Epoxy beads
- Please tell me how to separate FG beads (magnetic separation and centrifugation).
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In order to reduce the background (reduce non-specific adsorption), the following cautions are required in the experiments.
Immobilization
Perform centrifugal separation.
The nano size of FG beads gives them excellent dispersibility. As a result, magnetic separation in an organic solvent may be difficult, and it is necessary to recover the FG beads by centrifugal separation.
During affinity purification (screening)
Perform magnetic separation
When centrifugal separation is performed, heavy proteins and insoluble proteins are also precipitated, raising the level of the background. Because FG beads have high dispersibility, magnetic separation requires time (in some cases 5 minutes or longer). However using magnetic separation avoids the risk of intrusion by these impurities and provides clear results with a low background level.
- Please tell me how to disperse FG beads (ultrasonic method and manual method).
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In order to reduce the background (reduce non-specific adsorption), the following cautions are required in the experiments.
Immobilization
Disperse the beads well.
Centrifugal separation causes the FG beads to agglutinate strongly, making it difficult to disperse them. Ordinarily a manual dispersion method or ultrasound would be used to disperse the beads. Although ultrasound separates the beads easily, caution is required due to the possibility of damaging the proteins. Therefore in general it is recommended that ultrasound be used when binding low molecular weight compounds, and that manual dispersion be used when binding proteins. The manual method involves resting the bottom of the micro tube in a plastic test tube stand and moving it roughly to disperse the beads. Depending on the type of micro tube, when using the manual method to disperse the beads, the bottom of the tube may crack or leakage from the lid may occur. Use a micro tube that is strong and has a lid with a tight fit. We recommend the use of cap locks.
During affinity purification (screening)
Disperse the beads well.
If dispersion is insufficient following the FG beads washing process after the binding reaction with the proteins, there is the possibility of impurities remaining inside the bead clusters. Therefore it is necessary to disperse the beads well. Use the manual method to disperse the beads. (With ultrasound, there is the risk that the proteins will be damaged.)
- I mistakenly frozen some beads that were supposed to be stored in the refrigerator. Is it available?
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We do not recommend using the beads after freezing, as crystallization of water molecules may compromise the structure of the beads.
- What amount of the beads is required?
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20 mg of beads are sufficient for 2 studies of the affinity purification conditions. A total of approximately 50 mg of beads are necessary in order to secure a certain amount of the purified target proteins or other substance after the study of the conditions.
- What are the important points when designing a ligand?
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When designing a ligand in order to introduce a functional group for the parent compound of the protein you seek to identify, because the protein which will be purified varies depending on the location where the function group is introduced, it is recommended that receptor identification tests be performed by binding multiple ligands at different linker binding locations on the same parent compound. In the event that there is no functional group capable of binding to the ligand, it is necessary to change the chemical structure of the compound itself.
- How are beads stored after the ligands are bound to them?
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When the amount of bound ligands is large, the beads become hydrophobic and dispersibility decreases. Therefore the beads are stored in a 50% methanol solution, not in water.
- Are there any methods other than HPLC for verifying whether or not ligand binding has been successful?
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The proteins can be actually bound to the ligands in order to verify binding. At a binding concentration of 0 mM, there is no protein binding. If the protein band increases as the binding concentration increases, then good binding has been verified.
- How strong is the affinity for the proteins that are affinity purified?
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Ordinarily the affinity for the ligands of the proteins to be affinity purified is expressed as a dissociation constant (Kd) of 10-6 M or below. If the affinity is too low, there is the risk of a higher level of background noise.
- What is the purification efficiency?
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When ligands with a relatively strong bonding strength are used, the results of studies by our company show a purification efficiency of approximately 50%. However the actual efficiency is highly dependent on the properties of the individual ligands and varies on a case-by-case basis.
- How is the cell extract prepared?
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We recommend the Dignam method. However, since the Dignam method requires a large amount of cells, we recommend a method of solubilizing with a surfactant such as NP-40 when using a small amount. Please refer to the protocols 401 and 402 from the protocol page. If the above is difficult, you can also use commercially available kits (ProteoExtract Subcellular Proteome Extraction Kit (MERCK MILLIPORE), CelLytic M (SIGMA), etc.). However, if the detergent concentration is 1% or higher, it may prevent the ligand from binding to the target protein during affinity purification, so reduce the detergent concentration to 0.1% by dialysis or dilution before use.
- Is there any problem with using frozen stock homogenate?
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There is no problem. However perform centrifuge separation before using to remove any impurities (such as degenerated proteins).
- How much protein supply is necessary?
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Serum proteins or extracts from cultivated cells or tissues can be used. When using cultivated cells as the protein source, 109 nuclear proteins or 107 – 109 cellular proteins are required.
- Can affinity purification be used with membrane proteins such as GPCRs and ion channels?
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While it is not impossible, it is not easy to use affinity purification by this method for GPCRs and ion channels. There are cases when affinity purification can be used by solubilizing the membrane protein with a surfactant.
- There are many background bands. how can i reduce it?
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It is necessary to consider the optimal conditions by varying the concentration of ligand binding to the FG beads, the salt concentration in the buffer, and the surfactant concentration. This can sometimes be improved by carefully performing dispersion when screening. It is also important to centrifuge the protein solution before use to remove the insoluble fraction. Insufficient masking is also a possibility, so it is necessary to suitably mask the functional groups which are not bound by the ligands.
- What should be done when a large number of bound protein bands are detected?
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Change the buffer composition and detect a more highly specific band. Alternatively, it is necessary to perform competitive inhibition tests and drug elution, and verify whether or not the band is specific to that ligand. In addition, the use of active or inactive ligand-immobilized beads makes it easier to narrow down the target protein.
- Is it necessary to use the recommended buffer as the binding buffer?
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There are no problems with using TBS in place of HEPES, and NaCl in place of KCl.
- Why is it that both salt elution and boil elution are performed for elution?
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This is because bonds are divided between somewhat weaker bonds (which can be broken by salt) and strong bonds (which can be broken by a sample buffer plus heating). However there is no problem with performing boil elution alone.
- Does it happen that the band of bound protein becomes thin when the concentration of ligand is increased?
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When the amount of ligand immobilization is increased, the beads become hydrophobic and easily aggregate, which may reduce the reaction efficiency with the protein and thin the band. If bead agglomeration is seen, reduce the amount of ligand immobilization.
- Why can’t I see any bands of bound proteins?
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Since the amount of ligand immobilization may be small, consider increasing the immobilization concentration. The affinity between the ligand and the target protein may be weak, so try lowering the salt concentration in the buffer.
Also, if the target protein is low in the protein solution, you may need to consider increasing the concentration or volume of the protein solution.
- How long is the stable period of the ligand-immobilized beads?
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It depends on the stability of the ligand itself and the stability of the bond between the beads and the ligand. If the ligand itself is stable and the bond is an amide bond, there is no problem for at least one year.
- Is the optimal binding reaction time of 4 hours?
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In our experience, 4 hours is sufficient, but it is advisable to investigate the optimal reaction time as it depends on the nature of the ligand immobilized on the beads and the target protein.
- What is the optimal bead type for immobiliding antibodies?
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The optimum beads vary depending on the method of use and the bonding method. See here for details.
- Can I quantify the immobilization amount of the antibodies on the beads?
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It can be carried out by direct quantification of the immobilized antibodies. Please refer to Protocol 107 on our website.
- What is the efficiency when immobiliding antibodies?
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If binding is performed when 50 μg of antibodies is supplied to 1 mg of NHS beads, approximately 20 – 40 μg will be bound, although the result varies depending on the antibody animal, subclass, and clone. It is possible to increase the amount that is bound by increasing the amount on feed. When streptavidin beads are used, up to approximately 10 μg of biotin-modified antibodies will bond to 1 mg of Streptavidin beads or NeutrAvidin beads, although this also varies depending on the antibody.
- Is there a way to increase the antibody immobilization efficiency (immobilization amount)?
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There are several ways. Please refer to the following.
(1) In the case of NHS beads, the amount of immobilization may increase by lengthening the immobilization reaction time. However, the NHS group is susceptible to hydrolysis and causes non-specific adsorption, so the reaction time should be within 6 hours.
(2) In the case of NHS beads, the immobilization reaction may be hindered by a composition other than the recommended immobilization buffer. If the antibody storage solution contains additives such as salts or glycerol, buffer substitution will improve the immobilization efficiency.
(3) The amount of immobilization increases by increasing the amount of antibody added during the immobilization reaction.
- Can I disperse antibody-immobilized beads by ultrasonic device?
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Please perform the dispersion of the beads by the manual agitation. When you cannot disperse the beads easily, disperse them in a short time by using an ice-cold ultrasonic homogenizer or ultrasonic washer. Please refer to the following.
- When immobilizing antibodies to beads, the beads may adhere to the wall of the tube. Is there a way to suppress this?
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It may be improved by the following methods.
-Use a protein low adsorption tube.
-Agitate with a microtube mixer instead of overturning and mixing.
-After the immobilization reaction, the beads are dispersed by manual agitation and centrifugal (magnetic) separation is performed.
- How can I improve dispersibility of antibodies immobilized beads?
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Antibodies or proteins immobilized beads may easily precipitate. Please disperse the beads well before use. In addition, dispersibility improves if salt is removed from the buffer.
- What is the optimal bead type for binding DNA?
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Single-stranded and double-stranded DNA can be immobilized to the Plain beads.
- I want to analyze bound proteins with MS, but what should I do if the target protein band is thin?
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Increase the scale of affinity purification (ex: 2.5 mg / 1000 μL), or increase the number of samples under the same conditions to perform affinity purification.
- How much protein can be analyzed by MS?
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It depends on the type of equipment used, but it can be analyzed if the amount of protein is about 50 ng.
