Ettan DIGE System - Frequently Asked Questions for Minimal Dyes and Saturation Dyes
FAQs for Minimal Dyes
Experimental design
Sample preparation and labelling
Isoelectric focusing
Gel preparation and running
Image acquisition
Dynamic range and sensitivity
Image analysis and statistics
DeCyder™ Differential Analysis Software - general questions
Spot picking and downstream analysis
System validation
Experimental design
Q: What approaches to experimental design do you recommend?
A: To obtain valid statistical analysis there must be a common pooled internal standard sample in all gels. It is recommended that this internal standard is generated by pooling equal amounts of each of your experimental samples. The internal standard is labelled with one CyDye™ DIGE Fluor dye (typically Cy™2 if using the CyDye DIGE Fluor minimal dyes) and experimental samples labelled with one or both of the other dyes (Cy3 and/or Cy5). The internal standard is run on every gel, along with the experimental samples. It is good laboratory practise to distribute individual experimental samples evenly between CyDye DIGE Fluor dyes as illustrated below:
The system variation of this approach is very low, much lower than the inherent biological variation between individuals. Therefore, we recommend that biological replicates be tested in preference to gel replicates to maximize the value of the information that can be obtained from each experiment.
Use of an internal standard ensures that every spot on every gel is represented in an image common to all gels. Each protein spot in a sample can therefore be compared to its representative within the internal standard to generate a ratio of relative expression. Matching of these internal standards creates an intrinsic link between the protein spots from the samples on each of the different gels. Quantitative comparisons of proteins between samples are made based on the relative change of each protein spot to its own in-gel internal standard. This removes gel-to-gel (system) variation, a common problem associated with conventional “one sample per gel” 2-D electrophoresis studies, enabling accurate, statistical quantitation of induced biological change between samples. Ettan DIGE system is the only protein difference analysis technique that utilises this approach.
Linking every sample in-gel to a common standard offers many advantages:
· accurate quantitation and spot statistics for changes in protein abundance;
· increased confidence in matching between gels;
· flexibility of statistical analysis depending on the relationship between samples;
· separation of induced biological variation from system variation.
It is strongly advised that biological replicates are included in every group. This will enable accurate measurement of the induced biological change (differences in protein abundance due to a treatment/disease/life cycle stage) above a baseline of inherent biological variation (intrinsic differences that occur within populations e.g. differences from animal-to-animal, plant-to-plant or culture-to-culture which have been subjected to identical conditions). The more biological replicates, the more that inherent biological variation is accounted for and therefore, the more meaningful the results. Without biological replicates, results are not biologically relevant. In the example shown above, each treatment group contains 3 individuals A-C.
For more information on 2-D DIGE experimental design refer to the Ettan DIGE System User Manual (code no. 18-1173-17).
Q: Can we choose our own experimental design and still use DeCyder™ Differential Analysis Software?
A: You can choose your own experimental design and use DeCyder Differential Analysis Software. However, if you want to match across gels and achieve valid statistical results, the common internal standard must be on each gel. The most effective standard is a pool of all the experimental samples within the experiment. In this way, every protein in the experiment is represented within each gel.
Q: Why does using a pooled internal standard increase the precision of the technology?
A: With any biological technique, the precision is greatly improved if an internal standard is used. This is one of the key benefits of 2-D DIGE as this is the only 2-D technique that allows the user to run an internal standard. When using a pooled internal standard, every protein in the population appears on each gel. Each sample can then be compared internally to the same standard, to account for any gel-to-gel variation. This facilitates fully automated and accurate spot statistics when using DeCyder™ Differential Analysis Software. The inclusion of an internal standard on each gel also aids matching between gels.
Occasionally, protein expression changes between samples are so large that the expression of specific proteins can be switched off. The use of the pooled internal standard with spot co-detection of multiplexed samples avoids any problems caused by presence-absence events. The internal standard would always contain every protein within the experiment enabling the absence of specific proteins in individual samples to be analyzed accurately with confidence.
For more information please refer to the following paper, Alban et al. A novel experimental design for comparative two-dimensional gel analysis: two-dimensional difference gel electrophoresis incorporating a pooled internal standard. Proteomics,3, 36—44 (2003).Sample preparation and labelling
CyDye™ DIGE Fluor minimal dye characteristics
Q: How can I calculate the molecular weight (Mr) added onto a protein after labelling with each CyDye™ DIGE Fluor minimal dye?
A: The molecular weight of each CyDye DIGE Fluor is provided on the specification sheet supplied with each dye (TFA salt). The molecular weight added after labelling can be calculated from this by using the formulae below.
Note: The calculation for proteins labelled using CyDye DIGE Fluor minimal dyes is different from the calculation for proteins labelled with CyDye DIGE Fluor saturation dyes.
Example of a calculation for minimal dyes (CyDye DIGE Fluor Cy™2 minimal dye);
Quoted mass of dye (TFA salt) = 663.62
Mass of dye cation (-TFA) = 663.62 – 113.02 = 550.60
Mass added to protein (-NHS, -H from protein amine group) = 550.60 – 114.08 - 1
= 435.52
The masses added for each of the other two minimal dyes can be calculated in a similar manner;
CyDye DIGE Fluor Cy3 minimal dye, mass added to protein = 467.69
CyDye DIGE Fluor Cy5 minimal dye, mass added to protein = 465.67
Q: Why are proteins labelled with minimal dyes (stable for 3 months at -70 C) more stable than those labelled with saturation dyes (stable for 1 month at -70 C)?
A: The shorter storage time recommended for CyDye™ DIGE Fluor saturation dyes is a precautionary measure due to differences in labelled protein concentration. CyDye DIGE Fluor saturation dyes are designed for use with scarce sample types and therefore, the concentration of the protein sample is likely to be relatively low (compared to samples suitable for use with CyDye DIGE Fluor minimal dyes). Proteins are less stable when stored at low concentrations, hence a shorter storage time is recommended when labelling with CyDye DIGE Fluor saturation dyes. The shorter storage time is not due to differences in stability between the two types of CyDye DIGE Fluor dye.Reagent compatibility
Q: Does Amersham Biosciences provide a list of reagents compatible with minimal labelling?
A: Yes, a complete list of recommended reagents and consumables for use with Ettan™ DIGE system is available in the Ettan DIGE System User Manual (Appendix E2). However when testing new components as part of the sample preparation procedure always, handle samples on a case-to-case basis.
Q: Are there any components we should avoid in the labelling reaction?
A: For minimal labelling, omit all primary amines such as ampholytes that compete for available CyDye™ DIGE Fluor minimal dye. Reducing agents such as DTT and TCEP (Tris-[2-carboxyethyl]phosphine) should be avoided until after the labelling reaction. DTT is a thiol that competes for available dye at high concentrations. We have demonstrated that a concentration of 2 mg/ml DTT causes a slight reduction in labelling. At 5 mg/ml, DTT causes a 2-fold reduction in labelling efficiency. TCEP at concentrations between 0.5 mM and 1 mM cause a slight reduction in labelling. At 2 mM, the labelling efficiency is significantly reduced. Beta-mercaptoethanol should be omitted. The amount of CyDye DIGE Fluor minimal dye can be increased to counteract the presence of DTT or TCEP, although users should be aware that this may lead to non-minimal labelling of proteins.
For further information about compatible reagents please refer to the Ettan DIGE System User Manual.
Q: I thought that Tris is a primary amine, yet the protocols suggest using Tris to buffer the labelling reaction. Wouldn't the Tris compete for the available fluor?
A: You may use Tris base with 2-D DIGE labelling because although it is a primary amine, it is much less reactive than the epsilon primary amine of lysine. This is due to its structure and the arrangement of its functional groups (Fig 1).
Fig 1. Structures of (a) Tris base and (b) lysine.
The amine group of Tris is attached to a quaternary carbon atom. This carbon has three bulky groups attached to it that sterically interfere with nucleophilic attack by the amine group on other molecules (e.g. CyDye™ DIGE Fluor). In addition, the three hydroxyl groups are electron-withdrawing and so reduce electron density on the nitrogen lone pair, thereby reducing its reactivity. The epsilon primary amine group of lysine is sterically unhindered and is remote from any electron-withdrawing groups, making it a much more reactive nucleophile in comparison.
The poor nucleophilicity of the Tris amine group, enables the use of Tris as a buffer without compromising the efficiency of CyDye DIGE Fluor minimal dye labelling.
Q: What effect do SDS, NP-40, Triton X-100, and lipids have on the sample preparation procedure?
A: The standard lysis buffer recommended for use with 2-D DIGE labelling contains 4% CHAPS. However, proteins for certain sample types may be more efficiently solubilized by the use of alternative detergents. Some of these detergents have been shown to have a detrimental effect on CyDye™ DIGE Fluor minimal dye labelling efficiency. Labelling efficiencies can be reduced by the addition of 1% SDS, but 0.25% appears to have no effect. NP40 is compatible to 1%. However Triton X-100 at a concentration of 1% has been shown to reduce labelling by 17%. Lipids may be a problem but these affect protein solubility rather than labelling efficiency.
Always handle samples on a case-to-case basis. Test any non-standard lysis buffer components for their effect on labelling efficiency using 1-D gel analysis prior to 2-D DIGE experiments (see the Ettan DIGE System User Manual for detailed protocols).
Q: What is the effect of nucleic acids in the sample?
A: High nucleic acid content can cause gel smearing in the second dimension due to the clogging of pores in the gel. This can be reduced by sufficient sonication to shear the DNA. DNA can also be removed by treating the sample with DNAseI, or using Ettan™ 2-D Clean-Up Kit (Code no. 80-6484-51). Using DNase or RNase as part of the sample preparation may result in the enzymes appearing on the gel depending on the molecular weight and type of gel.
Significant amounts of DNA in the sample can aggregate and cause the sample to become viscous and awkward to pipette. We recommend that magnesium acetate be added to the lysis buffer to prevent this aggregation.
Q: Can you use DNase and RNase in the samples?
A: Yes, but if you add these prior to labelling, they may show up on the gel depending on the molecular weight and type of gel.
Q: Can you add protease inhibitors to the samples?
A: The following protease inhibitors have all been tested for compatibility with Ettan™ DIGE system: |
| Protease Inhibitor | Supplier | Product Code |
| Pefabloc SC (AEBSF) | Roche | 1585916 |
| Complete mini protease inhibitor cocktail | Roche | 1836153 |
| Complete mini protease inhibitor cocktail, EDTA-free | Roche | 1836170 |
It is important that different samples being compared in the same experiment are all prepared in the same way so if phosphatase inhibitors are to be used, they should be included in all experimental samples. For example, the use of phosphatase inhibitors can result in visualisation of new spots that could be mistaken for differences in protein expression.
When including new buffer components (e.g. phosphatase inhibitors), always check the protein lysate for successful labelling (instructions are provided in the Ettan DIGE System User Manual, code no. 18-1173-17)
Any of these protease inhibitors can be used to protect protein extracts prepared for labelling with CyDye™ DIGE Fluor minimal dyes. It is important to use them in conjunction with a protector reagent (Pefabloc® SCPLUS, AEBSF, Roche, Code no. 1873601) to prevent the formation of charge trains.
It is important that different samples being compared in the same experiment are all prepared in the same way so if protease inhibitors are to be used, they should be included in all experimental samples. For example, the use of protease inhibitors can result in visualization of new spots that could be mistaken for differences in protein expression.
We recommend that any new components added to the lysis buffer (e.g. protease inhibitors and blocking agents) should be tested for their effect on labelling efficiency using 1-D gel analysis prior to 2-D DIGE experiments (see Ettan DIGE System User Manual for detailed protocols).Labelling efficiency
Q: Is there a risk that labelling efficiency and therefore quantitation is performed unequally if the concentration of substances such as SDS differs in two samples?
A: Yes, compounds that have an adverse effect on labelling in one sample could affect the final quantitation. In addition, the cumulative effect of two or more compounds that have an adverse effect on labelling in one sample could also affect the final quantitation. In these instances the samples should be cleaned up and resuspended in the same lysis buffer. We recommend the Ettan™ 2-D Clean-Up Kit (Code no. 80-6484-51).
Q: Will the quality of my DMF affect labelling?
A: Yes, DMF must be high quality anhydrous (specification: 0.005% H2O, 99.8% pure) and every effort should be taken to ensure it is not contaminated with water. DMF, once opened, will start to degrade, generating primary amine compounds. Primary amines can react with the NHS ester group of CyDye™ DIGE Fluor minimal dyes, reducing the concentration of dye available for protein labelling. It is therefore recommended that only DMF that has been open for less that three months (and is within the expiry date) is used for all 2-D DIGE applications using the Ettan™ DIGE system.
Q: Can you explain the labelling ratio?
A: CyDye™ DIGE Fluor minimal dyes will label 1–2% of all lysine residues in the lysate. The minimal dyes are the limiting factor in the reaction so dual or multiple labelling of the same protein should not occur when the ratio is maintained at 50 µg protein:400 pmol CyDye DIGE Fluor minimal dye.
Q: One protein has more lysine residues. Won’t it label more than the other proteins?
A: This might happen, but this does not affect your quantitation. The same protein will label to the same percentage with the different CyDye™ DIGE Fluor minimal dyes so your within-spot analysis is not affected. We have never seen any direct evidence of this happening.
Q: What is the distribution of labelled:unlabelled protein after the labelling reaction?
A:If the labelling ratio is maintained within the recommended limits, such as 400 pmol of dye to 50 g protein the majority of protein will remain unlabelled. Labelled protein will contain only one dye molecule.
Q: Is it possible that differences in post-translational modifications between samples will cause differences in labelling efficiency?
A:If post-translational modifications affect the labelling, these will affect the labelling with each of the different CyDye™ DIGE Fluor minimal dyes to the same degree. However, we have never seen this phenomenon before.
Q: What happens if a single protein in the mixture makes up a significant percentage of the total protein content, e.g., albumin in serum?
A: As with traditional 2-D methods, proteins of low abundance may be harder to visualize, and the highly abundant protein will probably obscure spots of lower abundance. The single protein will effectively take up most of the available CyDye™ DIGE Fluor minimal dye during labelling. To reduce this titration effect, pre-fractionation of the sample can be carried out to remove the interfering protein. Protein concentration
Q: What protein concentration range is recommended prior to the start of labelling with CyDye™ DIGE Fluor minimal dyes?
A: Protein samples at higher concentrations (5–10 mg/ml) label more efficiently than dilute solutions (< 1 mg/ml). Protein concentration should not exceed 10 mg/ml as protein aggregation and precipitation may occur causing loss of protein and horizontal or vertical streaking (Gorg et al. The current state of two-dimensional electrophoresis with immobilised pH gradients. Electrophoresis 21, 1037–1053 [2000]). We recommend a concentration of 5 mg/ml for optimal labelling.
Q: Is it possible to make a quantitative comparison of two samples in which the total protein concentration is different?
A: Ideally, experimental samples should be diluted to the same concentration prior to labelling. However, if this is not possible (e.g. the concentration of one of the samples is very low) a quantitative comparison can still be made providing all samples are labelled using the recommended ratio of 50 µg protein per 400 pmol dye. The same amounts of sample labelled with each CyDye™ DIGE Fluor can then be combined for loading on the IPG strip for quantitative comparison.
Q: What method do you recommend for protein concentration determination?
A: If you have detergent in your sample such as CHAPS, use a detergent compatible method. We recommend the use of the Ettan™ 2-D Quant Kit (Code no. 80-6483-56).
Note that when using protein determination reagent with thiourea present in the lysis buffer, the aliquot used for protein concentration determination must be acidified. CyDye™ DIGE Fluor minimal dyes and post stains
Q: How long will my CyDye™ DIGE Fluor minimal dyes keep once I have reconstituted them?
A: Once reconstituted to a concentrated stock of 1 mM, the dyes will keep for two months if stored in the dark at -20 oC. The more dilute working stock at a concentration of 400 µM will keep for two weeks if stored in the dark at -20 oC.
Q: Are there differences in labelling between the three CyDye™ DIGE Fluor minimal dyes?
A: In our experience we have not seen any significant differences between the labelling for each of the CyDye DIGE Fluor minimal dyes, although samples without 2 M thiourea in the lysis buffer tend to show more variability than samples containing thiourea. Differences in spot patterns between the three dyes are more likely to arise due to differences in sample preparation prior to labelling. As a precaution, we recommend checking each new sample type, by labelling the same sample in separate reactions with each of the three CyDye DIGE Fluor minimal dyes, recombining, and running on a single gel. Spot patterns can then be compared to ensure that labelling is identical for each dye.
Q: Why can’t I pick directly from a gel containing CyDye™ DIGE Fluor labelled proteins?
A: When a CyDye DIGE Fluor is coupled to a protein it will add approximately Mr 500 to the proteins mass. This causes a shift between labelled and unlabelled proteins and is more marked for lower molecular weight proteins. Because of this shift, the most fluorescent point on a spot represents the highest concentration of the labelled protein, but does not correspond to the highest concentration of unlabelled protein. Thus, to maximize the amount of protein extracted from each spot, preparative gels are run using higher protein loadings and post-staining e.g., with SYPRO™ Ruby. These gels are matched to the quantitative data obtained from the analytical gels so that spots of interest can be accurately picked from the preparative gel(s).
Q: How does the staining pattern differ between CyDye™ DIGE Fluor minimal dyes, fluorescent post-labelling and silver-stained gels?
A: There is no difference between the 2-D spot pattern you see with CyDye DIGE Fluor minimal dye labelling, fluorescent post-labelling, and post-staining with silver. The mass increase resulting from attachment of a CyDye DIGE Fluor to a protein will shift the 2-D spot pattern to the same extent for each protein in the sample so all the labelled proteins will remain in the same place relative to one another. Isoelectric focusing
Q: What is the best method for applying protein samples to strips for isoelectric focussing?
A: Cup loading and in-gel rehydration are the most commonly used methods of sample application and can be used with Cup Loading Strip Holders (Code no. 80-6459-43). More CyDye™ DIGE Fluor minimal dye-labelled protein enters the strip using the cup-loading method. The use of longer rehydration times (e.g. 22 h) when using in-gel-rehydration can increase the number of spots detected (particularly in the high molecular weight range), although the overall amount of protein in the strip remains unchanged. The cup-loading method is favored but in-gel rehydration is useful for preparative gels where the sample volume may be too large to apply using the cup-loading approach.
Other strip holders available (e.g. Regular 24 cm Strip Holders, Code no. 80-6469-88) allow sample loading via gaps/wells at the sides of the strip holder. CyDye DIGE Fluor minimal dye labelled protein loaded using this method results in levels of protein entry intermediate between cup loading and in-gel rehydration.
Q: Does labelling the proteins with CyDye™ DIGE Fluor minimal dyes affect the isoelectric focusing parameters?
A: We recommend that a high salt compatible isoelectric focusing program be used to focus proteins labelled with CyDye DIGE Fluor minimal dyes (refer to the Ettan DIGE System User Manual for examples of focusing protocols). CyDye DIGE Fluor minimal dyes are designed to ensure that the net charge of the protein remains unchanged upon labelling so the isoelectric point is unaffected. Focusing times should likewise remain unchanged.
Q: Is the IEF of protein samples labelled with CyDye™ DIGE Fluor minimal dyes more difficult or technically demanding?
A: Because the pI is unaffected, IEF separation is no more technically demanding than for unlabelled proteins. CyDye DIGE Fluor minimal dyes have the advantage of being highly sensitive, with a wider dynamic range than other techniques such as CoomassieTM Blue and silver staining. However, with all fluorescence imaging techniques, consideration is required to avoid the inclusion of components that either fluoresce or have the potential to quench fluorescence.
Q: What is an ampholyte and what is its function?
A: Ampholytes are small soluble amphoteric molecules (carrying both positive and negative charges) that have a high buffering capacity near their isoelectric point. Their function is to generate a stable pH gradient allowing proteins to be separated according to their isoelectric points.
Q: Do you recommend anodic or cathodic cup loading?
A: It is recommended that anodic cup loading is used because the majority of proteins are more soluble under acidic conditions. However some sample types may benefit from cathodic loading. Cathodic loading can also give better results when using acidic narrow range strips. Whether anodic or cathodic loading is used depends upon the sample types and preparation. Separation conditions for specific sample types can be found in the Ettan DIGE System User Manual and on this website link.
Q: Can I use narrow-range IPG strips?
A: A wide selection of narrow-range Immobiline™ DryStrip gels (both 18 cm and 24 cm) have been tested with Ettan™ DIGE system and show comparable results to silver staining. For best results when using narrow-range Immobiline™ DryStrip gels, we recommend using Pharmalyte™ broad range pH 3–10 solution (Code no. 17-0456-01) in preference to narrow-range IPG buffers. For examples of gels prepared using narrow-range Immobiline DryStrip for the Ettan DIGE system, please refer to the range of E.coli samples given in the sample types tables on this web site link.
Spot resolution and reproducibility can be improved for basic narrow-range Immobiline DryStrip gels by using DeStreak™ Reagent (Code no. 17-6003-18) during the isoelectric focusing step. When using acidic narrow-range Immobiline™ DryStrip gels, results may be improved by implementing cathodic cup loading.
Q: Can I use Destreak Reagent to improve the resolution of basic proteins when using 2-D DIGE?
A: Yes. Destreak Rehydration Solution (code no. 17-6003-19) is compatible with CyDye™ DIGE Fluor minimal dyes (as well as CyDye DIGE Fluor saturation dyes). We recommend rehydrating all strips that contain basic pH ranges using Destreak Rehydration Solution.Gel preparation and running
Q: How does Amersham Biosciences PlusOne™ ReadySol IEF 40% acrylamide solution compare to Bio-Rad premixed acrylamide/bis-acrylamide powder for preparing 2-D gels?
A: PlusOne acrylamide solution gives comparable separation as the Bio-Rad premixed powder for the same percentage gel (both isocratic and gradient gels tested). In addition, PlusOne Ready Sol IEF 40% is in solution form so it is less hazardous and far more convenient to use.
Q: Can piperazine di-acrylamide (PDA) be used as an alternative cross-linker to acrylamide when making the gels?
A: Yes, PDA can offer increased gel strength and higher spot resolution and is compatible with Ettan™ DIGE system when substituted weight-for-weight with acrylamide (suitable for both isocratic and gradient gels).
Q: Is it essential to use low-fluorescence glass plates?
A: Yes, see the next FAQ.
Q: What happens if ordinary glass plates are used?
A: The background fluorescence will increase. The signal to background ratio will be reduced. This could result in the detection of fewer spots, especially for those present at a lower concentration.
Q: Is the thickness of the glass plates important?
A: Yes, the thickness of glass will affect the focal plane of the gel. As the center of the gel moves away from the focal plane of the imager, the intensity of the spots will decrease, therefore fewer spots will be seen. We recommend that the 3.4 mm-thick, low-fluorescence glass supplied for use with Amersham Biosciences equipment is used for all gels in the Ettan™ DIGE system.
Q: What is the best way to clean plates after use?
A: Wash with hot tap water (this will help dissolve the acrylamide) and detergent (we recommend a non-fluorescent detergent such as Hemo-sol glassware detergent, supplied by MERZ and DADA AG. Switzerland, code no. 56022 3E). Use a soft sponge to ensure all adhered polyacrylamide fragments are removed. Be careful not to use washing implements that may scratch or damage the glass. Rinse thoroughly with Milli-Q water and leave to air dry. Alternatively, plates can be dried using lint-free wipes (Kimberly-Clark, code no. 33370).
Q: Is the cleaning protocol different for plates that have been treated with PlusOne™ Bind-Silane?
A: Yes, scrape the gel off with a plastic wedge, soak the plates in 5% Decon (called Contrad in the US) overnight to remove any gel fragments that adhere to the plate. It is important that fresh Decon is used. If opaque deposits remain on the plate, the plate can be soaked for an hour in 1% HCl then rinsed in Milli-Q water. Leave to air dry or dry using lint-free wipes (Kimberly-Clark, code no. 33370). Note that etching of the plates may occur if they are left in Decon or HCl for longer than the recommended period.
Q: How long can I store the labelled gel before I have to scan it and what storage conditions should I use?
A: CyDye™ DIGE Fluor minimal dyes are very stable but after 24 hours there will be a noticeable amount of protein spot diffusion. We therefore recommend that once the gel run is complete, labelled gels are scanned on the same day. The gels should be removed from the electrophoresis tank and placed in a light excluding box, containing running buffer at room temperature. If it is not practical to scan gels on the same day, the gels are best stored at 4 oC in the dark, in running buffer and scanned within 24 hours. Fluorescence is temperature dependent so it is important that the gels are allowed to warm to room temperature prior to scanning.
It is not recommended that analytical gels are fixed as fixing affects the fluorescence of the CyDye DIGE Fluor minimal dyes. Fixed gels may not provide accurate quantitative information about protein levels.
After scanning, preparative gels can be fixed and stored in 5% ethanol (acid-free) until required for picking. Once fixed, gels can be stored for up to 1 week before picking. Storage for longer periods of time is not recommended as this may affect the number of proteins which can be identified by mass spectrometry.
Q: My experiment requires me to run 48 gels but we only have access to one Typhoon Variable Mode Imager. How can I ensure that all my gels are scanned within 24 hours of running?
A: We recommend that gels are run overnight in batches of 12 (i.e. one Ettan DALT run per day). Gels can then be scanned the following day when the electrophoretic separation has finished. An experiment of this size would require staggered gel runs over four days, followed by scanning on the day each run is completed. For IEF, all 48 strips can be focused simultaneously and the strips stored at –15 oC to –30 oC until required for 2-D electrophoresis.
Q: Why can’t I use commercially available plastic backed precast gels with CyDye™ DIGE Fluor minimal dyes?
A: These gels come attached to a plastic backing, which generally has high background fluorescence. This inherent fluorescence of the plastic will significantly increase the background level, lower the dynamic range and reduce the number of protein spots seen in the gel.
Q: What is the effect of changing the equilibration time prior to second dimension separation?
A: We recommend an equilibration time of 10 min for each equilibration solution. This can be increased up to 15 min but equilibration times over 15 min can result in vertical streaking due to over-alkylation. Equilibration times below 10 min should not be used as these are likely to lead to poor protein entry into the gel.
Q: Does it matter if I do not keep the first or second dimension run in the dark?
A: On a practical level, it will make very little difference to the results unless very strong light levels are present. It is good laboratory practice to avoid exposing fluorescent dyes unnecessarily to light to reduce the risk of photobleaching. We therefore recommended that you do keep the first and second dimension gel runs in the dark. Light-excluding Ettan™ IPGphor lid covers are available for Ettan IPGphor Isoelectric Focusing Unit (product code: 80-6496-48).
Q: I can see horizontal lines across my gel images, even if I do not load a sample?
A: Make sure that you pour the gels in one continuous motion without stopping and starting to adjust the final level of the mix.
Q: I have lots of dirt specks in my images.
A: The most common cause of this problem is dust particles from wipers used to dry the plates after washing. It is best to rinse plates in high quality distilled water and leave them to air-dry in a rack. If you have to wipe the plates, use lint-free wipes (Kimberly-Clark, product code 33370).
A second reason that may explain dirt specks in your images could be dust in the reagents. Prior to adding TEMED and APS, filter the acrylamide gel mix through a 0.2 mm filter before pouring the gels.
A third possibility could be crystalization of the post-stain if the gel has been post-stained with SYPRO™ Ruby. To minimize this problem, wash the gel in four changes of high quality distilled water over a 2 h period after staining in SYPRO Ruby. If the gel is attached to the glass plate with PlusOne™ Bind-Silane, the gel should not expand. If the gel is not attached to the glass plate, the gel will need to be incubated in gel fix until it shrinks back to the original size before scanning.
Q: What precautions can I take to ensure that my analytical and preparative gels have as similar spot patterns as possible?
A: To ensure preparative gels are as similar to analytical gels as possible a number of protocol variations can be tried. The sample can be processed after labelling to improve levels of protein solubilisation before IEF. For example, sonication of samples immediately prior to IEF (i.e. rehydration buffer, DTT and pharmalytes already added) can help to resolubilise proteins. Proteins can also be resolubilised after labelling by the addition of organic solvents (e.g. isopropanol) or increasing the concentration of IEF-compatible detergent (e.g. ASB14) in the sample. More consistent spot patterns can also be obtained by focusing preparative and analytical IPG strips together and using 2-D gel cast at the same time. Alternatively, a high loading of unlabelled protein can be run within a couple of analytical gels, and thus separated in the same 1st and 2nd dimension as the labelled protein. Following analysis of the CyDye DIGE Fluor labelled images, the gel is post-stained and scanned prior to matching to the analytical data-set. Use of narrow range strips (for analytical and preparative gels) can also improve matching as there are fewer spots and the spot separation is better, so spot patterns tend to be more consistent between different protein loadings.Image acquisition
Q: Can I use my existing TyphoonTM Variable Mode Imager with EttanTM DIGE system?
A: Yes. Depending on the Typhoon model you can either use two or three different CyDye™ DIGE Fluor minimal dyes. The Typhoon 9200 series will optimally scan CyDye DIGE Fluor, CyTM3 and Cy5 minimal dyes whereas the Typhoon 9400 series will optimally scan CyDye DIGE Fluor, Cy2, Cy3, and Cy5 minimal dyes. There is a software upgrade and accessory kit available that will improve the usability further.
Typhoon 8600 series models may also be used for CyDye DIGE Fluor Cy3 and Cy5 minimal dyes but will need a hardware upgrade before the software upgrade can be installed. Please contact your local Amersham Biosciences sales representative for more details.
Q: When imaging the SYPRO™ Ruby stained gel, can I still image for the original CyDye™ DIGE Fluor minimal dyes and get three (or four) related images?
A: No. SYPRO Ruby has very broad excitation and emission wavelengths. Once the gel has been stained with SYPRO Ruby, if you try and scan again using CyDye DIGE Fluor parameters, the SYPRO Ruby will be excited by the 532 nm laser (used for Cy3) and emit light, interfering with quantitation in the Cy3 and Cy5 channels.
Q: CyDye™ DIGE Fluor Cy™2, Cy3 and Cy5 minimal dyes have different quantum yields. Does this effect the accuracy with which protein abundance changes can be quantified?
A: No. The quantum yield of Cy5 is greater than that of Cy2 and Cy3 so the signal from a sample labelled with CyDye™ DIGE Fluor Cy5 minimal dye would be expected to be the most intense. Selecting the correct PMT when scanning will ensure that the signal for each dye falls within the linear dynamic range of the Typhoon Variable Mode Imager. For example, using a higher PMT for Cy3 images should result in similar signal intensities in both channels. Images are then normalized in DeCyder Differential Analysis Software by setting the total signal for each image at the same level. This normalization process negates differences in signal intensity between the dyes.
Q:How do I clean the Typhoon Variable Mode Imager platen and sample press?
A: The scanning of gels (especially naked gels) and blots containing stains or other fluorescent compounds may leave fluorescent residues on the platen and/or sample press. These may show up in subsequent scans, obscuring or altering the intensity/profile of protein spots in the 2-D DIGE gel. To achieve best results from 2-D DIGE gels, ensure the platen and sample press are clean before scanning.
The platen is usually cleaned by using Milli-Q water and then dried using tissues, followed by lint-free wipes (Kimberly-Clark, code no. 33370). To remove insoluble material, ethanol can be used instead of water. For stubborn residues, wipe the platen with a tissue dampened with 10% hydrogen peroxide. Rinse well several times with Milli-Q water. It is important to rinse thoroughly at this stage. Failure to do this may leave hydrogen peroxide on the platen which could cause bleaching of fluorescent dyes used in subsequent scans. Dry the platen with a lint-free tissue.
The sample press can be cleaned using the same procedures as the platen, described above. However, liquid should not be squirted directly onto the sample press surface. Instead, apply the appropriate cleaning fluid (Milli-Q water, ethanol or hydrogen peroxide) to a tissue and then wipe the surface of the sample press. Dry using a lint-free tissue.
Q: The images for each color channel do not seem to overlay properly anywhere on the gel. What is the cause of this?
A: Ensure that you have checked the sample press box on the Typhoon Variable Mode Imager. Movement of the laser during scanning will cause the instrument to vibrate slightly. If the sample press option is not selected, these vibrations may cause the plates to move during the scan. This may result in a slightly shifted image between the first and second color channels which are always scanned in succession for analytical scans. An example where plate slippage has occured due to failure to use the sample press is shown below.
Figure 1. Image of a gel where the user failed to select the sample press option on the Typhoon Variable Mode Imager.
To troubleshoot a gel where you think this may have happened, rescan the gel selecting the “speed” option for auto link mode within the Fluorescence setup window in the Typhoon Scanner Control Software. This will link Cy™3 and Cy5 scans, so both images are taken simultaneously. If the channels show good overlay for this linked scan, then the poor overlay in the original scan was due to a failure to select the sample press option.
Note: the “speed” option should never be used for analytical scans that are to be quantified using DeCyder Differential Analysis Software. Linked scans experience cross-talk between Cy3 and Cy5 channels so the spot volumes may not accurately represent differences between the two dyes being
imaged.Dynamic range and sensitivity
Q: What is the limit of sensitivity and how was it determined?
A: Limits of sensitivity were determined on a 1-D gel with a single labelled protein. The limit is currently reported at 125 pg for transferrin using the CyDye™ DIGE Fluor minimal dyes.
Q: What factors affect the dynamic range?
A: The dynamic range of the system is fixed. The TyphoonTM Variable Mode Imager offers a linear dynamic range of five orders of magnitude. Altering the photomultiplier tube (PMT) voltage will shift this dynamic range window to accommodate gels with different levels of fluorescent intensity. The capability of CyDye™ DIGE Fluor minimal dyes, the fluorescent background of the gel, and other factors will influence the effective dynamic range.
Q: How does the sensitivity compare with other detection methods?
A: CyDye™ DIGE Fluor minimal dyes, silver staining and SYPRO™ Ruby staining are all comparable. For more information please refer to the following paper: Gharbi et al. Evaluation of 2D-DIfferential Gel Electrophoresis for proteomic expression analysis of a model breast cancer cell system. Mol. Cell Proteomics 1, 91–8 (2002). Image analysis and statistics
Normalization
Q: How important is it to achieve the same or similar maximum intensity of pixel values in the multiplexed CyDye™ DIGE Fluor minimal dye images in order to get reliable results using DeCyder™ Differential Analysis Software?
A: For example, say that for one image, the maximum pixel value is 35 000 and the second image is 45 000. How can we be sure we can compare these two images with reliability?
Pixel values of 35 000 and 45 000 will cause no problem in the comparison of the images. However, it is always good to use as much of the dynamic range as possible as this will increase the sensitivity of the system. Typhoon™ imager has a very broad linear range, therefore it is possible to accurately detect even low signals.
Of course it would be preferable to aim for similar signals for both images. This will improve data quality and reduce reliance on DeCyder Differential Analysis Software to perform normalization. However, DeCyder Differential Analysis Software will normalize images even if they have very different pixel values, and the outcome should not be affected by virtue of the internal standard present in each gel.
Ideally, we recommend that the maximum pixel value is optimized on the first two gels. Aim to achieve a maximum pixel value between 50 000 and 70 000 in each of these gels. This will give scope in subsequent gels to avoid saturating spots (values above 100 000) or producing very low maximum pixel values which may lead to poor detection of low abundance material.
Q: When you use DeCyder™ Differential Analysis Software to calculate spot volumes, how do you normalize the absolute values so that they can be compared from one gel to another?
A: Spot volumes are normalized within a co-detected image pair in DIA using the blue model histogram that is visible in the graph (see Fig 2). A plot is generated of the number of spots against log (volume ratio) and this should show a normal distribution (red line in Fig 2). As the samples are related, most of the spots will show no change (ratio = 1, log[volume ratio] = 0) so the peak should be centered on 0. The software locates this peak, fits it to a model histogram, and then centers this at zero.
Fig 2. Normalization of spots in DIA.
In BVA, gel-to-gel spot ratios are compared. Because these ratios are of a sample compared to a common standard, they do not need to be normalized further. The ratio is a relative measurement against a standard. For a more detailed explanation of DeCyder Differential Analysis Software, please refer to the DeCyder Differential Analysis Software User Manual.Statistical approach to 2-D DIGE
Q: Can we compare the intensity of different protein spots and relate that to their abundance?
A: Within a single gel it is not possible to compare the intensity of different protein species. Fluorescence can become non-linear for some highly abundant spots and when using broad-range IPG strips, many spots have overlapping boundaries. Neither of these phenomena effect quantitation between different images in the same gel (i.e. for 2-D DIGE experiments) but they do prevent accurate quantitation between spots in the same image.
Q: Can we choose our own experimental design and still use the DeCyder™ Differential Analysis Software?
A: You can choose your own experimental design and use DeCyder Differential Analysis Software. However to match across gels and achieve valid statistical results, the internal standard must be run on each gel. We recommend a pooled internal standard as of all the samples within the experiment. In this way, every protein in the experiment is represented within each gel.
Q: Why does using a pooled internal standard increase the precision of the technology?
A: With any biological technique the precision is greatly improved if an internal standard is used. This is one of the key benefits of 2-D DIGE, the only 2-D electrophoresis technique that allows the user to run an internal standard. When using a pooled internal standard, every protein in the population appears on each gel. Each experimental sample can then be compared internally to the same standard, thereby accounting for any gel-to-gel (system) variation. This facilitates fully automated data processing and accurate spot statistics when using DeCyder Differential Analysis Software. The inclusion of an internal standard on each gel also aids matching between gels.
Occasionally, protein expression changes between samples are so large that the expression of specific proteins can be switched off. The use of the pooled internal standard with spot co-detection of multiplexed samples avoids any problems caused by presence-absence events. The pooled internal standard would always contain every protein within the experiment enabling the absence of specific proteins in individual samples to be analysed accurately with confidence.
For more information please refer to the following paper, "A novel experimental design for comparative two-dimensional gel analysis: two-dimensional difference gel electrophoresis incorporating a pooled internal standard.", Alban A., David S., Bjorkesten L., Andersson C., Sloge E., Lewis S. and Currie I. Proteomics 2003, Vol 3 (1) 36-44.
Q: Can you explain what the system variability of the technique is?
A: We ran five replicate gels containing a standard labelled with Cy™3 and a sample labelled with Cy5. If there were no variations, the spot ratio from gel-to-gel of standard against sample would be exactly the same for every spot, i.e. 0% variation. We compared the ratios for each spot across the five gels. The average standard deviation (SD) of the ratios for all spots was 1.05-fold or a 5% variance. Protein levels needed to differ by at least 1.13-fold (13% variance) to be significant at the 95% confidence level (2 SD). This compares with the conventional 2-DE approaches where it has been shown that protein levels may need to differ by at least 1.947-fold (94.7% variance) to be significant at the 95% confidence level (2 SD). For more information, refer to Asirvatham et al. Analytical and biological variances associated with proteomic studies of Medicago truncatula by two-dimensional polyacrylamide gel electrophoresis. Proteomics 2, 960–968 (2002).DeCyder™ Differential Analysis Software; general questions
Q: What statistical tests are available in DeCyder™ Differential Analysis Software?
A: The statistical tests available in the software include: Independent and paired two-tailed Student's t-test (assuming independent variances) in addition to 1-way and 2-way ANOVA (analysis of variance).
Q: What is Model Standard Deviation?
A: In DIA, a plot is generated of the number of spots against log[volume ratio] and this should show a normal distribution (red line in graph view, DeCyder™ DIA see Fig 2). As the samples are related, most of the spots will show no change (ratio = 1, log[volume ratio] = 0) so the peak should be centered on 0. The software locates this peak, fits it to a model histogram and then centers this at zero. The model standard deviation is the standard deviation of the model histogram (blue line in graph view, DeCyder DIA see Fig 2). This gives the user an idea of the variation associated with the largest peak of spot ratios. For further information please refer to the Ettan DIGE System User Manual.
Q: In BVA what does the standardized log abundance refer to?
A: To quantify spot abundance, each spot value is compared as a function of the internal standard, i.e, the ratio value of a spot pair. This is referred to as the standardized abundance and this process occurs within the DIA module of DeCyder™ Differential Analysis Software. Within DeCyder BVA module, spot maps from different gels are analyzed. The log10 of the standardized abundance, referred to as the standardized log abundance, is used to aid scaling and ensures that the data set is normally distributed, a prerequisite for the statistical analysis.
Q: Is spot editing required in DeCyder™ Difference Analysis Software?
A: A spot merging function is available in version 5.0 of DeCyder Differential Analysis Software. All versions of the software allow exclusion of gel artefacts and editing of matches can be performed in DeCyder BVA module.
Q: Is there manual override for gel to gel matching in the BVA?
A: Yes. Spot matching can be manually performed in the Match Table (MT) view.
Q: Are there any data export features?
A: Data can be exported in an XML format from both the DIA and BVA modules. The DeCyder™ XML toolbox enables users to extract and convert the exported data into tab separated text or html files for downstream report generation or database storage.
Picklists, which are text files utilized by the Ettan™ Spot Picker, can be exported from both the DIA and BVA modules. In addition, users can program their own functionality into the XML toolbox.
Q: Can you annotate the gel images?
A: Yes, gel images can be annotated on screen in BVA with the master spot number, protein ID, comment or denote a protein of interest.
Q: Sometimes I see some really noisy peaks in the background — is it worth including them in the analysis if it looks like there really is a peak there, not just random noise? Do you have a recommended peak height or volume below which you would not include those "peaks"?
A: We do not have a recommended peak height or volume as it will be different for different sample types and gel sizes. Excluding real peaks causes more problems than leaving in non-protein peaks. If in doubt, leave it in.
Q: Why is it that sometimes peaks that appear on the gel image as very dark spots do not show up as tall peaks in the 3-D image or have a smaller volume than might be expected based on the intensity in the gel image?
A: The gel image is subject to contrast adjustment so what may appear as an intense spot may not really be so. Also, the 3D view is adjusted relative to the area surrounding the spot. This means that if a spot of interest is next to a much more intense spot the 3D view is scaled to the most intense spot and the spot of interest may appear as a very short peak.
Q: If a dust peak happens to fall right on top of a peak of interest, should that spot be included in the analysis even though the dust particle will skew the results for that peak?
A: Dust particles effect the accuracy of quantitation and the Student’s T-test result but you may wish to include the spot in the analysis since it occurs on a protein spot that is of interest. Therefore you should include it and add a comment in the spot comment field such as ‘includes dust’. If you have replicate gels this result can then be checked to see if it is a consistent difference in the other gels, either by matching in BVA or manually having a look at the other gels in DIA. In DeCyder Differential Analysis Software, quantitation is based upon spot volume. The volume of a dust particle is very small in comparison to the majority of spots and therefore dust should not impact too greatly on the analysis of that particular spot. However, it should be remembered that low abundance proteins, with smaller spot volumes, will be affected to a greater extent. Subsequently if the dust particles skews the accuracy of quantitation by a large amount it should ultimately be excluded from the analysis. Ultimately, if the T-test result is significant, the effect of the dust particle can be ignored.Spot picking and downstream analysis
Q: When imaging a SYPRO™ Ruby stained gel, can I still image for the original CyDye™ DIGE Fluor minimal dyes and get three (or four) related images?
A: No. SYPRO Ruby has very broad excitation and emission wavelengths. Once the gel has been stained with SYPRO Ruby, if you try and scan again using CyDye DIGE Fluor parameters, the SYPRO Ruby will be excited by the 532 nm laser (used for Cy™3) and emit light, interfering with quantitation in the Cy3 and Cy5 channels.
Q: If I define an Area of Interest, does it have to include the reference markers?
A: No. In DeCyder DIA, after spot detection, define the area of interest which should include all the protein spots on the preparative gel, but may or may not include the reference markers. Then run the Spot Filter to remove all spots outside the Area of Interest. At this stage, the user can also include their own filter parameters to simultaneously remove gel artefacts within the area of interest. The reference markers are taken as a separate entity and are manually assigned using the “define reference markers” option under the edit menu. Any protein spots outside the area of interest can be added back by clicking on them and selecting include and confirm.
Q: Do the CyDye™ DIGE Fluor minimal dyes cause suppression in MS ionization?
A: Generally, spot picking is performed on gels containing unlabelled samples stained with SYPRO™ Ruby. If you wish to perform MS on gels containing labelled proteins, we have no evidence that the dyes will cause suppression in MS ionization. However, the majority of proteins within the sample will remain unlabelled due to the minimal labelling approach. Therefore it is unlikely that a labelled protein will be detected by the mass spectrometry analysis.
Q: Is the sensitivity of mass spectrometry affected by CyDye™ DIGE Fluor minimal dye labelling?
A: We have no evidence of this.System validation
Q: How many different sample types have been analyzed using the CyDye™ DIGE Fluor minimal dye labelling approach?
A: Currently we know of at least 28 different samples that have been tested using this technique. These include mammalian cell lines, mammalian tissue, mammalian biopsy, bacteria, yeast, plant, Drosophila, and biological fluids such as plasma and serum. For a complete list please refer to the Ettan DIGE System User Manual.
Q: Are there any sample types that cannot be labelled with this technology?
A: We have not found any to date. Ensure that proteins are in a recommended lysis buffer (8 M urea, or 7 M urea/2 M thiourea, 4% CHAPS, 30 mM Tris pH 8.5) and there should be no problem. For best results, the concentration of protein in the labelling solution should be 1–10 mg/ml. The optimal concentration is 5 mg/ml.
Q: What publications are available to validate use of this method?
A: We have a list of publications and citations on this website. |
