280 nm (See Table 1-1 in Module 1). There is a relationship between protein concentration/quantity and their aromatic amino acid content; the protein quantity in the sample is generally proportional to its measured absorbance at 280 nm. However, this method is not ideal for protein mixtures as different proteins contain different proportions of aromatic amino acid content and thus vary absorbances. In addition, any non-protein content that absorbs at or near 280 nm interferes with the measured absorbance. Therefore, various colorimetric and fluorescent reagent-based protein quantification assays have been developed, where the reagent is added to the protein sample, producing a color change or increased fluorescence in proportion to the protein quantity in the sample. The quantity/concentration of the protein sample is determined by referencing a standard curve constructed with known quantities/concentrations of a purified reference protein.

Copper-, dye-, and fluorescence-based protein assays are three commonly used methods for protein quantification based on the chemistry and detection method involved.¹¹ Copper-protein assays, such as the Lowry protein assay,¹² is a two-step process of protein oxidation to reduce Cu²⁺ to Cu⁺ in an alkaline solution (the Biuret test). The Cu⁺ is detected by reducing the Folin-Ciocalteu (phosphomolybdic/phosphotungstic acid) reagent to produce an intense blue molecule, heteropoly-molybdenum blue, which is measured by reference to a standard curve. Bicinchoninic Acid (BCA) is also a copper-based assay. It replaces the Folin-Ciocalteu reagent to detect Cu⁺ after the Biuret test to produce a water-soluble, purple-colored product (absorbance at 562 nm).¹³ BCA is stable under alkaline conditions, and thus, it can be included in the copper solution to make the assay a one-step procedure. The BCA/copper complex exhibits a strong linear absorbance at 562 nm with increasing protein concentrations.

A Bradford assay is a dye-protein assay involving protein binding to an acidic dye, Coomassie® Brilliant Blue G-250.¹⁴ As shown below, this dye-binding assay is based on a differential color change of the protein-binding dye in response to different protein concentrations/quantities of samples. The absorbance maximum for an acidic dye solution shifts from 470 nm (red) to 595 nm (blue) when the dye binds to proteins made up of mainly basic and aromatic amino acids, especially arginine.

A standard curve can be made following a Bradford assay based on the OD₅₉₅ readings from known quantities or concentrations of a specific protein, such as bovine serum albumin (BSA). It provides a relative measurement of protein quantities/concentrations of samples. The unknown protein quantity of a sample can be determined by its OD₅₉₅ reading and the equation derived from the standard curve (Figure 5-2). Although the Bradford protein assay is less susceptible to interference from contaminants in samples, diluting a sample before starting the assay is often necessary due to the short linear detection ranges of two common standard proteins used, 200 to 900 µg/ml for BSA and 200 to 1500 µg/ml for immunoglobulin G (IgG), the most common class of immunoglobulin (a type of antibody) in blood and other body fluids (see introduction on antibodies in Week 3 of this module). In the process of making dilutions, errors in one dilution can be compounded in further dilutions, resulting in a linear relationship that may not always be accurate. Detergents, such as SDS, an essential reagent for protein isolation, may also affect the assay’s results.

Both copper- and dye-based protein assays are colorimetric detections. The other protein quantification method is a fluorescence-based assay with a higher sensitivity, meaning less protein needed for quantification and more downstream applications available, but different detecting equipment(s) may be required.¹¹ For example, when colorimetric assays cannot be used due to color interference in the sample, a fluorescence-based assay using a fluorometer or fluorescence microplate reader is required for detection instead of a standard spectrophotometer. Meanwhile, it can also be adapted for automation in high-throughput applications since the timing of this assay is not as critical as that of the colorimetric ones. It is critical to have a correct assay for protein quantification. Considerations for selecting a proper protein quantification assay include but are not limited to i) compatibility with common substances in samples (e.g., detergents, reducing agents, chaotropic agents, inhibitors, salts, and buffers), ii) standard curve linearity, iii) assay range and available sample quantity, iv) protein-to-protein variation, v) speed and convenience for the number of samples to be tested, and vi) availability of detection equipment, spectrophotometer or fluorometer.¹⁵ Chemicals used in protein isolation, such as the reducing agent, DTT, detergent, and Triton X-100, can interfere with some assays more than others. The Coomassie-dye–based assays (Bradford) method can be used to analyze samples containing reducing or copper-chelating agents because the dye is compatible with reducing agents and does not involve copper-protein binding reactions.

Protein assays can also be classified by their detection concentration range. Low protein concentration samples (< 20 µg/ml) may need a modified microplate protocol or a specific assay for dilute samples. If the protein concentration in the sample is high (> 2,000 µg/ml) with interfering substances, sample dilution may solve the problem. However, removing the interfering molecules may also be the only solution if a sample contains molecules incompatible with all assays. In general, the BCA and Coomassie (Bradford) protein assays complement with each other and can accommodate most samples.

For this lab exercise, hemolymph proteins are isolated from fruit fly larvae by grinding them in a cold Ephrussi-Beadle Ringer’s (EBR) buffer, which consists of a saline solution (0.129 M NaCl, 4 mM KCl, 2 mM CaCl₂, and 35 mM Tris.HCl pH 6.85). After centrifugation, the supernatant of the protein homogenate is collected and quantified by Bradford protein assay before being prepared for SDS-PAGE later. The protein isolation step is simple compared to other protein isolation protocols. Samples of BSA, bovine calf serum (BCS), and fetal bovine serum (FBS) are also quantified and prepared for SDS-PAGE.

Procedure

The protocol for this lab module follows the procedure described in a previous paper by Chang and Lovett.¹⁶

A. Surface sterilization of larvae

1. Obtain a culture of Drosophila larvae. The larvae should be ~5 days old (third instar). These are the largest soft ones often found crawling above the media on the glass wall (not the pupae).
2. For the following protocol, select 25 larvae by picking them up with fine forceps and then transfer them into a well of a spot plate filled with Ephrussi-Beadle Ringer’s (EBR) solution (0.129 M NaCl, 4 mM KCl, 2 mM CaCl₂, 35 mM Tris. HCl pH 6.85). Please handle them with special care!
3. Gently wash the larvae for about 1 min, then transfer them to a second well containing 70% ethanol (EtOH) and wash for about 1 min, followed by transferring to a third well filled with EBR.

B. Protein Isolation for SDS-PAGE

Due to endogenous proteases, everything must be kept on ice to prevent protein degradation.

1. Transfer the larvae to a pre-chilled 1.5 ml microcentrifuge tube.
2. Add 200 µl cold EBR to rinse the larvae to the bottom.
3. Homogenize gently by grinding thoroughly with a plastic pestle fitted to the 1.5 ml microcentrifuge tube. If needed, return the tube to ice and add another 100 µl cold EBR.
4. Centrifuge the tube for 1 min at top speed in the microcentrifuge.
5. Transfer the supernatant to a clean, pre-chilled 1.5 ml microcentrifuge tube and discard the pellet. Note: Keep the supernatant-containing tube on ice to prevent protein degradation.

C. Protein quantification with Bradford protein assay

1. Obtain 500 µl of 1 mg/ml BSA.
2. Place six 13 x 100 mm glass tubes in a rack and label them as 0, 20, 40, 60, 80, and 100 µg. Add an appropriate volume of 1 mg/ml bovine serum albumin (BSA) standard protein solution to each tube (Table 5-1).
3. Place three more 13 x 100 mm glass tubes in the rack and label them. Separately, add 10 µl of the isolated Drosophila protein, 10% BCS, and 10% FBS to each of the three glass tubes.
4. In each glass tube, bring the volume to 100 µl with RO H₂O and then add 3 ml of Bio-Rad protein assay dye.
5. Obtain a square piece of parafilm (5 cm x 5 cm) and use it to mix the content in each tube by firmly pressing one spot of parafilm against the rim of one of the tubes with your thumb, followed by inverting the tube 3–4 times. Repeat the steps with a clean spot on the same parafilm piece for the rest of each tube. (Ask the instructor if you do not know how to do this.)
6. Let the tubes stand for 15 min, measure the OD₅₉₅ of all tubes in the spectrophotometer, and record the data in Table 5-1.

   Table 5-1. OD₅₉₅ Readings from Bradford Protein Assay

   Tube	1mg/ml BSA (µl)	H2O (µl)	OD595	µg/ glass tube
   #1	0	100	0.00	0
   #2	20	80
   #3	40	60
   #4	60	40
   #5	80	20
   #6	100	0
   Drosophila	10	90
   10% BCS	10	90
   10% FBS	10	90

7. Prepare a standard curve using the OD₅₉₅ values (Y-axis) vs. the known amount of BSA (0, 20, 40, 60, 80, 100 µg) (X-axis) in Table 5-1.
8. Obtain the linear regression equation from the standard curve to determine the protein quantity in each glass tube (the last column of Table 5-1 and the 3^rd column of Table 5-2), and convert it to concentration by dividing it by 10 µl to obtain the original protein concentration of Drosophila, 10% BCS, and 10% FBS (the 4^th column of Table 5-2).
9. Calculate the volume (µl) of each sample needed for the protein quantities indicated in the last column of Table 5-2.

D. Preparation of protein samples for SDS-PAGE

All the steps below are based on your calculations, as shown in Table 5-2.

1. All groups: Make 20 µg in 30 µl of the Drosophila hemolymph protein solution in a 0.2 ml PCR tube. If needed, dilute the protein sample to 10% (See the last column of Table 5-2).
2. Odd-numbered groups:

1. 1. Sequentially, make 10% Bovine Serum Albumin (BSA) (10 µl of BSA plus 90 µl of water), 1% BSA (10 µl of 10% BSA plus 90 µl of water), and 0.1% BSA (10 µl of 1% BSA plus 90 µl of water) separately into three 1.5 ml microcentrifuge tubes. Mix well before each dilution!
   2. For SDS-PAGE, prepare three PCR tubes containing 6.6 µg BSA, 0.66 µg (10% BSA), and 0.066 µg (1% BSA), respectively. (For volume, see the last column in Table 5-2.)

2. Even-numbered groups:
   1. Sequentially, make 100 µl of 1% Bovine Calf Serum (BCS) (10 µl of 10% plus 90 µl of water), 0.1% BCS (10 µl of 1% BCS plus 90 µl of water), and 0.01% BCS (10 µl of 0.1% BCS plus 90 µl of water) separately into each of three 1.5 ml microcentrifuge tubes. Mix well before each dilution!
   2. For SDS-PAGE, prepare three 0.2 ml PCR tubes containing 6.6 µg BCS (1% BCS), 0.66 µg(0.1% BCS), and 0.066 µg (0.01% BCS), respectively.
   3. Repeat the step to make another three samples with 1.0%, 0.1%, and 0.01 % FBS.

3. All groups: bring the total volume in each 0.2 ml PCR tube to 30 µl with RO H₂O and then add 10 µl of 4X SDS-Laemmli sample buffer (187.5 mM Tris pH 6.8, 6% sodium dodecyl sulfate (SDS), 30% glycerol, 15% β-mercaptoethanol (β-ME), 0.015% bromophenol blue (BPB)) to each tube. Mix well by pipetting.
4. The instructor will collect all tubes, heat them at 95°C for 10 mins in a thermocycler, and store them at -20°C for the next lab.

Data Analysis and Discussion

1. Prepare a calibration curve using the OD₅₉₅ values (Y-axis) versus quantities/concentrations of the known samples (0, 20, 40, 60, 80, and 100 µg, or 0, 0.2, 0.4, 0.6, 0.8, and 1 µg/ml (mg/ml) of BSA) (X-axis) with the derived linear regression equation and R² based on the results in Table 5-1.
2. Calculate the concentration (µg/ml) of each SDS-PAGE sample in the 0.2 ml PCR tube you stored away. (Hint: You added the 4X SDS-Laemmli sample buffer as an additional volume to the samples!)
3. Watch the following videos: “How to load and run an SDS-PAGE gel” by Science@Waikato and “Western Blotting” by Bio-Rad Laboratories.