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We do a lot of bioconjugation chemistry (click chemistry in particular but also NHS and Maleimide chemistries). Our method to valid the conjugation reactions have been to use SDS-PAGE gels followed with densitometry. However one particular challenge with such analyses is that proteins conjugated with non-proteins like DNA or PEG migrate with an inconsistent MW and also in a smear making it hard to isolate the relevant bands. I assume that this has to do with having a poly-disperse radius of gyration. Are there tricks that I can do to get tighter bands?
The first step in troubleshooting is to run controls - run lanes with your protein input alone, and the conjugate alone (you may need to play with the type/percentage of the gel if the conjugate molecule is much smaller than your target protein - attaching biotin to a 50 kDa protein, for example) to see if the smear shows up. Depending on the type of conjugation you're using, the number of acceptor sites on the substrate, and the time and conditions of the conjugation reaction, you may be adding a highly variable number of conjugate molecules to your substrate. For example, when I first started working with adding fluorophores like Alexa and DyLight dyes to antibodies, the fluor/protein ratio was highly variable from reaction to reaction and between antibodies in the same reaction.
This is most likely what is happening in your case - the number of conjugate molecules being added to each substrate molecule is highly variable. If you haven't already, I'd suggest getting in touch with your conjugation kit vendor's tech support, as they likely encounter this problem all the time. You may need to change the type of conjugation chemistry, get a higher-quality conjugate, or purify the substrate before and/or after the reaction. One of the solutions to my antibody labeling problem I mentioned above was to purify the reaction products through a size-exclusion column. This had the dual effect of separating out the un-reacted antibody and fluorophore, and allowing us to pick different fractions of the conjugated product and decide which were best for our needs.
SDS-PAGE is a reliable method for determining the molecular weight (MW) of an unknown protein, since the migration rate of a protein coated with SDS is inversely proportional to the logarithm of its MW. The key to accurate MW determination is selecting separation conditions that produce a linear relationship between log MW and migration within the likely MW range of the unknown protein. See Molecular Weight Estimation Protocol for more information.
To ensure accurate MW determination:
- Separate the protein sample on the same gel with a set of MW standards (see Protein Standards for information regarding selection of protein standards)
- For statistical significance, generate multiple data points (>3 gels)
- Use a sample buffer containing reducing agents (DTT or &beta-ME) to break disulfide bonds and minimize the effect of tertiary structure on migration
- Include SDS in the sample buffer. SDS denatures secondary, tertiary, and quaternary structures by binding to hydrophobic protein regions, and it confers a net negative charge on the proteins, which results in a constant charge-to-mass ratio SDS binds proteins in the proportion of 1.4 g SDS/g protein
After separation, determine the relative migration distance (Rf) of the protein standards and of the unknown protein. Rf is defined as the mobility of a protein divided by the mobility of the ion front. Because the ion front can be difficult to locate, mobilities are normalized to the tracking dye that migrates only slightly behind the ion front:
Rf = (distance to band)/(distance to dye front)
Using the values obtained for the protein standards, plot a graph of log MW vs. Rf (see figure below). The plot should be linear for most proteins, provided they are fully denatured and that the gel percentage is appropriate for the MW range of the sample. The standard curve is sigmoid at extreme MW values because at high MW, the sieving effect of the matrix is so large that molecules are unable to penetrate the gel at low MW, the sieving effect is negligible and proteins migrate almost freely. To determine the MW of the unknown protein band, interpolate the value from this graph.
The accuracy of MW estimation by SDS-PAGE is in the range of 5&ndash10%. Polypeptides like glyco- and lipoproteins are usually not fully coated with SDS and will not behave as expected in SDS-PAGE, leading to inaccurate molecular weight estimations. For more details about protein molecular weight determination using SDS-PAGE, refer to bulletin 3133.
Typical characteristics of a log MW vs. Rf curve for protein standards.
Running agarose and polyacrylamide gels
One of the most widely used tools in molecular biology, electrophoresis provides a simple, low-cost way to separate nucleic acids based on size for quantification and purification. Get some tips on running your gels. From here you can also access a detailed PAGE troubleshooting guide.
Electrophoresis with agarose and polyacrylamide gels is one of the most widely used tools in molecular biology. Gels provide a simple, low-cost way to separate nucleic acids based on size for quantification and purification.
Agarose gels can be used to resolve large fragments of DNA. Polyacrylamide gels are used to separate shorter nucleic acids, generally in the range of 1&minus1000 base pairs, based on the concentration used (Figure 1). These gels can be run with or without a denaturant. Gels that are run without a denaturant are referred to as native gels. The DNA or RNA will migrate at different rates, depending on its secondary structure. Native gels allow the DNA or RNA to remain double stranded. Adding a denaturant to the gel, such as urea, will generally make all of the nucleic acids single stranded. Secondary structure will not form in denaturing gels and, therefore, only the length of the DNA will affect mobility.
Different concentrations of agarose and acrylamide are necessary to optimize resolution of nucleic acids with different lengths. Suggested concentrations are shown below in Table 1.
Table 1. Gel concentrations for size separation.
|Agarose Gels||Polyacrylamide Gels|
|% agarose||Size Range for Optimum Resoultion (bp)||% acrylamide||Size Range for Optimum Resoultion (bp)|
Acrylamide is a potent neurotoxin and, in its powdered form, can easily be aerosolized. Make sure to wear the appropriate personal protection, including gloves and a mask, when weighing out the material. Many companies sell acrylamide dissolved in water or pre-cast gels. These products are slightly more costly but reduce the risk of acrylamide inhalation.
Ethidium bromide is the most common DNA stain available it is also toxic if inhaled, decomposes when heated to produce toxic gases, and is suspected of causing genetic defects . Always wear gloves and avoid microwaving liquids containing ethidium bromide. Non-mutagenic fluorescent dyes available as an alternative include Bio-Safe&trade (Bio-Rad), SYBR-safe&trade (ThermoFisher), and GelRed Nucleic Acid Stain (Phenix Research Products). While more costly than ethidium bromide, these stains reduce the need for isolating and decontaminating gel electrophoresis stations. Note that ethidium bromide only intercalates into double-stranded DNA and, therefore, is not a good stain for single-stranded DNA analysis.
Tips for acrylamide gel electrophoresis
- Use fresh Ammonium Persulfate (APS). APS catalyzes the polymerization of acrylamide. Using old APS or APS stored above -20°C will result in slow or incomplete polymerization. Keep small, fresh aliquots in the freezer.
- Know how your tracking dye(s) will migrate.In agarose gels, Bromophenol Blue and Xylene Cyanol will migrate at approximately 3000 and 300 bp, respectively. These dyes will migrate at different rates in acrylamide gels depending on the gel density. Table 2 provides the approximate migration rate in terms of the relative size of single-stranded/denatured DNA.
- TAE or TBE? Agarose gels commonly use Tris-Acetic Acid-EDTA (TAE) or Tris-Boric Acid-EDTA (TBE) buffers. TAE buffer has the advantage that it can be made in 50X stock solutions. However, it buffers less efficiently and, in some cases, its use results in smeared bands. TBE buffered gels yield sharper bands, particularly when using small-sized DNA fragments, and can be run at higher voltages. However, the borate in TBE can inhibit some enzymes&mdashincluding T4 DNA ligase&mdashin DNA purified from these gels.
- What voltage to use? Agarose gels can be run at a large range of voltages&mdashfrom 0.25&ndash7 V/cm. High voltages save time but can result in overheating of the gel, even leading to melting of low percentage agarose gels. High voltages can also cause band smearing, particularly of fragments >10 kb. The sharpest bands can be obtained by running gels in TBE overnight at 0.25&ndash0.5 V/cm.
Table 2. Dye migration rates in acrylamide gels.
|Nondenaturing gels||Denaturing gels|
|% Acrylamide||Bromophenol Blue (nucleotides)||Xylene Cyanol (nucleotides)||Bromophenol Blue (nucleotides)||Xylene Cyanol (nucleotides)|
*Adapted from Sambrook J, Fritsch EF, Maniatis T. (1989) in: Molecular Cloning: A Laboratory Manual, Cold Spring harbor Laboratory.
Troubleshooting gel electrophoresis
- Blurry bands? Too much DNA or excess salt will create smeared bands and/or streaking in the gel. Loading the correct amount of DNA (usually a maximum of 100&minus250 ng/mm well width) and desalting samples with a spin column prior to loading will prevent this.
- Bands in the wrong place?Do not heat nucleic acids before running on a native gel, and do not exceed 20 V/cm (measured from anode to cathode, rather than entire gel length) or allow the gel to exceed 30°C. For the sharpest bands, run the gel slowly, at 5 V/cm.
- Loading buffer floats away? Rinsing wells with running buffer just before loading is essential failure to do so may prevent the loading mixture from sinking to the bottom of the well, resulting in an uneven band and delayed migration.
Adam Clore, PhD, Director of Synthetic Biology Technical Support & Development, IDT.
Published Jun 17, 2011
Revised/updated Sep 20, 2017
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SDS-PAGE Question: Uneven dye front?
I've posted here a few times about SDS-PAGE issues I've been having with success in the past so here goes again.
Ran two gels today. Checked on them after 30 minutes and the dye front is uneven. One of them looks like a slight frown and is probably okay but the other looks totally distorted. What are some causes of this? Any help would be greatly appreciated.
Update: After running for more than hour the machine gave an error message and stopped. The ladder made it maybe half way down the gel. Seriously have no idea what could be causing this. Samples were loaded fine. As far as I know the gels were fine - I made sure that they had solidified, etc. I make my gels using the 10% recipe. I usually pour two at a time. Any ideas would be awesome.
EDIT: Looked up what the error message said. It apparently means "no load detected." Still have no clue. The samples were pretty dilute but I ran gels with these same samples not long ago with no issues.
Troubleshooting bioconjugates migration in a SDS-PAGE gel? - Biology
Western Blot Troubleshooting
The following troubleshooting guide is summarized to explain causes and possible solutions for common problems observed in western blot (WB) assay. Though the tips provided here cover many different problems you may encounter in WB, we hope that you will find the information beneficial to you and useful as a reference guide. The intact Western Blot Protocol is available for you or view WB procedure step-by-step. For further assistance, please contact us.
No Signal or Weak Signal
- Increase the concentration of the primary antibody.
- Increase the incubation time to 4°C overnight.
- Use fresh antibody to improve signal.
- Load more protein.
- Choose the appropriate lysis buffer.
- Use immunoprecipitation (IP) if necessary to increase the concentration of a non-abundant protein.
- Add protease inhibitors into the lysis buffer.
- Make sure the sample has not degraded.
- Make sure the proteins were successfully transferred to the membrane.
- Confirm equal transfer by analyzing loading control expression or positive control.
High Uniform Background
- Increase blocking time and/or temperature.
- Increase the concentration of blocking reagent.
- Changing the blocking agent.
- Lower the concentration of the antibody.
- Add blocking agent in antibody buffers.
- Confirm the secondary antibody is specific by performing a secondary antibody control.
Non-specific Bands/Wrong Size or Multiple Bands
Of all methods available for protein quantitation (including UV spectroscopy at 280 nm, colorimetric dye-based assays, and electrophoresis in combination with image acquisition analysis), only protein quantitation by electrophoresis enables evaluation of purity, yield, or percent recovery of individual proteins in complex sample mixtures.
Two types of quantitation are possible: relative quantitation (quantitation of one protein species relative to the quantity of another) and absolute quantitation (quantitation of a protein by using a calibration curve generated by a range of known concentrations of that protein). Because proteins interact differently with protein stains, staining intensity of different proteins at identical protein loads can be very different, so only relative quantitative values can be determined in most cases. Absolute protein measurements can be made only if the protein under investigation is available in pure form and used as calibrant.
Troubleshooting: Weak/No Signal & Other
Titrate the antibody to determine optimum concentration.
The antibody may have lost activity – perform a dot blot to determine activity and optimal concentration.
Include a positive control (e.g., overexpressed protein, purified protein, positive cell line, etc. Adjust protein loading accordingly).
Change incubation time and temperature (4°C, overnight).
Target protein abundance is too low
Load more protein per well.
Enrich low-abundance proteins by immunoprecipitation, fractionation, etc.
Use appropriate treatment to induce target protein expression or modification.
Ensure sample has not degraded.
Include protease inhibitors in the lysis buffer.
Use the optimum lysis buffer for the target protein’s subcellular localization.
Check protein loading with an internal loading control antibody.
Select PVDF or NC membranes based on hydrophobicity/ hydrophilicity of the target antigen.
Blocking buffer issues
Blocking for too long can mask specific epitopes and prevent antibody binding.
Reduce the percentage of, or remove, the blocking reagent from the antibody incubation buffers.
Switch to using an alternative blocking reagent.
Low molecular weight targets
Use a Tris-tricine gel for protein targets <20kDa.
Reduce transfer times and/or use smaller pore size membranes (0.22 μm) for low MW proteins <30kDa.
Wet transfer is recommended for small proteins (10kDa).
Ensure proper transfer set-up (e.g., no air bubbles trapped between the gel and the membrane).
Thicker gels can result in incomplete transfer of high molecular-weight-proteins.
Check the quality of protein transfer with a reversible, universal protein stain, e.g., Ponceau-S.
Wet transfer produces higher-resolution transfers over semidry transfer.
Sodium azide contamination
The presence of sodium azide inhibits the activity of HRP.
Use sodium azide-free buffers.
Ensure sufficient washing.
Film exposure too short / detection reagent not sensitive enough
Check several exposure times to achieve optimum detection.
Try different detection reagent compositions and/or brands.
Dilute chemiluminescent reagents in high-purity water.
Other blotting issues
Ghost hollow bands
Too much antibody
Inverse staining (i.e., white bands on a dark blot)
Too much primary and/or too much secondary antibody.
Use antibodies at higher dilutions.
Molecular weight marker staining
The antibody reacts with the MW marker.
Migration through the gel was too hot or too fast.
Reduce the voltage applied to run the SDS-PAGE gel or run the gel in a cold room.
Blank areas/white spots
High protein concentrations can result in diffuse protein bands.
Uneven protein loading: assay protein samples and load by protein amount. Check for even protein loading by stripping and reprobing the blot with an internal control antibody (or use an HRP-conjugated loading control antibody).
Uneven gel composition (gel has set too quickly while casting or buffer was mixed inadequately).
Uneven bands can be due to insufficient buffer being added to the tank during running.
This problem can be caused by antibodies binding to the blocking reagent in the blocking buffer.
Change to another blocking reagent.
Filter the blocking buffer.
Wash excess detection reagent from the membrane before exposure.
Find your product-specific protocols for WB, IHC, IP and more!
Optimize your experiment with our product-specific protocols for WB, IHC, IP, IF, and FC. You can search by either catalog number or antibody name.
Standard protocols are also available
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Where agarose gels are best for running larger molecules, like DNA, SDS-PAGE is better suited for running smaller ones, like proteins.
SDS-PAGE has a number of uses, which include:
- Establishing protein size
- Protein identification
- Determining sample purity
- Identifying disulfide bonds
- Quantifying proteins
- Blotting applications
SDS-PAGE stands for sodium dodecyl (lauryl) sulfate-polyacrylamide gel electrophoresis. The SDS portion is a detergent. You may recognize it if you read the ingredients lists on your shampoo, soap, or toothpaste. The purpose of the SDS detergent is to take the protein from its native shape, which is basically a big glob, and open it up into a linear piece. It's kind of like taking a wadded up ball of string and untangling it into one straight, long piece. This will allow it to run more efficiently down the gel and will get you better results, since it's easier to compare two linear pieces of something rather than two wads of the same thing.
In more scientific terms, it is an anionic detergent that binds quantitatively to proteins, giving them linearity and uniform charge, so that they can be separated solely on the basis if their size. The SDS has a high negative charge that overwhelms any charge the protein may have, imparting all proteins with a relatively equal negative charge. The SDS has a hydrophobic tail that interacts strongly with protein (polypeptide) chains. The number of SDS molecules that bind to a protein is proportional to the number of amino acids that make up the protein. Each SDS molecule contributes two negative charges, overwhelming any charge the protein may have. SDS also disrupts the forces that contribute to protein folding (tertiary structure), ensuring that the protein is not only uniformly negatively charged, but linear as well.
The polyacrylamide gel electrophoresis works in a similar fashion to an agarose gel, separating protein molecules according to their size. In electrophoresis, an electric current is used to move the protein molecules across a polyacrylamide gel. The polyacrylamide gel is a cross-linked matrix that functions as a sort of sieve to help "catch" the molecules as they are transported by the electric current. The polyacrylamide gel acts somewhat like a three-dimensional mesh or screen. The negatively charged protein molecules are pulled to the positive end by the current, but they encounter resistance from this polyacrylamide mesh. The smaller molecules are able to navigate the mesh faster than the larger one, so they make it further down the gel than the larger molecules. This is how SDS-PAGE separates different protein molecules according to their size.
Once an SDS-PAGE gel is run, you need to fix the proteins in the gel so they don't come out when you stain the gel. Acetic acid 25% in water is a good fixative, as it keeps the proteins denatured. The gel is typically stained with Coomassie blue dye R250, and the fixative and dye can be prepared in the same solution using methanol as a solvent. The gel is then destained and dried.
Western Blot Protocols (part 1) - Sample & Gel Preparation
Here we list some recommended sample extract method for several kinds of materials in Table 1.
Table 1. Overview of extraction options for different cells and tissues
|Sample||Typical lysis options|
|Tissue culture||Detergent lysis|
|Most plant and animal tissues||Mechanical homogenization|
|Soft animal tissues and cells||Manual or mechanical homogenization|
|Bacterial and mammalian cells||Freeze/thaw lysis|
|Bacteria, erythrocytes, cultured cells||Osmotic shock lysis|
|Solid tissues and plant cells||Manual grinding with mortar and pestle|
|Cell suspensions, yeast cells||Grinding with abrasive component|
|Bacteria, yeast, plant tissues, fungal cells||Enzymatic digestion|
|Bacteria, yeast, plant cells||Nitrogen cavitation|
|Microorganisms with cell walls||French press|
|Plant tissues, fungal cells||Glass bead milling|
The extracted protein must be dissolved and preserved in lysis buffer to further conduct the following electrophoresis process. There are many recipes for lysis buffers but a few will serve for most western blotting experiments. In brief, they differ in their ability to solubilize proteins, with those containing sodium dodecyl sulfate and other ionic detergents considered to be the harshest and therefore most likely to give the highest yield. However, these detergent can cause protein denature more or less. So if your antibody used cannot recognize denatured proteins, you should avoid the detergent in buffer such as SDS, deoxycholate, Triton X-100 and NP-40 or try to use some relatively mild non-ionic detergents.
Protease and phosphatase inhibitors
Protease inhibitors must be included in lysis buffers to prevent degradation of proteins following the release of endogenous proteases during the process of cell lysis. As soon as lysis occurs, proteolysis, de-phosphorylation and denaturation begin. These events can be slowed down tremendously if samples are kept on ice or at 4°C at all times and appropriate inhibitors are added fresh to the lysis buffer. Cocktails of inhibitors from various suppliers are available, but still you can make your own inhibitor mixture. Here we list some inhibitor formula in Table 2.
Table 2. Inhibitor formula
|Inhibitor||Protease/phosphatase inhibited||Final concentration in lysis buffer||Stock (store at -20°C)|
|Aprotinin||Trypsin, Chymotrypsin, Plasmin||2 μg/ml||Dilute in water, 10 mg/ml. Do not re-use once defrosted.|
|Leupeptin||Lysosomal||5-10 μg/ml||Dilute in water. Do not re-use once defrosted.|
|Pepstatin A||Aspartic proteases||1 μg/ml||Dilute in methanol, 1 mM.|
|PMSF||Serine, Cysteine proteases||1 mM||Dilute in ethanol. You can re-use the same aliquot.|
|EDTA||Metalloproteases that require Mg2+ and Mn2+||5 mM||Dilute in H2O, 0.5 M. Adjust pH to 8.0.|
|EGTA||Metalloproteases that require Ca2+||1 mM||Dilute in H2O, 0.5 M. |
Adjust pH to 8.0.
|Na Fluoride||Serine/Threonine phosphatases||5-10 mM||Dilute in water. Do not re-use once defrosted.|
|Na Orthovanadate||Tyrosine phosphatases||1 mM||Dilute in water. Do not re-use once defrosted.|
Preparation of lysate from cell culture:
Place the cell culture dish in ice and wash the cells with ice-cold PBS.
Drain the PBS, then add ice-cold lysis buffer (1 ml per 107 cells/100 mm2 dish/150 cm2 flask 0.5ml per 5x106 cells/60 mm2 dish/75 cm2 flask).
Scrape adherent cells off the dish using a cold plastic cell scraper, then gently transfer the cell suspension into a pre-cooled micro centrifuge tube.
Maintain constant agitation for 30 minutes at 4°C.
Spin at 16,000 x g for 20 minutes in a 4°C pre-cooled micro centrifuge.
Gently remove the tubes from the micro centrifuge and place on ice. Transfer the supernatant to a fresh tube kept on ice, and discard the pellet.
Preparation of lysate from tissues:
Dissect the tissue of interest with clean tools, on ice preferably, and as quickly as possible to prevent degradation by proteases.
Place the tissue in round-bottom micro centrifuge tubes or Eppendorf tubes and immerse in liquid nitrogen to “snap freeze”. Store samples at -80°C for later use or keep on ice for immediate homogenization.
300 ml lysis buffer rapidly to the tube, homogenize with an electric homogenizer, rinse the blade twice with another 2 X 300 ml lysis buffer, then maintain constant agitation for 2 hours at 4°C. Volumes of lysis buffer must be determined in relation to the amount of tissue present. Protein extract should not be too dilute, to avoid the need to load large volumes per gel lane. The minimum protein concentration is 0.1 mg/ml optimal concentration is 1-5 mg/ml.
Centrifuge for 20 minutes at 16,000 rpm at 4°C in a micro centrifuge for 20 minutes. Gently remove the tubes from the centrifuge and place on ice. Transfer the supernatant to a fresh tube kept on ice discard the pellet.
After extracting from cells or tissue lysate, protein samples should be quantification so that to compare the amount of protein from samples run in different lanes within the same gel or between gels and make all the lanes have been loaded with the same total amount of protein. Several spectrophotometric methods are routinely used to determine the concentration of protein in a solution. These include measurement of the intrinsic ultraviolet (UV) absorbance of the protein as well as methods based on a protein-dependent color change, such as the classic, copper-based Lowry assay, the Smith copper/bicinchoninic assay (BCA) and the Bradford dye assay.
Table 3 compare of three different protein quantification methods.
|The Lowry Method||Relies on the reaction of copper with proteins, but the sample is also incubated with the Folin-Ciocalteu reagent. Reduction of the Folin-Ciocalteu reagent under alkaline conditions results in an intense blue color (heteropolymolybdenum blue) that absorbs at 750 nm. The Lowry method is best used with protein concentrations of 0.01–1.0 mg/mL.||Easy to use |
Sensitive and broad linear range
|Need to do a standard curve for every assay |
Timing and mixing of reagents must be precise
|Bradford Assay||Binding of Coomassie Brillant Blue G-250 to proteins, causes a shift of the dye from red (465 nm) to blue (595 nm) under acidic conditions. It is compatible with more common reagents, although detergents can cause interference. Proteins with a concentration of 20-2000 μg/mL can be measured using the Bradford assay.||Easy to use |
Sensitive and broad linear range
Compatible with many buffers
|Need to do a standard curve for every assay |
Reagent stains cuvettes
Often need to dilute samples prior to analysis
Depends strongly on amino acid composition
|Bicinchoninic Acid Assay (BCA)||After reduction of Cu2+ ions, two molecules of BCA chelate with each Cu+ ion resulting in formation of an intense purple color that absorbs at 560 nm. BCA is as sensitive as the Lowry method and works well with protein concentrations from 0.5 μg/mL to 1.5 mg/mL.||Easy to use |
Sensitive and broad linear range
|Need to do a standard curve for every assay |
Color continues to develop over time, but is stable for measurement after 30 minutes at 37°C
Once the concentration of each sample have been determined, you can freeze them at -20°C or -80°C for later use or prepare for loading onto a gel or other usage.
Polyacrylamide gels are inert, crosslinked structures. The pore sizes in these gels are similar to the molecular radius of many proteins. As molecules are forced through the gel in an electric field, larger molecules are retarded by the gel more than smaller molecules. When separated on a polyacrylamide gel, the procedure is abbreviated as SDS-PAGE (for Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis). The technique is a standard means for separating proteins according to their molecular weight. Polyacrylamide gels are formed from the polymerization of two compounds acrylamide and N, N-methylenebis-acrylamide (Bis, for short). Bis is a cross-linking agent for the gels. The polymerization is initiated by the addition of ammonium persulfate along with either DMAP or TEMED. The gels are neutral, hydrophilic, three-dimensional networks of long hydrocarbons crosslinked by methylene groups. The separation of molecules within a gel is determined by the relative size of the pores formed within the gel. The pore size of a gel is determined by two factors: the total amount of acrylamide present and the amount of cross-linker. As the percentage of acrylamide increases, the pore size decreases. With cross-linking 5% gives the smallest pore size. Any increase or decrease in cross-linking percentage increases or decreases the pore size. Gels are designated as percent solutions and will have two necessary parameters. The total acrylamide is given as a percentage (w/v) of the acrylamide plus the bis-acrylamide. The average pore size is determined by the percentage of the amount of cross linker and total amount of acrylamide used. Polyacrylamide is used to separate most proteins, ranging in molecular weight from Mr >5000 to <200,000.
Table 4 Recommended acrylamide concentration for protein target within defined size ranges.
|Target protein size range (Mr)||Recommended acrylamide concentration|
|36 000 to 205 000||5%|
|24 000 to 205 000||7.5%|
|14 000 to 205 000||10%|
|14 000 to 66 000||12.5%|
|14 000 to 45 000||15%|
The percentage of polyacrylamide used in the gel along with the buffer system will influence the mobility of the proteins through the gel as current is applied. The expected size of the target protein can be used to select the best gel/buffer system to achieve optimal separation and resolution (Figure 2).
Figure 2. Migration of protein by molecular weight varies by gel type, concentration, and running buffer. Use the estimated migration patterns provided here to assist in the selection of gel type and running buffer based on the predicted MW of the target protein and the desired separation
The SDS-PAGE system we used is usually a discontinuous system. It consists of two different gel: the stacking gel and the separating/resolving gel (Figure 3.). Stacking gel usually with low pH (6.8) and Acr Bis concentration (4%), that makes the stacking gel have higher porosity and could hardly influence the movement of proteins. Separating gel with higher pH (8.8) and Acr Bis concentration (12.5%). Use a gel comb to form sample loading cells on stacking gel. Load the sample in the cells and switch on the power. By initially running the samples through a lower density stacking gel, proteins are concentrated in a matter of minutes into a thin starting zone by the time the sample contents reach the resolving gel by a process known as isotachophoresis. The interface between the two gel densities may thus be regarded as the starting line for all the proteins in each well and on entering the resolving gel, the proteins begin to separate according to size. A reference recipe of these two kind of gel preparation is shown in Table 5 and Table 6.
Friday, 7 March 2014
Evolution - Is there any evidence that sexual selection may lead to extinction of species?
There is a dearth of actual experimental evidence. However:
there is at least one study that confirmed the process ([STUDY #7] - Myxococcus xanthus by Fiegna and Velicer, 2003).
Another study experimentally confirmed higher extinction risk as well ([STUDY #8] - Paul F. Doherty's study of dimorphic bird species an [STUDY #9] - Denson K. McLain).
Theoretical studies produce somewhat unsettled results - some models support the evolutionary suicide and some models do not - the major difference seems to be variability of environmental pressures.
Also, if you include human predation based solely on sexually selected trait, examples definitely exist, e.g. Arabian Oryx
First of all, this may be cheating but one example is the extinction because a predator species specifically selects the species because of selected-for feature.
The most obvious case is when the predator species is human. As a random example, Arabian Oryx was nearly hunted to extinction specifically because of their horns.
Please note that this is NOT a simple question - for example, the often-cited in unscientific literature example of Irish Elk that supposedly went extinct due to its antler size may not be a good crystal-clear example. For a very thorough analysis, see: "Sexy to die for? Sexual selection and risk of extinction" by Hanna Kokko and Robert Brooks, Ann. Zool. Fennici 40: 207-219. [STUDY #1]
They specifically find that evolutionary "suicide" is unlikely in deterministic environments, at least if the costs of the feature are borne by the individual organism itself.
Another study resulting in a negative result was "Sexual selection and the risk of extinction in mammals", Edward H. Morrow and Claudia Fricke The Royal Society Proceedings: Biological Sciences, Published online 4 November 2004, pp 2395-2401 [STUDY #2]
The aim of this study was therefore to examine whether the level of
sexual selection (measured as residual testes mass and sexual size dimorphism) was related to the risk of extinction that mammals are currently experiencing. We found no evidence for a relationship between these factors, although our analyses may have been confounded by the possible dominating effect of contemporary anthropogenic factors.
However, if one takes into consideration changes in the environment, the extinction becomes theoretically possible. From "Runaway Evolution to Self-Extinction Under Asymmetrical Competition" - Hiroyuki Matsuda and Peter A. Abrams Evolution Vol. 48, No. 6 (Dec., 1994), pp. 1764-1772: [STUDY #3]
We show that purely intraspecific competition can cause evolution of extreme competitive abilities that ultimately result in extinction, without any influence from other species. The only change in the model required for this outcome is the assumption of a nonnormal distribution of resources of different sizes measured on a logarithmic scale. This suggests that taxon cycles, if they exist, may be driven by within- rather than between-species competition. Self-extinction does not occur when the advantage conferred by a large value of the competitive trait (e.g., size) is relatively small, or when the carrying capacity decreases at a comparatively rapid rate with increases in trait value. The evidence regarding these assumptions is discussed. The results suggest a need for more data on resource distributions and size-advantage in order to understand the evolution of competitive traits such as body size.
As far as supporting evidence, some studies are listed in "Can adaptation lead to extinction?" by Daniel J. Rankin and Andre´s Lo´pez-Sepulcre, OICOS 111:3 (2005). [STUDY #4]
The first example is a study on the Japanese medaka
fish Oryzias latipes (Muir and Howard 1999 - [STUDY #5]). Transgenic males which had been modified to include a salmon growth-hormone gene are larger than their wild-type counterparts, although their offspring have a lower fecundity (Muir and Howard 1999). Females
prefer to mate with larger males, giving the larger
transgenic males a fitness advantage over wild-type
males. However, offspring produced with transgenic
males have a lower fecundity, and hence average female
fecundity will decrease. As long as females preferentially
mate with larger males, the population density will
decline. Models of this system have predicted that, if
the transgenic fish were released into a wild-type
population, the transgene would spread due to its mating
advantage over wild-type males, and the population
would become go extinct (Muir and Howard 1999).
A recent extension of the model has shown that
alternative mating tactics by wild-type males could
reduce the rate of transgene spread, but that this is still
not sufficient to prevent population extinction (Howard
et al. 2004). Although evolutionary suicide was predicted
from extrapolation, rather than observed in nature, this
constitutes the first study making such a prediction from
In cod, Gadus morhua, the commercial fishing of large
individuals has resulted in selection towards earlier
maturation and smaller body sizes (Conover and Munch
2002 [STUDY #6]). Under exploitation, high mortality decreases the
benefits of delayed maturation. As a result of this,
smaller adults, which mature faster, have a higher fitness
relative to their larger, slow maturing counterparts
(Olsen et al. 2004). Despite being more successful
relative to slow maturing individuals, the fast-maturing
adults produce fewer offspring, on average. This adaptation,
driven by the selective pressure imposed by
harvesting, seems to have pre-empted a fishery collapse
off the Atlantic coast of Canada (Olsen et al. 2004). As
the cod evolved to be fast-maturing, population size was
gradually reduced until it became inviable and vulnerable
to stochastic processes.
The only strictly experimental evidence for evolutionary
suicide comes from microbiology. In the social
bacterium Myxococcus xanthus individuals can develop
cooperatively into complex fruiting structures (Fiegna
and Velicer 2003 - [STUDY #7]). Individuals in the fruiting body are
then released as spores to form new colonies. Artificially
selected cheater strains produce a higher number of
spores than wild types. These cheaters were found to
invade wild-type strains, eventually causing extinction of
the entire population (Fiegna and Velicer 2003). The
cheaters invade the wild-type population because they
have a higher relative fitness, but as they spread through
the population, they decrease the overall density, thus
driving themselves and the population in which they
reside, to extinction.
Another experimental study was "Sexual selection affects local extinction and turnover
in bird communities" - Paul F. Doherty, Jr., Gabriele Sorci, et al 5858 PNAS May 13, 2003 vol. 100 no. 10 [STUDY #8]
Populations under strong sexual selection experience
a number of costs ranging from increased predation and
parasitism to enhanced sensitivity to environmental and demographic
stochasticity. These findings have led to the prediction that
local extinction rates should be higher for speciespopulations
with intense sexual selection. We tested this prediction by analyzing
the dynamics of natural bird communities at a continental
scale over a period of 21 years (1975), using relevant statistical
tools. In agreement with the theoretical prediction, we found
that sexual selection increased risks of local extinction (dichromatic
birds had on average a 23% higher local extinction rate than
monochromatic species). However, despite higher local extinction
probabilities, the number of dichromatic species did not decrease
over the period considered in this study. This pattern was caused
by higher local turnover rates of dichromatic species, resulting in
relatively stable communities for both groups of species. Our
results suggest that these communities function as metacommunities,
with frequent local extinctions followed by colonization.
This result is similar to another bird-centered study: Sexual Selection and the Risk of Extinction of Introduced Birds on Oceanic Islands": Denson K. McLain, Michael P. Moulton and Todd P. Redfearn. OICOS Vol. 74, No. 1 (Oct., 1995), pp. 27-34 [STUDY #9]
We test the hypothesis that response to sexual selection increases the risk of extinction by examining the fate of plumage-monomorphic versus plumage-dimorphic bird species introduced to the tropical islands of Oahu and Tahiti. We assume that plumage dimorphism is a response to sexual selection and we assume that the males of plumage-dimorphic species experience stronger sexual selection pressures than males of monomorphic species. On Oahu, the extinction rate for dimorphic species, 59%, is significantly greater than for monomorphic species, 23%. On Tahiti, only 7% of the introduced dimorphic species have persisted compared to 22% for the introduced monomorphic species.
Plumage is significantly associated with increased risk of extinction for passerids but insignificantly associated for fringillids. Thus, the hypothesis that response to sexual selection increases the risk of extinction is supported for passerids and for the data set as a whole. The probability of extinction was correlated with the number of species already introduced. Thus, species that have responded to sexual selection may be poorer interspecific competitors when their communities contain many other species.