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Troubleshooting SDS-PAGE of trypsin-treated BSA

Troubleshooting SDS-PAGE of trypsin-treated BSA


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I am currently working on SDS-PAGE technique having 20% acrylamide concentration for hydrolyzed BSA protein. Here I attached a gel photo for trypsin hydrolyzed BSA. I don't know , is it a good result and how do I interpreted this result?


I'm going to treat this as a partial homework question but provide some guidance as to how you can potentially address your question and have solid theory to back it up. Chymotrypsin preferentially cleaves peptide amide bonds where the carboxyl side of the amide bond (the P1 position) is a large hydrophobic amino acid (tyrosine, tryptophan, and phenylalanine). So one solution which is often used by experimental biologists (such as myself) performing mass spectrometry (MS) is to put their protein sequence (in this case BSA) in an algorithm such as PeptideCutter (from ExPASy), which performs a theoretical digest of the protein sequence having chosen the desired enzyme or combination of enzymes) and see what peptide fragments are expected to be produced and work out their length/molecular weight and compare that to your experimental data.

Having had an initial look, I noticed that your protein marker/ladder is missing, which is quite important in figuring out the peptide sizes produced! Also what is 1-8 lanes? are they different concentration of enzymes? if so then it looks ok to me since the higher the enzyme concentration from left to right, the higher the amount of digest products you have. If the peptide sizes obtained are not the ones expected then perhaps increase or decrease the digestion time since you could have a problem due to either incomplete digest or unspecific digestion, depending. I guess since BSA MW is ~ 66kDa your gel concentration is appropriate since you have a problem with the 15% gel as the products are too small and too abundant to resolve.


COmplete™ His-Tag Purification Column Protocol & Troubleshooting

cOmplete His-Tag Purification Columns (Product No. COHISC-RO) are compatible with automated chromatography systems such as ÄKTAexplorer.

Purification Under Native Conditions

Purification of native proteins should be performed using optimal buffer conditions for the target protein. Buffers recommended in this document are well established examples and can be adapted to achieve optimal conditions for a specific target protein.
cOmplete His-Tag Purification Columns offer flexibility in selecting the optimal buffer conditions.
Warning: The binding capacity may drop significantly if the buffer composition is suboptimal.
Note: For best results, load with a low flow rate to bind the target proteins more efficiently to the resin.
Warning: cOmplete His-Tag Purification Columns have been optimized using Buffer A and Buffer B specified in the table below. Other buffers might function as well, but need to be tested before use with cOmplete His-Tag Purification Columns.
Buffer A: 50 mM NaH2PO4, pH 8.0 300 mM NaCl
Buffer B: 50 mM NaH2PO4, pH 8.0 300 mM NaCl 250 mM imidazole
Note: The following protocol describes the experimental procedures when using an ÄKTAexplorer 100 System (Cytiva) for FPLC purification.

  1. Wash the pump with 10 to 20 mL Buffer A using the System Wash function of the ÄKTAexplorer System at a flow rate of 10 mL/min. Ensure that all air is displaced from the pumps and tubings of the system.
  2. Remove the plug at the column outlet and attach it to the outlet tubing of the ÄKTAexplorer System. Note: Save the plug of the column outlet in case the column needs to be stored or is planned to be reused.
  3. As soon as Buffer A is running out of the inlet tubing of the ÄKTAexplorer System, remove the upper plug from the column and immediately attach it to the inlet tubing of the ÄKTAexplorer System. Continuously measure OD280 values on the ÄKTAexplorer System. Note: If a fluorescent protein is purified, continuously measure the OD values at the absorption maximum of the fluorescence dye e.g., OD485 for GFP (Green Fluorescent Protein) or OD435 for CFP (Cyan Fluorescent Protein). Note: Save the plug of the column inlet in case the column needs to be stored or is planned to be reused.
  4. Define the flow rate as 10 mL/min for the 5 mL column or 2 mL/ min for the 1 mL column and equilibrate the column with 10 column volumes of Buffer A.
  5. Pause the run. Load the cleared sample (e.g., after an ultracentrifugation or filtration step) onto the column with a volumetric flow rate of 2.5 mL/min for the 5 mL column or 0.5 to 1 mL/min for the 1 mL column. Warning: To prevent blockage of the column, remove insoluble material before loading the column. Warning: Since the binding specificity of the resin is high, the kinetics of adhesion of the protein to the resin is slower than other available resins. If using high volumetric flow rates for loading, protein yield can decrease.
  6. Wash the column with Buffer A until the OD280 value reaches the baseline level (approximately 10 column volumes).
  7. Elute the His-tagged protein with a gradient of Buffer A (without imidazole) and Buffer B (250 mM imidazole). Warning: Protein peaks can be expected between 25 to 45 mM imidazole. Due to the specific characteristics of cOmplete His-Tag Purification Columns, a protein can already be eluted with approximately 25 mM imidazole. Warning: The amount of imidazole required in the elution buffer for efficient release of the target protein from the resin depends on various parameters, such as: • the length of the His-tag, • the accessibility of the His-tag.
  8. Wash and equilibrate for the next run. For details, refer to section Cleaning Procedures. Warning: If the column is not immediately reused, clean the column with 2 column volumes of 2 M imidazole to remove nonspecific binding of proteins. Equilibrate the column in a 20% ethanol solution and tightly close the column at both threads with plugs. Store at +2 to +8 °C to prevent cell growth.

Note:Refer to sections Purification Process Optimization and Troubleshooting for technical advice in optimizing the purification results.

Purification Under Denaturing Conditions

Purification of denatured proteins should be performed using optimal buffer conditions for the target protein. Buffers recommended in this document are well established examples and can be adapted to achieve optimal conditions for a specific target protein.
cOmplete His-Tag Purification Columns offer flexibility in selecting optimal buffer conditions.
Denature the protein or dissolve the inclusion bodies in a buffer containing 6 M guanidinium-HCl or 8 M urea.
Warning:The addition of urea to buffered solutions will cause the pH to drop. It is essential to adjust the pH of the buffer with NaOH after urea addition.
Warning: The binding capacity may also drop significantly if the buffer composition is suboptimal.
Note: For best results, load with a low flow rate to bind the target proteins more efficiently to the resin.
Warning: cOmplete His-Tag Purification Columns have been optimized using Buffer C, Buffer D, Buffer E, and Buffer F specified in the table below. Other buffers might function as well, but need to be tested before use with cOmplete His-Tag Purification Columns.
Buffer C: 100 mM NaH2PO4 10 mM Tris-HCl 8 M urea pH 8.0
Buffer D: 100 mM NaH2PO4 10 mM Tris-HCl 8 M urea pH 6.3
Buffer E: 100 mM NaH2PO4 10 mM Tris-HCl 8 M urea pH 5.9
Buffer F: 100 mM NaH2PO4 10 mM Tris-HCl 8 M urea pH 4.5
Note: The following protocol describes the experimental procedures when using an ÄKTAexplorer 100 System (Cytiva) for FPLC purification.

  1. Wash the pump with 10 to 20 mL Buffer C using the System Wash function of the ÄKTAexplorer System at a flow rate of , e.g., 10 mL/min. Ensure that all air is displaced from the pumps and tubings of the system.
  2. Remove the plug at the column outlet and attach it to the outlet tubing of the ÄKTAexplorer System. Note: Save the plug of the column outlet in case the column needs to be stored or is to be reused.
  3. As soon as Buffer C is running out of the inlet tubing of the ÄKTAexplorer System, remove the upper plug from the column and immediately attach it to the inlet tubing of the ÄKTAexplorer System. Continously measure OD280 values on the ÄKTAexplorer System. Note: If a fluorescent protein is purified, continously measure the OD values at the absorption maximum of the fluorescence dye e.g., OD485 for GFP (Green Fluorescent Protein or OD435 for CFP (Cyan Fluorescent Protein). Note: Save the plug of the column inlet in case the column needs to be stored or is to be reused.
  4. Define the flow rate as 10 mL/min for the 5 mL column or 2 mL/min for the 1 mL column and equilibrate the column with 10 column volumes of Buffer C.
  5. Pause the run. Load the cleared sample (e.g., after an ultracentrifugation or filtration step) onto the column with a volumetric flow rate of 2.5 mL/min for the 5 mL column or 0.5 to 1 mL/min for the 1 mL column. Warning: To prevent blockage of the column, remove insoluble material before loading the column. Warning: Since the binding specificity of the resin is high, the kinetics of adhesion of the protein to the resin is slower than other available resins. If using high volumetric flow rates for loading, protein yield can decrease.
  6. Wash the column with Buffer C until the OD280 value reaches the baseline level (approximately 10 column volumes).
  7. Wash with 10 to 20 column volumes of Buffer D.
  8. Wash with 10 to 20 column volumes of Buffer E.
  9. Elute the His-tagged protein with 10 to 20 column volumes of Buffer F. Note: Alternatively, the elution can also be performed with a gradient up to 200 mM imidazole solution using Buffer A and Buffer B instead of the pH shift option (refer to the elution step within section Purification Under Native Conditions).
  10. Wash and equilibrate for the next run under denaturing conditions with Buffer C or wash with Buffer A to remove the denaturing agents if the column will next be used under native conditions. For details, refer to section Cleaning Procedures. Warning: If the column is not immediately reused, clean the column with 2 column volumes of 2 M imidazole to remove nonspecific binding of proteins. Equilibrate the column in a 20% ethanol solution and tightly close the column at both threads with plugs. Store at +2 to +8 °C to prevent cell growth.

Note: Refer to section Purification Process Optimization and Troubleshooting for technical advice in optimizing the purification results.

Cleaning Procedures

cOmplete His-Tag Purification Columns can be used multiple times without loss of binding capacity. Over time, however, some protein aggregates might accumulate, leading to a decrease of efficiency of the resin within the columns. This can be identified by a slower flow rate or a higher back pressure.
The cleaning procedures remove aggregates for further efficient use of the columns. Different cleaning procedures can be carried out, based on the different applications. Once the cleaning procedure is completed, the resin should be transferred to 20% ethanol.

Stringent Native Cleaning

This method is appropriate when non-aggregating proteins have been purified, and if the column is used again for purifying the same protein.

  • Wash with 10 column volumes of 1 M imidazole/HCl, pH 7.5,
  • Wash with 10 column volumes of 4 M imidazole/HCl, pH 7.5,
  • Equilibrate the column with binding buffer and proceed to the next round of purification or transfer the material to 20% ethanol.

Denaturing Cleaning with SDS

This method is appropriate to remove aggregated proteins and lipids.
Warning: This cleaning procedure has to be performed at +15 to +25 °C because the solubility of SDS is more effective at this temperature than at +2 to +8 °C.
Note: The SDS buffer may also contain 50 mM DTT.
Warning:Avoid using K + in this buffer to prevent precipitation with SDS.

  • Wash with 10 column volumes of 1 M imidazole/HCl, pH 7.5,
  • Wash 2 times with 10 column volumes of 1 M imidazole/HCl, pH 7.5, 20% ethanol, 2 to 4% SDS,
  • Remove SDS with 3 times 10 column volumes of 20% ethanol.

Denaturing Cleaning with Guanidinium-HCl

This method is appropriate to remove aggregated proteins.
Note:The guanidinium-HCl buffer may also contain 50 mM DTT.

  • Wash with 10 column volumes of 1 M imidazole/HCl, pH 7.5,
  • Wash twice with 10 column volumes of 6 M guanidinium-HCl, 1 M imidazole, pH7.5,
  • Wash twice with 10 column volumes of 20% ethanol.

Note: In general, the choice of cleaning method depends on the protein type.
Note: The denaturing cleaning procedure with guanidinium-HCl presents fewer constraints than the denaturing cleaning method with SDS.

Purification Process Optimization

The parameters allowing for the maximal protein yield and purity might vary significantly depending on the characteristics of a given target protein. To optimize the protein purification procedure for highest protein purity, determine the optimal operating conditions for the specific target protein. Both purity and yield of a protein preparation depends on the sample amount. If the amount of sample is too high, the resin's binding capacity may not be sufficient to bind all target protein, and this will result in a suboptimal protein yield. If the amount of sample is too low, the remaining binding sites on the resin may enable background binding of lysate components. Optimal results are obtained when the amount of target protein matches the amount of resin within the columns. The capacity for a given target protein depends on several factors such as target protein size, conformation, multimerization status, length and accessibility of the His-tag, expression level and solubility of the His-tagged protein, lysate concentration, as well as the buffer pH and composition. For best results, determine the optimal ratio of the volume of lysate and resin within the columns required for the purification of a specific protein of interest, which is dependent on the expression rate of the protein:

  • Incubate the columns with varying volumes of lysates, in parallel test experiments,
  • Wash the columns and elute the bound proteins,
  • Determine the amount of target protein in the unbound fractions and in the eluate by SDS-PAGE,
  • The volume of lysate is optimal when only a small amount of target protein remains in the flow through and the maximal amount of protein is detected in the eluate fractions.

Note: The yield of the target protein can be optimized by allowing more time for the protein to bind to the resin. This can be performed by reducing the flow rate during the loading step of the chromatography purification.
Note: The optimal concentration of imidazole during binding, washing and elution steps can also be determined during pretrial experiments.
Note: Optimal results can typically be achieved with buffers containing a high salt concentration (300 mM) at pH 8.0 for target proteins compatible with those conditions.


Transfer of high molecular weight proteins - (Nov/14/2008 )

Hi
I am trying to probe for a high molecular weight complex but am not getting consistent detection. I use gradient gels and PVDF membrane for probing my 350kDa complex. Many times the background is immense though the bands are also present. Any suggestions please??

Make sure that it is running into gel and not stuck in the well.
-Gradient is good.
-Run for a longer time.
-Boil or not boil depending upon whatever works.

Make sure of complete transfer.
-PVDF is good.
-confirm by Staining the gel. PonceauS for membrane.
-Use thinner gel.
-Longer transfer time.
-May think of adding extra SDS to transfer buffer.
-Increased Methanol in transfer buffer may also help.

If transfer is complete, the background problem is a as good as any westerns.
-Refer to discussions on how to reduce background.

Blot cool (4 degrees C) and long (overnight -24 hours). It is best if you try several different times to see what works best for you.

I am working with a high molecular weight protein (MW 512kDa). For protein detection, I tried 5% SDS PAGE gel but the protein got stuck at the interface between the stacking and the running gel. I am using 1% SDS (TGS based) for running gel and 1% SDS and 20% methanol in the transfer buffer. If any one has experience working with such a high molecular weight proteins, please give me suggestions.

Seems kind of obvious but you don't have to use a stacker. They aren't absolutely necessary, especially for high MW proteins, though they do make small proteins run more evenly. I would also have said that 5% was a bit high for a 500 kDa protein, you may need to switch to some other gel system.

I would also lower the methanol concentration in your transfer, it reduces the mobility during transfer.

reduce the sds in your transfer buffer. more than 0.05% will interfere with binding.

the 20% methanol will help strip sds from the protein to aid binding, you can leave it alone if you want (you can extend transfer time, if necessary).

we used to run a 3.5-5% gradient gel for high mw proteins (with a 8M-0 urea gradient, to sharpen the bands).

Oh great, many huge protein fans here. I also need your help

I try to detect a 310 kDa protein. Up till now I used 6% resolving gel, 5% stacking gel and a Tris-glycine buffer system.
I stained the gel and it seems that there are proteins > 250 kDa on my gel (that's the largest protein of my marker). I didn't stain the PVDF membrane after transfer as I read this might interfere with detection afterwards (right? wrong?), but I will do it next time. I had problems with blocking and I got a very nice secondary antibody binding up to 250 kDa, but not beyond. So I think my main problem is the transfer.

You mentioned not using stacking gels - does this also apply if I can't use gradient gels? Would you suggest 5% or 6% gels? Or even less?

For the transfer I used a semi-dry blotting unit, but as I read that this might be suboptimal, I ordered a tank blot unit. Do you have any suggestions for a good transfer buffer? Is SDS necessary in the transfer buffer? Mine has just methanol, but no SDS.

And one question for blocking: We usually use 5% BSA in TBS-T. In the data sheet of my primary antibody they suggest 5% milk powder in TBS-T. How do I sterilze this solution? I was told not to autoclave it, but filtration was. a bit. ineffective

Oh great, many huge protein fans here. I also need your help

I try to detect a 310 kDa protein. Up till now I used 6% resolving gel, 5% stacking gel and a Tris-glycine buffer system.
I stained the gel and it seems that there are proteins > 250 kDa on my gel (that's the largest protein of my marker). I didn't stain the PVDF membrane after transfer as I read this might interfere with detection afterwards (right? wrong?), but I will do it next time. I had problems with blocking and I got a very nice secondary antibody binding up to 250 kDa, but not beyond. So I think my main problem is the transfer.

You mentioned not using stacking gels - does this also apply if I can't use gradient gels? Would you suggest 5% or 6% gels? Or even less?

For the transfer I used a semi-dry blotting unit, but as I read that this might be suboptimal, I ordered a tank blot unit. Do you have any suggestions for a good transfer buffer? Is SDS necessary in the transfer buffer? Mine has just methanol, but no SDS.

And one question for blocking: We usually use 5% BSA in TBS-T. In the data sheet of my primary antibody they suggest 5% milk powder in TBS-T. How do I sterilze this solution? I was told not to autoclave it, but filtration was. a bit. ineffective

if you start with 5-6% in separating gel you do not need a stacking gel if used it should be lower than <5%

you need a reference protein >/= 310 kDa

why using sterile milk solution? through it away after usage.

if you start with 5-6% in separating gel you do not need a stacking gel if used it should be lower than <5%

you need a reference protein >/= 310 kDa

why using sterile milk solution? through it away after usage.

I'm still waiting for the new blotting unit, but then I will try a 6% gel and a buffer with SDS.

Reference protein. hm. I just want to show size differences and the second form of the protein is in the range of my marker and I have a positive control (cell line).

I thought sterile milk solution might be better if there's anything inside the powder (fibres or whatever) that might interfere the blot. But you're right, I will just prepare it fresh, that's the easiest way.


Restriction Enzyme Troubleshooting Guide

The following guide can be used for troubleshooting restriction enzyme digestions. You may also be interested in reviewing additional tips for optimizing digestion reactions.

We are excited to announce that we are in the process of switching all reaction buffers to be BSA-free. Beginning April 2021, NEB will be switching our current BSA-containing reaction buffers (NEBuffer&trade 1.1, 2.1, 3.1 and CutSmart ® Buffer) to Recombinant Albumin (rAlbumin)-containing buffers (NEBuffer r1.1, r2.1, r3.1 and rCutSmart&trade Buffer). We anticipate that this switch may take as long as 6 months to complete. We feel that moving away from animal-containing products is a step in the right direction and are able to offer this enhancement at the same price. Find more details at www.neb.com/BSA-free.

During this transition period, you may receive product with BSA or rAlbumin-containing buffers. NEB has rigorously tested both and has not seen any difference in enzyme performance when using either buffer. Either buffer can be used with your enzyme. All website content will be switched in April to reflect the changes, although you may not receive the new buffer with your product immediately.

Videos

Restriction Enzyme Digestion Problem: DNA Smear on Agarose Gel

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Why is My Restriction Enzyme Not Cutting DNA?

Not getting the cleavage you expected? Let an NEB scientist help you troubleshoot your reaction.

Restriction Enzyme Digest Problem: Too Many DNA Bands

Are you finding unexpected bands in your digestion reaction? Here are some tips to help you determine the cause.

Restriction Enzyme Digest Protocol: Cutting Close to DNA End

When cutting close to the end of a DNA molecule, make sure you know how many bases to add to the ends of your PCR primers.

TIME-SAVER &trade Protocol for Restriction Enzyme Digests

Need a protocol to digest quickly and completely? Try this protocol for Time-Saver&trade qualified enzymes from NEB.

Reduce Star Activity with High-Fidelity Restriction Enzymes

NEB has engineered HF® enzymes to eliminate star activity. Learn how, and what this means for your digests.

Standard Protocol for Restriction Enzyme Digests

Let one of NEB's restriction enzyme experts help you improve your technique and avoid common mistakes in digest setup.

  • Check the methylation sensitivity of the enzyme(s) to determine if the enzyme is blocked by methylation of the recognition sequence
  • Use the recommended buffer supplied with the restriction enzyme
  • Clean up the DNA to remove any contaminants that may inhibit the enzyme
  • When digesting a PCR fragment, make sure to have at least 6 nucleotides between the recognition site and the end of the DNA molecule
  • Lower the number of units
  • Add SDS (0.1&ndash0.5%) to the loading buffer to dissociate the enzyme from the DNA or use Gel Loading Dye, Purple (6X) (NEB #B7024)
  • Use fresh, clean running buffer
  • Use a fresh agarose gel
  • Clean up the DNA (NEB #T1030)
  • Check the methylation sensitivity of the enzyme(s) to determine if the enzyme is blocked by methylation of the recognition sequence
  • DNA isolated from a bacterial source may be blocked by Dam and Dcm methylation
  • If the enzyme is inhibited by Dam or Dcm methylation, grow the plasmid in a dam-/dcm- strain (NEB #C2925)
  • DNA isolated from eukaryotic source may be blocked by CpG methylation
  • Enzymes that have low activity in salt-containing buffers (NEBuffer r3.1) may be salt sensitive, so clean up the DNA (NEB #T1030) prior to digestion
  • DNA purification procedures that use spin columns can result in high salt levels, which inhibit enzyme activity. 1 To prevent this, DNA solution should be no more than 25% of total reaction volume.
  • Clean up the PCR fragment prior to restriction digest (NEB #T1030)
  • Use the recommended buffer supplied with the restriction enzyme
  • Use at least 3&ndash5 units of enzyme per &mug of DNA
  • Increase the incubation time
  • Some enzymes have a lower activity on supercolied DNA. Increase the number of enzyme units in the reaction.
  • Some enzymes can exhibit slower cleavage towards specific sites. Increase the incubation time, 1&ndash2 hours is typically sufficient.
  • Some enzymes require the presence of two recognition sites to cut efficiently
  • Assay substrate DNA in the presence of a control DNA. Control DNA will not cleave if there is an inhibitor present. Mini prep DNA is particularly susceptible to contaminants.
  • Clean DNA with a spin column (NEB #T1030) or increase volume to dilute contaminant
  • Lower the number of units in the reaction
  • Add SDS (0.1&ndash0.5%) to the loading buffer to dissociate the enzyme from the substrate
  • Use the recommended buffer supplied with the restriction enzyme
  • Decrease the number of enzyme units in the reaction
  • Make sure the amount of enzyme added does not exceed 10% of the total reaction volume. This ensures that the total glycerol concentration does not exceed 5% v/v
  • Decrease the incubation time. Using the minimum reaction time required for complete digestion will help prevent star activity.
  • Try using a High-Fidelity (HF) restriction enzyme. HF enzymes have been engineered for reduced star activity.
  • Enzymes that have low activity in salt-containing buffers (e.g., NEBuffer r3.1) may be salt sensitive. Make sure to clean up the DNA (NEB #T1030) prior to digestion.
  • DNA purification procedures that use spin columns can result in high salt levels, which inhibit enzyme activity. 1 To prevent this, DNA solution should be no more than 25% of total reaction volume.
  • Clean-up the PCR fragment prior to restriction digest (NEB #T1030)
  • Use the recommended buffer supplied with the restriction enzyme
  • Use at least 5&ndash10 units of enzyme per &mug of DNA
  • Digest the DNA for 1&ndash2 hours


1 Monarch Kits (NEB #T1010, NEB #T1020, and NEB #T1030) use columns that have been designed to minimize salt carry over into the eluted DNA, so using them can minimize this issue.


Quantitation

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.


Sample denaturation

Various sample buffers have been used for SDS-PAGE but all use the same principles to denature samples. We obtain good denaturation by preparing a sample to a final concentration of 2 mg/ml protein with 1% SDS, 10% glycerol, 10 mM Tris-Cl, pH 6.8, 1 mM ethylene diamine tetraacetic acid (EDTA), a reducing agent such as dithiothreitol (DTT) or 2-mercaptoethanol, and a pinch of bromophenol blue to serve as a tracking dye (

We prepare a 2x concentrate of sample buffer consisting of 2% SDS, 20% glycerol, 20 mM Tris-Cl, pH 6.8, 2 mM ethylene diamine tetraacetic acid (EDTA), 160 mM dithiothreitol (DTT), and 0.1 mg/ml bromphenol blue dye. I prefer DTT to 2-mercaptoethanol because the latter has a much stronger unpleasant odor and it doesn't denature our blood fractions very well. Part of the problem is that our water baths don't reach the boiling point, and boiling may be necessary with 2-mercaptoethanol. We prepare all of our unknowns to the same concentration then mix 1 volume prepared sample to 1 volume 2x buffer.

So, what do the various components do? EDTA is a preservative that chelates divalent cations, which reduces the activity of proteolytic enzymes that require calcium and magnesium ions as cofactors. The tris acts as a buffer, which is very important since the stacking process in discontinuous electrophoresis requires a specific pH. Glycerol makes the sample more dense than the sample buffer, so the sample will remain in the bottom of a well rather than float out. The dye allows the investigator to track the progress of the electrophoresis.

SDS, DTT, and heat are responsible for the actual denaturation of the sample. SDS breaks up the two- and three-dimensional structure of the proteins by adding negative charge to the amino acids. Since like charges repel, the proteins are more-or-less straightened out, immediately rendering them functionless. Some quaternary structure may remain due to disulfide bonding (covalent) and due to covalent and noncovalent linkages to other types molecules. By the way, another name for SDS is lauryl sulfate. Your shampoo may contain lauryl sulfate - now doesn't that inspire confidence in the product?

Many proteins have significant hydrophobic properties and may be tighly associated with other molecules, such as lipids, through hydrophobic interaction. Heating the samples to at least 60 degrees C shakes up the molecules, allowing SDS to bind in the hydrophobic regions and complete the denaturation.

The amino acid cysteine contains a sulfhydryl (-SH) group that spontaneously forms a disulfide bond (-S-S-) with another sulfhydryl group under normal intracellular conditions. Disulfide bonding is covalent and is not disrupted by SDS. DTT is a strong reducing agent. Its specific role in sample denaturation is to remove the last bit of tertiary and quaternary structure by reducing disulfide bonds.

Most sample buffers do not remove covalently attached carbohydrate or phosphate groups, and some associations with other types macromolecules are difficult to disrupt. Polypeptides contain varying amounts of basic and acidic amino acids that add charge to the molecules, and individual amino acids vary in molecular weight although they may bind SDS with the same affinity. Therefore, charge to mass ratio and the relative mobility of many proteins is affected by factors other than strictly the molecular weight. SDS-PAGE is very effective in providing reproducible results, but don't count on precise values for MW determination.


6 Important FAQs About Bovine Serum Albumin (BSA)

Bovine Serum Albumin (BSA) is universal in the lab, having use in Western blot, cell tissue culture, PCR and more, but BSA’s versatility has led researchers to ask many common questions. Rather than chasing answers through the Internet, we have identified 6 very common questions regarding BSA and provided some very detailed answers to serve as a helpful guide.

1. What Type of BSA Should I Use for My Experiment?

In our earlier article on this very topic we actually addressed this question in greater detail. This question comes up often because there are so many types of BSAs available which are directed toward a wide range of laboratory applications, each having characteristics that better optimize it for a lab technique. For instance, you might want an IgG free BSA if you need a blocking agent.

Deciding on which BSA to use will be based on the type experiment you’re doing and your specific needs. After taking a look at the above mentioned article and its selection guide table, it’s still good to refer to other sources such as ResearchGate for more information and a more tailored answer.

2. How Does BSA Work as a Blocking Agent?

Blocking agents are meant to prevent the nonspecific binding of your antibodies. In general, as long as a protein doesn’t have affinity for your probe, it can theoretically be used as a blocking agent. But if you choose anything at random, your experiment will likely be inefficient. Therefore, there are certain proteins that are directed for this type of work. These proteins bind more consistently to the membrane, improving assay sensitivity and reducing background interference.

So why BSA then? First of all, you don’t want just any BSA. Many types of BSAs contain IgGs (ok for other applications) which can lead to nonspecific binding. The goal is to work with a protein that will keep your experiment in optimal conditions, so in the case of a blocking agent, particularly BSA, you want to consider types that are free of immunoglobulins.

3. Should I Use BSA or Milk for My Western Blot?

The two most frequently used blocking agents in labs are nonfat milk and BSA. And there are pros and cons to each. Milk is usually more affordable and easily prepared from powder compared to BSA however, milk is not good to use in avidin-biotin systems since milk contains biotin, and milk should not be used on phosphorylated proteins.

Typically, when working with phosphorylated proteins, BSA tends to work better as a blocking agent. This is because milk has a variety of proteins, one being phosphoprotein casein, which leads to a higher background. Of course with all advice, there are special cases. The best rule of thumb when working with phosphorylated proteins is to start with BSA, and then optimize from there.

4. Why Does Protein Concentration Estimation Usually Require BSA?

In order to estimate the protein concentration of an unknown sample, you need to compare it to a reference that is most similar to your sample. However, this isn’t always easy to find, so BSA plays the role of reference. There are a few factors that make BSA an appropriate reference to use: it’s very abundant, it’s very affordable, it’s very stable and it won’t have a huge impact in biochemical reactions.

5. Is Bovine Serum Albumin (BSA) and Fetal Bovine Serum (FBS) the Same Thing?

Nope. It’s not the same. Fetal Bovine Serum is a commonly used serum supplement for eukaryotic cell culture. The benefit of FBS for cell culture is its lower antibody levels and higher growth factor levels. Bovine Serum Albumin (BSA) is used in a variety of laboratory applications including its function as a protein concentration standard, its function as a cell nutrient and its ability to stabilize enzymes during restriction digest. The important takeaway here, however, is that while they often both come up in Google searches when you’re only looking for one or the other, and sound very similar, they are indeed different.

6. Why is the It Important for BSA to Be Manufactured in the USA?

The importance of sourcing your BSA from either the US or Australia is due to concern about the prion infection bovine spongiform encephalitis (BSE). There has not been an outbreak in the United States which makes these products more trustworthy. GoldBio particularly stresses that their BSA is made in the United States in a closed loop system from USDA inspected animals, and is certified to be BSE/TSE free.


2-D Electrophoresis Workflow

The 2-D Electrophoresis Workflow provides a comprehensive selection of product solutions and educational resources that give you better results for biomarker discovery, protein characterization, and HCP assay development.

Sample Preparation

1st Dimension Electrophoresis

2nd Dimension Electrophoresis

Staining, Imaging, and Analysis

2-D Electrophoresis Equipment and Reagents

Complete systems for 2-D electrophoresis including first-dimension isoelectric focusing (IEF) cells, "2DE" buffers and reagents, precast gels, and mini, midi, and large format second-dimension electrophoresis cells.

Preparative Electrophoresis

Bio-Rad&rsquos preparative electrophoresis systems can fractionate and purify nanogram to gram quantities of proteins or nucleic acids by electrophoresis or continuous electroelution from gels.


Relative activity of restriction enzymes in universal and basal buffers

Our restriction enzymes are supplied with an optimal universal buffer (one of five universal buffers indicated in blue in the table below). The relative activity in each of the other universal buffers is normalized to the optimal buffer, where the activity of each enzyme in the optimal buffer is expressed as 100%. Values in ( ) indicate buffers that are likely to be affected by star activity. In order to avoid these effects, use of buffers highlighted in blue or pink is recommended.

A few specific enzymes (AccIII, BalI, BcnI, BglI, Bpu1102I, Cfr10I, Eco52I, NruI, PshBI, SnaBI, SspI, TaqI, and VpaK11BI), are each supplied with a basal buffer specialized for the particular enzyme. The compositions of these basal buffers vary depending on the enzyme.

Restriction enzyme Relative activities (%)
LMHKT + BSABasal***
AatII <20 <20 <20 <20 100 120
AccI 20 100 <20 (<20) 160 80
AccII (260) 100 <20 20 200 160
AccIII (<20) (<20) 20 (80) (<20) 100
AfaI 60 60 40 60 100 100
AflII 20 80* <20 <20 140 120
AluI 100 100 <20 40 200 120
Aor13HI <20 20 <20 80 * 80 100
Aor51HI 80 100 <20 20 120 120
ApaI 100 <20 <20 <20 <20 120
ApaLI 100 20 <20 <20 120 120
AvaI (<20) 100 20 40 100 120
AvaII 80 100 <20 20 100 100
BalI 20 20 <20 <20 40 100
BamHI (<20) <20 40 100 (<20) 80
BanII (120) (120) 100 80 (100) 100
BcnI <20 20 40 60 60 100
BglI <20 <20 20 40 <20 100
BglII <20 20 100 (100) (60) 100
BlnI <20 20 40 100 20 120
BmeT110I <20 <20 20 100 <20 140
BmgT120I <20 <20 100 40 <20 240
Bpu1102I <20 <20 <20 40 60 100
BspT104I 100 60 <20 <20 100 120
BspT107I <20 20 80 100 20 100
Bsp1286I 100 20 <20 <20 60 100
Bsp1407I 20 60 20 20 100 100
BssHII 100 100 60 20 140 100
BstPI (<20) (60) 100 (100) (100) 100
BstXI <20 40 100 <20 <20 120
Bst1107I (<20) 60 100 100 40 100
Cfr10I (<20) (<20) (<20) 40 (20) 100
ClaI 40 100 120 100 60 100
CpoI <20 <20 80 100 <20 100
DraI 100 100 60 100 80 80
EaeI 60 100 <20 <20 120 160
EcoO65I (20) (60) 60 * 40 40 100
EcoO109I 100 60 <20 <20 100 160
EcoRI (20) (100) 100 (120) (80) 120
EcoRV (<20) (40) 100 (120) (40) 100
EcoT14I (<20) (40) 100 120 (60) 100
EcoT22I <20 20 100 (140) (20) 120
Eco52I <20 <20 <20 <20 <20 100
Eco81I <20 100 <20 <20 100 160
FbaI (<20) (<20) (80) 100 (20) 100
FokI (20) (60) <20 <20 (200) 100
HaeII 80 100 <20 80 140 100
HaeIII 60 100 100 60 100 100
HapII 100 60 <20 <20 100 80
HhaI 80 100 100 120 120 100
HincII 20 100 20 40 100 80
HindIII (60) 100 <20 200 (100) 80
HinfI 80 100 100 160 60 100
Hin1I 40 80* <20 20 60 160
HpaI <20 (40) 20 100 (80) 100
KpnI 100 60 <20 <20 (100) 80
MboI 20 40 60 100 40 100
MboII 100 60 <20 <20 60 100
MflI 100 80 <20 <20 80 100
MluI 60 60 100 (100) 60 100
MspI 80 80 <20 100 100 80
MunI (200) 100* <20 <20 160 100
MvaI (<20) (40) 80 100 (20) 120
NaeI 100 <20 <20 <20 100 120
NcoI (40) (60) 20 60* (60) 160
NdeI <20 40 100 100 80 100
NheI (120) 100 <20 <20 (160) 100
NotI (<20) (<20) 20** <20 (<20) 100
NruI 0 <20 20 20 <20 100
NsbI 40 20 <20 60 100 100
PmaCI 100 80 <20 <20 100 100
PshAI 20 40 <20 100 60 160
PshBI (20) (40) 20 40 40 100
Psp1406I 20 60 <20 <20 100 100
PstI (<20) (60) 100 80 (20) 80
PvuI (<20) (20) (40) 80* (40) 120
PvuII (80) 100 40 <20 (40) 100
SacI 100 60 <20 <20 80 80
SacII 40 20 <20 <20 100 40
SalI <20 <20 100 (20) <20 120
Sau3AI (60) 80 100 <20 (80) 100
ScaI (<20) (<20) 100 (60) (<20) 100
SfiI (40) 100 <20 <20 100 100
SmaI <20 <20 <20 <20 100 100
SmiI <10 <20 100 40 <10 100
SnaBI (20) (40) <20 <20 (40) 100
SpeI (80) 100 80 100 (80) 100
SphI (20) (40) 100 120 (20) 100
Sse8387I (120) 60* <20 <20 (60) 100
SspI (<20) (60) 40 (100) (80) 100
StuI 60 100 60 80 140 100
TaqI 40 80 60 60 80 100
Tth111I (20) 80 40 100 (80) 120
Van91I <20 (20) 60 100 (60) 100
VpaK11BI <20 <20 60 (40) <20 100
XbaI <20 80* 20 <20 120 120
XhoI <20 60 100 160 60 100
XspI <20 60 <20 100 160 100

Blue: buffer supplied with the restriction enzyme
Pink: alternative buffer recommended for use

*+0.01% BSA &rarr 100% AflII, EcoO65I, FokI, Hin1I, MunI, NcoI, PvuI, SplI, Sse8387I, XbaI
**+0.01% BSA + 01% Triton X-100 &rarr 100% NotI
*** The compositions of the basal buffers are enzyme-specific.


Troubleshooting SDS-PAGE of trypsin-treated BSA - Biology

on a mission to make biochemistry fun and accessible to all

Could that be a massive protein I see? We can separate proteins by size by sending them traveling through an SDS-PAGE gel, but in order to find our protein treasure we need to send in some treasure hunters. And the one we use most often is a dye called Coomassie Brilliant Blue (CBB), either in the classical way or in the form of faster colloidal stains (often advertised as some trademarked version of &ldquoinstant blue&rdquo).

note: this post is refurbished from September 2019, about 300 SDS-PAGEs ago (literally, not exaggerating &ndash today I ran my 732nd, 733rd, & 734th!)

SDS-PAGE (Sodium Dodecyl Sulfate &ndash PolyAcrylamide Gel Electrophoresis) is a technique we use to separate proteins in a mixture by size (well, length) using electricity to send the proteins through a gel mesh. The goal here isn&rsquot to purify the proteins, instead the goal is just to see what&rsquos there. But protein&rsquos are invisible to the naked eye, so we need some sort of dye in order to protein-spy.

note: the blue you see while the gel is running is just a tracker dye in the sample loading buffer which helps you know when to turn off the electricity and start the staining.

You can think of the SDS-PAGE gel&rsquos matrix as maze with protein &ldquotreasure&rdquo hidden throughout. Unlike the proteins, with moved straight down the gel because the electric field was compelling them that way, the dyeing step happens after you turn off the electricity and transfer your gel from its plate sandwich to a staining box. Here, there&rsquos no directionality to movement of things, so the dye will move wherever the heck it wants! (though the protein will be stuck in place through chemical-induced clumping as we&rsquoll see).

The dye travels randomly through the gel maze by diffusion (the molecules move around randomly, ricocheting off the things they run into with the NET RESULT that they move from areas of high concentration to areas of low concentration). When it finds protein treasure, it latches on. And since the dye&rsquos blue it tells us where the protein is.

But it&rsquos really hard to find treasure that&rsquos trying to escape! So we need to get the treasure to stay put &ndash the FIXATION step uses an alcohol and/or acid to make the protein to precipitate (clump up) so it gets stuck in place so our treasure-hunting dye can find it. We also have to watch out for overly-eager treasure hunters that latch onto &ldquofool&rsquos gold&rdquo (stain the gel itself) causing high background. This leads to an overall treasure-hunting scheme of:

  1. GEL ELECTROPHORESIS &ndash separate proteins by size by unfolding them, coating them in negatively-charged detergent (SDS) & using that negative charge to motivate them to travel through a polyacrylamide gel mesh towards a positive charge. The bigger (longer) proteins get tangled up more, so they travel slower and have progressed less (so higher up on gel) when you turn off the power. More here: http://bit.ly/2GZc3tG
  2. WASH &ndash remove free SDS, etc.
  3. FIX &ndash trap the treasure &ndash the gel has to allow proteins to move when you want them to, but not too easily &ndash Ideally they&rsquod only move when power&rsquos on, but molecules like to move and if they can they will &ndash so (especially the small ones) can start wandering off even when the power&rsquos off). To prevent that wandering, you add an alcohol and/or acid to get them to clump and get stuck in place.
  4. STAIN &ndash send in the treasure hunters &ndash stick the gel in a bath of dye &ndash the dye enters, latches onto the protein and gets stuck too (this step&rsquos often combined w/the fixing step)
  5. DESTAIN &ndash call off the hunt &ndash get the treasure-less treasure hunters to leave so you can better see where the treasure-full ones are (some rapid stains have low backgrounds & you just destain in water, letting the dye diffuse out, but some methods use more complex destaining)

As I alluded to, there are lots of different formulations including &ldquoClassic Coomassie&rdquo which has really eager treasure-hunters that can find tiny amounts of treasure but are also fool&rsquos-gold-happy &ndash it&rsquos super sensitive and eventually gives you nice crisp bands, but takes a lot of destaining to reveal them. Alternative &ldquoColloidal Coomassie&rdquo recipes are becoming more and more common because they&rsquore faster & more eco-friendly &ndash these forms keep groups of treasure-hunters hanging out outside the gel and gradually send them in until all the treasure&rsquos found and bound.

Let&rsquos take a closer look at our treasure hunter, Coomassie Brilliant Blue (CBB). Most scientific reagents have boring (though descriptive) names, so you might wonder how Coomassie Brilliant Blue (CBB) got its name. If you&rsquore kinda nerdy, you might look it up. And if you&rsquore even nerdier like I am, (embrace it!) you might then tell other people about it!

&ldquoCoomassie&rdquo is actually a trademark name &ndash owned by a company that no longer even makes it &ndash & a town name (modern-day Kumasi, Ghana). CBB wasn&rsquot discovered or produced there, nope &ndash a British company thought naming their product after the capital of the Ashanti empire they recently conquered would be good business strategy. That makes me mad, so I&rsquom going to call it CBB most of the time.

&ldquoCoomassie&rdquo was initially used to market a wide range of wool dyes, with CBB first made in 1913 & first used to stain proteins in 1960s. Dye-ing to know more about the dye itself? It has a pinwheel-like chemical structure and is characterized as a triphenylmethane dye. Unlike the common name a company decided to give a product, this is an example of a functional name &ndash it describes the chemical makeup &ndash in this case three (tri) phenylmethane groups &ndash phenyl is a type of resonance-stabilized ring (a ring or atoms that share electrons in a delocalized fashion that makes them good at absorbing light and thus making things look colored) &ndash and methyl is a -CH₃ group.

There are 2 main forms of this &ldquotriphenylmethane&rdquo dye, R-250 & G-250. Both are blue, but R&rsquos more &ldquoreddish&rdquo & G&rsquos more &ldquogreenish&rdquo (although the color depends on the pH and whether and what it&rsquos bound to which is why it can be used to measure protein concentrations in something called a &ldquoBradford assay&rdquo http://bit.ly/bradforduv ). &ldquo250&rdquo was originally a purity/strength indicator. R-250 is more sensitive , but G-250 can be made into forms that produce lower background, with faster protocols.

We get the G-250-based &ldquoquick stain&rdquo we use most of the time pre-made, but we make our own &ldquoClassic&rdquo R-250 stain. It&rsquos a really simple recipe &ndash only 4 ingredients: water, acetic acid (AcOH), methanol (MeOH), & CBB &ndash but it takes a while to prepare&hellip more here: http://bit.ly/2QJNwLy

But none of that really matters if the dye&rsquos not where you want it &ndash and only where you want it. Where we want it is stuck to our protein (which is itself stuck in our gel). And where we don&rsquot want it is anywhere in the gel there isn&rsquot any protein. So how does it stick to our protein but not the gel? The bumbling biochemist&rsquos here to tell!

CBB has sulfuric acid groups that can be negative or neutral depending on pH. Under the conditions of the staining solution it has overall ➖charge (anionic), so it binds (reversibly) to ➕-charged parts of proteins (basic amino acids like Arg, Lys, & His) through electrostatic interactions (opposites attracting). note: those side chains aren&rsquot always positive, but we stain the gel in acidic conditions, where they&rsquore more likely to be &ndash more here: http://bit.ly/30qzHH6

CBB also binds to non-charged protein parts (especially the ring-y (aka aromatic) amino acids like Phe, Tyr, & Trp) &ldquogenerically&rdquo through &ldquoVan der waals&rdquo interactions, which involve shifting around of electrons when molecules get close together. These interactions are individually weak but they add up (they&rsquore what allow geckos to walk up walls!). Proteins with unusually high proportions of ring-y amino acids tend to stain better. An example is BSA (bovine serum albumin), which recruits twice as many treasure hunters per weight of protein than your average protein.

Speaking of weight, different proteins have different weights because they have different #s of amino acid letters. We commonly talk about protein weights in terms of &ldquomolecular weight (MW)&rdquo and units of &ldquokiloDaltons&rdquo (kDa). BSA&rsquos molecular weight is 66.5 kDa, meaning that 1 mole (6×10^23) of BSA molecules would weigh 66.5 kg. More on moles here: http://bit.ly/2KQLw4k

But the key thing here is that bigger proteins have &ldquomore to love&rdquo in CBB&rsquos eyes &ndash they offer more binding sites and thus will more BSA per protein molecule than a smaller protein. So your bands would look darker for bigger proteins than smaller proteins if you ran the same # of protein molecules of each.

CBB&rsquos protein-binding ability also makes it a good wool dye because wool&rsquos chock full of a protein called keratin. And that keratin-binding ability which makes it good at dyeing wool also makes it good at dying your skin &ndash so wear gloves & avoid splashing. CBB can also bind SDS (that detergent we used earlier to solubilize and negatively-charge the proteins) which can mess up results. So you want to wash your gel in water before staining to remove the SDS.

The classical CBB method goes something like this (apologies for scribbled-note-like formatting)&hellip

  1. water wash -> get buffer components to diffuse out of gel -> start with a &ldquoclean slate&rdquo
  2. fixation/stain -> immerse gel in a staining solution (CBB + methanol (MeOH) + acetic acid (AcOH)) (some more eco-friendly methods use ethanol)
    • we dissolve CBB in a MeOH/AcOH solution to combine the fixation & staining steps. note: It takes a really, really long time for the CBB powder to dissolve, so let it stir for several hours on a magnetic stir plate and then filter through a coffee filter to remove any undissolved stuff
    • fixation: protein precipitates so is trapped in the gel matrix
    • stain: dye diffuses into gel
      • when dye meets protein -> dye binds -> gets trapped because the protein&rsquos trapped
      • where there&rsquos not any protein, the dye still fills the pores of the gel, but it&rsquos not trapped (it can flow in and out but at this point its flowing mainly in because the dye concentration ([dye]is higher in the stain than in the gel).
      • When [dye outside gel] = [dye inside gel] you&rsquove reached &ldquoequilibrium.&rdquo Equilibrium isn&rsquot a stop point &ndash it&rsquos actually a dynamic process (the individual dye molecules are still moving around but the rate of molecules entering the pores = rate exiting the pores, so you can&rsquot tell anything&rsquos changing.
  3. destain -> swap out the stain solution with a destain solution (MeOH + AcOH but NO CBB) -> remove the non-protein-bound CBB
    1. the stain can&rsquot go down the drain &ndash instead we collect it in hazardous waste containers that our environmental health & safety (EH & S) workers dispose of properly
    2. at the start, there&rsquos high CBB in gel, NO CBB in bath -> CBB diffuses out of gel into bath
    3. this makes CBB concentration in bath increase -> less of a difference between gel & bath -> CBB diffuses more slowly

    As you can hopefully see from the pics, Classic CBB initially stains your whole gel blue so you can&rsquot see the bands until you destain it and de-blue what&rsquos not supposed to be blue. This is because in order to find tiny treasure you unleash a ton of treasure hunters that race in from the dye bath, where there&rsquos a high concentration of dye, to the pores of the gel where there&rsquos more room &ndash and hopefully protein! But they&rsquoll keep snooping around even if there&rsquos no protein left to find because you have high concentrations of free dye inside and outside the gel.

    But Colloidal CBB doesn&rsquot require as much destaining (though it helps make the bands crisper) because, instead of sending in a ton of treasure hunters you then have to kick out, it has groups of hunters hang out outside the gel and only go in &ldquoas needed&rdquo

    A colloid is a solution where instead of having individual molecules spread throughout, you have &ldquoclumps&rdquo of those molecules, but still evenly spread out like in a &ldquonormal&rdquo solution. And those clumps really are &ldquodissolved&rdquo (they don&rsquot come &ldquoundone&rdquo when you let the solution sit).

    The colloids are too big to enter the gel so you don&rsquot &ldquooverstain.&rdquo BUT the FREE molecules can come & go from the gel as they please &ndash and they do, just as before, except now they&rsquore at a lower concentration. So how do you get enough to stain the protein? No problem &ndash the free CBB population can constantly get replenished from colloidal &ldquostock rooms&rdquo (you have a sort of equilibrium between free & colloidal).

    The result? Instead of flooding the gel with dye (like the &ldquoclassical&rdquo method) you supply a steady, mild flow of CBB molecules &ndash enough to quench the proteins&rsquo thirst, but not so much that it can stick to the gel.

    Thus, you can see the bands even without destaining. It also helps that, although we think of CBB as blue, it&rsquos color depends on its charge. At low pH, CBB G-250&rsquos brownish, but when it binds proteins, it gets stabilized in blue state so you can tell background brown from treasure blue. http://bit.ly/bradforduv

    At least, the stain we used to get was more brown-y. That stain&rsquos been on back-order for months. And months. And months. So we&rsquove been experimenting with a few other formulations but they seem to start bluer and give higher background.

    Speaking of color, a great thing about coomassie stains is that, unlike the DNA gels that use fluorescent dyes we have to stick on a UV tray, http://bit.ly/2zjHH0K. CBB is &ldquocolorimetric&rdquo &ndash the readout is color, so the results are literally right there to see, no special equipment required (unless you count the eye, which is a pretty spectacular tool!) (and the light tray helps too 🙂 ).

    Another colorimetric protein stain is silver staining, which is more sensitive, but also more finicky. http://bit.ly/2JGHuto

    There are also fluorescent protein stains. CBB & silver stain are both &ldquoall protein stains&rdquo &ndash they stain all proteins (though sometimes not equally well). There are also fluorescent all protein stains. But the really cool fluorescent stains only stain specifically-modified proteins like glycoproteins (which have sugar chains added) or phosphoproteins (which have phosphate groups added)

    some sources on CBB history and use:

    more on topics mentioned (& others) #365DaysOfScience All (with topics listed) 👉 http://bit.ly/2OllAB0

    #scicomm #biochemistry #molecularbiology #biology #sciencelife #science #realtimechem


    Watch the video: SDS PAGE Introductory Procedures (May 2022).