Two-dimensional difference gel electrophoresis (DIGE) experiments on tomato leaves and roots
We can analyze the differential expression of proteins in two different tissues and organs of the same plant by two-dimensional fluorescence difference gel electrophoresis (2D-DIGE).
Principle
This method involves the collection of leaves and roots from ripe tomatoes, preparation of protein extracts from the collected tissues, fluorescent labeling of the samples prior to differential gel electrophoresis, one- and two-dimensional electrophoretic separation, and image quantification of differential protein expression.
Operation method
Two-dimensional difference gel electrophoresis (DIGE) of tomato leaves and roots
Materials and Instruments
Leaf suspension buffer EDTA aqueous solution Thiourea buffer Ammonium sulfate solution SDS solution Move 3.1 Collection and precipitation of proteins from leaf and root tissues For more product details, please visit Aladdin Scientific website.
Cellular isoelectric focusing electrophoresis instrument Isoelectric focusing tray and cover
( 1 ) Cut green leaves from the middle of the plant and quickly cool them in a plastic bag. If a refrigerator is not available, place on ice before storing at -20°C.
( 2 ) For root tissue, cut a one-inch piece of root from the bottom of the stem, shake the root and quickly place it in water to wash off the soil three times in a row before quickly placing it in a plastic bag to freeze or place it on ice for temporary storage.
( 3 ) Weigh 2.5 g of flash-frozen root and leaf tissue, remove excess stems and other impurities, and cut these tissues into small pieces with precooled scissors, place in a precooled mortar, and grind in liquid nitrogen.
( 4 ) Place the crushed root and leaf tissues into a 40 ml centrifuge tube, suspend the powder with 25 ml of suspension buffer, shake vigorously to mix, store at -20°C for 45 min, shake again, and centrifuge at 35,000 g for 15 min.
( 5 ) Remove the supernatant, wash the precipitate with the same volume of aqueous EDTA (see Note 1) without stirring, and place on ice for 3-4 min. centrifuge at 35,000 g for 15 min, and then at least twice more until the precipitate is no longer green.
( 6 ) The precipitate is freeze-dried until it turns grayish brown. It contains proteins as well as other substances such as cell walls and fibers.
3.2 Preparation of protein samples from TCA/acetone powder
( 1 ) Weigh 15 mg of TCA/acetone powder (15. 3.1 prepared precipitate) into a 1.5 ml centrifuge tube.
( 2 ) Add 1 ml of 100 mmol/L Tris-HCl (pH 8.5) to each tube, spin for 1 min, centrifuge at 18000 g for 10 min, and remove the supernatant (see Note 2).
( 3 ) Add 1 ml of thiourea buffer to each centrifuge tube, extract the proteins from the powder, vortex for 5 min, sonicate for 10 min, and shake for 30 min in a shaker.
( 4 ) Centrifuge at 18000 g for 10 min and remove the supernatant. The supernatant contains protein extracted from leaf tissue at a concentration of 0.5~1 μg/μl.
3.3 Preparation of CyDyes dye and protein fluorescent labeling
( 1 ) Prepare CyDyes in fresh DMF (see Note 3), because DMF is easily oxidized and degraded, so fresh DMF should be used to prepare CyDyes dye.
(2) Prepare CyDyes solution by adding 1.5 μl of DMF to 1 μl of CyDyes dye. 1 ml of CyDyes solution is used to label 50 μg of protein (see Note 4). Since CyDyes is light sensitive, wrap the centrifuge tube in tin foil and store in the dark.
( 3 ) Pipette 100 μl of each sample (containing 50 μg of protein) into a 0.6 ml centrifuge tube (see Note 5), keeping the samples separate, with 1 μl of Cy3 Workup in one sample and 1 μl of Cy5 Workup in the other (see Note 6).
( 4 ) Vortex for 10 s, centrifuge briefly and repeat twice.
( 5 ) Place on ice for 30 min, avoiding light.
( 6 ) Terminate the reaction by adding 1 μl of 10 mmol/L lysine to each sample and leave on ice for 10 min.
( 7 ) Add 250 μl of IPG buffer for every 100 μl of fluorescently labeled sample to a final volume of 450 μl (see Note 7).
3.4 First direction protein separation: isoelectric focusing
( 1 ) Place the Bio-Rad focusing tray into the focusing chamber of the Bio-Rad. Pipette the sample into the center tray to avoid air bubbles.
( 2 ) Use tweezers to remove the plastic skin of the dry IPG strip and place it into the tray, making sure that the anode is attached to the anode of the tray and the cathode is facing the cathode. Carefully raise one end of the strip and lower the other end of the strip, making sure that the entire strip is soaked with buffer and that no air bubbles are generated.
( 3 ) Cover the strip with mineral oil to prevent the strip from drying out when energized.
( 4 ) Normally a minimum of 67000 Vh is required for full focusing.
a. Recommended Procedure: Hydration 50 V 12 h, 500 V 1 h, 1000 V 1 h, 8000 V 9 h, 100 V 5 h.
b. The first step is hydration, which should not be less than 12 h. Since Cy dyes are sensitive to light, the focusing tray should be covered to avoid light, or the entire instrument should be used in a dark room.
c. The last step is to remove the window, during which the adhesive strip can be removed. A low voltage ensures that the adhesive strip can be fully focused until it is removed from the tray.
3.5 Preparation of SDS-PAGE two-way gels
( 1 ) A low fluorescence glass plate reduces interference from the background.
a. For ease of handling during scanning, apply affinity silane to the side of the glass plate in contact with the gel. The other side of the glass plate in contact with the gel should be coated with stripping silane for good gel removal.
b. Wipe the non-gel-contact side of the low-fluorescence glass plate with Milli-Q ultrapure water and a paper towel, and wipe again with ethanol and a paper towel.
c. Add 0.5 ml of Affinity Silane Work Solution at a time to the non-glue-contacting side, dispersing it evenly with a paper towel until there is a total of 1 ml of Work Solution per plate.
( 2 ) Wipe the glass plate with ethanol, then with water, gradually add 0.5 ml of stripping silane until each glass plate is coated with 1 ml of stripping silane, dry for 5~10 min, remove the excess liquid.
( 3 ) Put the glass plates in a clean place to dry for at least 3 h. Pay attention to the fact that the glass plates treated with affinity silane and stripping silane should be placed separately. Blow away the dust on the glass plate with nitrogen before gluing.
( 4 ) Place paper dot markers on the side treated with affinity silane. Each mark should be placed at the midpoint of the short side approximately 1.5 cm from the edge of the plate. Depending on the numbering system, the markers should also be placed at the bottom of the plate about 1.5 cm from the edge. This makes it possible to distinguish the different gels in each subsequent processing step.
( 5 ) Preparation of 12.5% gels (900 ml total to fill 12 gels): 281 ml 40% acrylamide reservoir, 225 ml 1.5 mol/L Tris-HCl pH 8.8, 376 ml Milli-Q H2O, 9 ml 10% SDS, 9 ml 10% ammonium persulphate, 125 μl TEMED. acrylamide first, Tris-HCl, Milli-Q H2O, and SDS were thoroughly mixed in a large beaker, and then this mixture was filtered (0.2 μmol/L ) and placed in a reservoir, which was connected to the perfusion chamber by a peristaltic pump.
( 6 ) Turn on the stirrer, quickly add ammonium persulfate and TEMED (within 15 s), and turn on the peristaltic pump.
a. When almost all of the mixture has been pumped in, open the side of the mixer to allow as much of the mixture as possible to be pumped in.
b. Add replacement fluid to the hose chamber before air enters the tubing.
c. Continue to fill slowly until the 12.5% solution is just below the top of the non-touching glue plate.
( 7 ) Turn off the pump, add water-saturated butanol to ensure that the top and bottom of the gum are covered (butanol is on top of the Displacement Solution), let stand for 1 h, then pour off the butanol and cover with Milli-Q H2O.
( 8 ) The glue is best left at room temperature overnight when thoroughly solidified. Remove the glue-filling apparatus and wash off the acrylamide from the glass plate. Use the glue immediately or put it in the refrigerator for temporary storage (up to one week).
3.6 Two-way protein separation: SDS-PAGE
( 1 ) Remove the isoelectric focusing strip from the tray and gently wipe the front and back of the strip with a Kimwipe to remove excess mineral oil, being careful not to touch the Kimwipe to the strip, if needed dry the strip can be stored in a -80°C refrigerator.
( 2 ) Place the tape in a balancing tray with one end slightly elevated.
a. On a shaker, wash the strips for 30 min with 2 ml of fresh Reducing Re-equilibration Buffer.
b. Discard Reduction Re-equilibration Buffer and add 2 ml of fresh Alkylation Re-equilibration Buffer.
c. Shake on shaker for 30 min and discard Alkylation Equilibration Buffer.
d. Add 2 mL of SDS Electrophoresis Buffer and shake for 5 min.
( 3 ) Carefully place the strip on a pre-prepared 24 cm gel (see Section 15.3.5), place the plastic end of the strip against the end of the glass plate so that the strip is easy and convenient to place on the gel, there should not be any air bubbles between the strip and the gel. Seal the strip with 0.5% ( m/V ) agarose sealing solution, aspirate the sealing solution, place it on the top of the strip, and allow it to solidify at room temperature.
( 4 ) Place the gel's gel plate into the Ettan DALTsix electrophoresis chamber connected to a circulating cold water system.
a. Place the gels on the outside of the small tank, and when running several gels at the same time, place them evenly on either side, facing the same direction.
b. Place empty gel plates in the pool until each slot is full.
c. Fill with electrophoresis buffer until it reaches the bottom of the maximum scale.
d. Connect the lid to the bottom power supply and begin electrophoresis.
The power supply should be set to 2 W/gel for overnight electrophoresis. e. For daytime electrophoresis only, the voltage can be increased, but do not exceed 8 W/gel. f. If electrophoresis is to be performed during the day, the power supply should be set to 2 W/gel.
3.7 Fluorescence Image Scanning
( 1 ) Turn on the Typhoon Scanner and warm up the scanner for 30 min before use, the purpose of warming up is to scan more accurately.
( 2 ) The glass panel must be wiped clean with ethanol and paper towel before and after use. Do not use paper towel to wipe as it will easily scratch the glass surface.
( 3 ) Remove the glass plate from the gel bath, wipe the surface clean, and place the fluorescent labeling gel in the glass cassettes on the glass plate in the correct orientation.
( 4 ) Scan the image with appropriate filters and wavelengths, the wavelengths of each marker are as follows: Cy3 with a filter of 580BP, 532 nm green excitation light; Cy5 with a filter of 670BP, 633 nm red excitation light; Cy2 with a filter of 520BP, 488 nm blue excitation light. The initial parameters were depth+ 3 mm, 600 V photomultiplier setting and 500 μm pixels, and the high-resolution scanning was repeated for 50~100 μm pixels.
( 5 ) Images can be acquired with the Typhoon scanner software and saved in dataset format in the relevant folder containing the gel images, so that they can be opened with ImageQuant and then re-saved in dataset format for further processing in DeCyder. Alternatively, in Tiff format, the images can be further processed using other image analysis packages, such as Progenesis (Nonlinear Dynamics).
3.8 Image Analysis
Images obtained at different wavelengths are compared and analyzed by DeCyder (see Note 8). Due to the complexity of the procedure, which is not described in detail here, the basic steps are described below.
( 1 ) In order to ensure that the image sizes match exactly, a bar graph showing the number of unchanged, increased, and decreased protein spots in the two images was measured and normalized using Process Image.
( 2 ) Use an exclusion filter, or manually remove any obvious non-protein background spots or streaks. Adjust the critical parameters of the histogram to display protein spots as needed. These protein spots are the specified amount of up- or down-regulated expression, e.g., greater than 2-fold difference.
( 3 ) Manually examine the histogram and images to determine which spots have significant expression changes and are of sufficient amount for mass spectrometry identification. In most cases, many proteins, although significantly differentially expressed, are so small that they cannot be reliably quantified and feasibly identified. The most interesting protein spots to study are usually of medium to high abundance and are very clearly differentially expressed.
( 4 ) In order to quantify differential expression more accurately, especially in cases of very weak differential expression, the procedure needs to be repeated 3 times for the results to be statistically significant (see Note 8).
3.9 Re-staining for further processing
The next step is to perform image analysis. The spacer strips in the glass box are removed and the gel is silver-stained [ 29, 30]. Silver staining is performed on a stationary glass plate, which allows for easy visualization and manual scooping of protein spots for the next step of mass spectrometry [6]. Silver staining ensures that the majority of proteins are labeled as opposed to CyDye labeling, which only labels a small number of proteins [8]. With silver staining, most of the proteins labeled with CyDye are visible, although there is some variation between samples. Compare the silver-stained image with the CyDye image, taking care to make sure you dig into the right spots. Some small molecular masses of CyDye often cause some drift, and some proteins can cause some variation with different methods (see Note 9).
3.10 Analysis of results: differential determination of protein expression in tomato roots in response to salt stress
1. Salt stress treatment
( 1 ) In a temperature-controlled greenhouse, tomatoes were allowed to grow to a height of 4 inches (1 inch = 25.4 mm).
( 2 ) At the time of trial planting, 20 seedlings were grown in a Hoglan fertilizer mix containing MiracleGro fertilizer.
( 3 ) All 20 plants were watered normally every day. 20 days later, 10 of the plants were labeled and treated with 25 mmol/L NaCl for one week. The NaCl concentration was gradually increased to 100 mmol/L over a period of one month.
( 4 ) The 10 control plants were watered throughout the period.
2. Collection of leaf and root tissues
( 1 ) During 7 weeks, after two weeks of continuous watering with 100 mmol/L NaCl, 3 to 5 g of leaves of control and treated healthy plants were cut twice a week until the leaves were necrotic.
( 2 ) - Once the leaves were necrotic, the plants were uprooted, the roots were rinsed with water, divided into proximal and distal parts from the middle of the root, and immediately snap-frozen in liquid nitrogen.
3. Extraction of proteins from the near stem end
( 1 ) Crush the collected proximal roots with a hammer.
( 2 ) Prepare the protein precipitate according to the TCA acetone method mentioned in section 15.3.1.
( 3 ) Since the gel showed that the roots contained less protein than the leaves, it was necessary to extract the proteins three times as described in section 15.3.2. The proteins from these three extractions were placed in a centrifuge tube and labeled with CyDye.
( 4 ) Pool the samples and run the gel for analysis using the methods in sections 15.3.3 and 15.3.6. The results are shown in Figure 15-1 to Figure 15-4. 
4. Comparative image analysis of DIGE gels of control and salt-stressed near-root tissue sections
Data from 4.3 were analyzed using the DIA model, which is described in Section 15.3.8. The starting data output from the DeCyder program is represented by virtual colors as shown in Figure 15-1. Cy 5-tagged protein spots of control plants are in red, Cy3-tagged protein spots of salt-stressed plants are in green, and protein spots that are present in both plants and are expressed in similar amounts are in yellow. This makes it easy to recognize proteins for which there is a clear difference in expression. In this experimental example, the vast majority of protein spots are yellow, indicating that the difference in protein expression is not very pronounced in both samples.
Different images of the two fluorescence wavelength scans are shown in Figure 15-2, and these images can be used to further analyze the protein expression differences. These two images look similar, but there are more protein spots on Figure 15-2B. There is a clear difference in overall protein loading or labeling efficiency, which is a significant advantage of this method. The software can be used to normalize the expression of these two images to all protein spots included, and only those protein spots with higher or lower than average expression are labeled as differential protein spots.
The quantitative analysis of the two images produces the histogram shown in Figure 15-3. As shown in the Protein Spot Detection window, after normalization and processing, the software detected 421 protein spots, of which 384 were unchanged, 34 had elevated expression for Cy3 markers, and 3 had elevated expression for Cy5 markers. x-axis is the ratio of the log value of the volume of the protein spots in the two images, the y-axis on the left is the frequency of the spots, and the y-axis on the right is the ratio of the log value representing each of the protein spots. The x-axis is the ratio of the logarithmic value of the volume of the protein spots in these two images. The blue curve represents the histogram of the protein spot frequency pattern, which was generated based on the actual histogram of the red curve.
The two perpendicular lines represent the x-axis points with 1.5-fold difference in protein expression and the protein points with no change in expression. The coloring strategy is similar to that of Figure 15-1, with yellow representing the protein points with no change, green representing the protein points with increased expression of the Cy3 marker, and red representing the protein points with increased expression of the Cy5 marker.
In this type of analysis, many protein sites are detected with a change in expression, but the change is so weak that it is not easy to identify. In the analysis of Figure 15-3, we applied an absolute fluorescence threshold of 106 fluorescence units (shown as a dotted line) to exclude these protein spots. There are four protein spots, labeled a to d, that are outside the normal distribution curve on the x-axis and beyond 106 units of fluorescence on the y-axis, so these are Cy3-tagged protein spots with more than 1.5-fold expression, which can be used for mass spectrometry identification. There are also many proteins in the histograms that are significantly differentially expressed, but below the thresholds we have specified.
The 4 highlighted proteins represent those proteins that were up-regulated more than 1.5-fold in response to salt stress. The first 3 protein spots, labeled a, b, and c, highlight the shortcomings of this approach, as they are part of a low-resolution protein that has been manually divided into protein spots using the analysis software. The protein labeled d, however, is a high-resolution protein with a fluorescence volume ratio of 1.79, as shown in Figure 15-4, and can be cut down for mass spectrometry.