Protein Analysis FAQ Summary
For protein samples that have been added to the Loading Buffer but have not yet undergone boiling denaturation, long-term storage requires a balance between protein stability and compatibility with subsequent experiments. Specific recommendations are as follows: 1. Recommended storage conditions Temperature It is recommended to change to -80°C to minimize protein degradation and protease activity. Freezing form: The samples can be stored directly in the original tube without the need for aliquoting, but if the sample volume is large, it is advisable to aliquot into single-use amounts to avoid repeated freeze-thaw cycles. Precautions: Ensure to avoid repeated freeze-thaw during the freezing process, as the presence of SDS in the Loading Buffer can lead to SDS precipitation, sample inhomogeneity, and abnormal migration on gels. 2. Pre-use treatment recommendations Retrieve the sample on the day of the experiment and boil immediately (95–100°C, 5 minutes); if SDS precipitation or layering is observed in the frozen sample, gently vortex or briefly warm (37–50°C) before boiling to dissolve it. 3. Other additional recommendations If the sample contains proteases (e.g., if no Protease Inhibitor has been added) or is of a less stable protein type (e.g., tissue lysate), consider adding protease inhibitors before freezing; avoid repeated freeze-thaw before freezing, especially applicable for preparing multiple batches of Western blots.
• How long is the protein re-boiling temperature and time? Can it be boiled after dilution?
Protein sample re-boiling (i.e., heating denaturation) is usually used to destroy the higher structures of proteins before loading on SDS-PAGE, ensuring that the electrophoretic migration mainly reflects molecular weight. The specific re-boiling temperature and time are as follows: 1. Regular re-boiling conditions Temperature: 95°C or 100°C (boiling water) Time: 5 minutes If the sample is viscous or of high concentration, or contains complex components like membrane proteins, it can be extended to 10 minutes, but avoid excessive boiling that leads to protein degradation. 2. Can it be boiled after dilution? Yes, but pay attention to the following points: 1. Ensure the Buffer system is complete After dilution, the sample should still contain reducing agents (such as DTT or β-mercaptoethanol) and SDS to ensure adequate denaturation and reduction. Otherwise, the boiling is not very meaningful. 2. The dilution factor should not be too large Excessive dilution may lead to too low protein concentration, affecting band clarity; generally, it is recommended to maintain >0.5 μg/μL after dilution. 3. It is not recommended to leave the boiled samples for a long time It is recommended to load the re-boiled samples as soon as possible, especially those containing SDS and reducing agents, as they are prone to degradation or precipitation.
For research on SUMO (Small Ubiquitin-like Modifier) modification, the experimental design needs to be precisely differentiated based on specific research objectives (e.g., whether focusing on global SUMOylation, modification at specific sites, or specific substrate proteins). The following layered explanations are provided: 1. Common strategies for SUMO-related experimental design 1. Detection of total SUMOylation levels Objective: To observe the dynamic changes of SUMOylation in cells/tissues under conditions such as stress or drug treatment. Method: Western blot to detect the overall SUMOylated protein profile; commonly used SUMO1 and SUMO2/3 antibodies to detect smear signals. 2. Detection of SUMOylation at specific sites/specific substrate proteins Objective: To verify whether a specific protein is SUMOylated, or to identify specific modification sites. Method: Co-IP + WB or Ni-NTA pull-down + WB to detect forms of SUMOylation, along with mutants (e.g., K→R) to validate modification sites. 3. SUMO E1/E2/E3 enzyme system intervention experiments Objective: To study the regulatory mechanisms of SUMOylation. Method: Gene knockout/overexpression of SUMO enzymes such as SAE1/2, UBC9, PIAS family, etc., to observe changes in substrate modifications. 4. Cellular localization and functional experiments Objective: To observe the impact of SUMOylation on protein function. Method: GFP fusion proteins + mutants (WT...
• What should I do next after completing the peptide sequence identification by mass spectrometry?
After obtaining the peptide sequence identification results from mass spectrometry, the next analysis should be conducted step-by-step based on the research objectives (qualitative, quantitative, functional studies). The general process is as follows: 1. Quality control and filtering of results. Check the scoring indicators of the database search results (such as Peptide/Protein FDR, Score, number of Unique peptides). Exclude low-confidence proteins or results supported by only one peptide to ensure data reliability. 2. Organization of protein identification and quantification. If only identification is performed, organize a list of identified proteins, UniProt IDs, coverage, etc.; if quantification is involved (Label-free, TMT, etc.), compile the protein abundance matrix for each sample, and perform normalization and missing value treatment. 3. Differential analysis (if group comparisons are involved). Perform statistical tests (t-tests, ANOVA, etc.) to screen for differential proteins; generally, set Fold Change and significance thresholds (e.g., FC≥1.5 or 2, p<0.05). 4. Functional annotation and pathway enrichment. Use databases (GO, KEGG, Reactome) for functional classification; enrichment analysis assesses the biological significance of differential proteins. 5. Further data mining. Protein interaction networks (STRING, Cytoscape); integration with transcriptomic and metabolomic data to explore regulatory mechanisms.
It is necessary to distinguish between two aspects: the effect of the protein's own structural state and the preservation conditions on stability. 1. Degradation risk of non-denatured proteins: Proteins in their natural conformation retain their complete spatial structure and usually maintain activity. If there are residual proteases or trace metal ions in the solution, such proteins degrade quickly due to their susceptibility to recognition and cleavage. Denaturing treatments (such as adding SDS, urea, or heating) cause loss of structure and activity, making it difficult for most proteases to cleave; however, denatured states are generally unsuitable for functional testing or structural studies. Therefore, non-denatured proteins require strict inhibition of protease activity (by adding a protease inhibitor cocktail) and should be kept at low temperatures. 2. Storage duration at -80°C: Under conditions containing glycerol (commonly 10-50%) or a small amount of buffering salt, proteins can be stably stored at -80°C for several months to a year, depending on the protein's own stability (structural complexity, disulfide bonds, whether it is a membrane protein, etc.). Without protectants, stored only in PBS or aqueous solution, freeze-thaw cycles can severely affect structure, and typically the activity noticeably declines within weeks to 1-2 months. To avoid frequent freeze-thaw: it is recommended to aliquot small volumes for single use. It is advised to clarify the type of protein and subsequent use (activity testing, structural studies, or mass spectrometry analysis), choose an appropriate buffering system and protectants, and add broad-spectrum protease inhibitors during preparation to significantly extend the storage period.
In liquid chromatography (LC) analysis, the relative concentration of the target substance in the sample is usually assessed through peak area, while time (Retention Time, RT) is primarily used to determine the identity of the components rather than directly indicating concentration. To compare the concentration levels of the target substance under different intervention conditions, the following steps should be taken: 1. Lock in the retention time of the corresponding compound. Retention times may drift slightly under different conditions, so confirmation of the same compound should be done using standard samples or characteristic ions (if using mass spectrometry). Only after confirming that they are the same peak can the areas be compared. 2. Use peak area to reflect relative concentration. Peak area is proportional to the amount injected or the concentration of that component in the sample (within the method's linear range). When comparing different intervention groups, simply compare the areas of the same compound's peaks. Larger area → higher relative concentration. 3. Normalize or correct with an internal standard. If there are slight differences in injection volume or detection conditions among different groups, normalization using an internal standard or total peak area is required to avoid systematic errors. 4. Pay attention to baseline, resolution, and detector linear range. Ensure peak integration consistency to avoid misjudgment of area due to tailing or unresolved peaks. If peak intensity approaches detector saturation, dilution is necessary for quantification. 5. Summary. The relative concentration of different intervention groups is mainly judged by the peak area of the same compound, and retention time is only used to confirm the identity of the compound, not for concentration comparison.
• Is the capillary tube for collecting tear fluid a disposable micro blood collection tube?
In general, the capillary tube used for collecting tear fluid is not an ordinary disposable micro blood collection tube, but a specialized glass or plastic capillary tube that is additive-free and uncoated. Common specifications are an inner diameter of 0.5–1.0 mm, a length of about 5–10 cm, and a capacity usually between 1–10 µL. The main differences are: 1. No anticoagulants or other additives: blood collection capillaries are usually coated with heparin or other anticoagulants, which may interfere with subsequent proteomics or metabolomics analyses. 2. No adsorptive coating or inert treatment: tear fluid samples are extremely small, and the concentrations of proteins and metabolites are low; any coating may cause loss of analytes or introduce background noise. 3. Precise capacity control: tear fluid collection often requires quantification (for example, for tear flow testing or quantitative omics), necessitating the use of capillary tubes with known inner diameters and precise graduations. Recommendations: 1. Use uncoated borosilicate capillary tubes specifically designed for collecting tear fluid or micro biological fluids, avoiding blood collection tubes. 2. Seal both ends promptly after collection to prevent evaporation, and store at low temperatures to minimize sample loss.
• How to determine the content of dimers under conditions with and without drug treatment?
To accurately compare the content of dimers under drug treatment and non-drug treatment conditions, it is essential to clarify: the properties of the protein (whether it is soluble, whether it is prone to aggregation), the detection environment (in vitro purified protein or cell/tissue lysate), and whether the objective is qualitative or quantitative analysis. The following strategies are generally considered: 1. Gel electrophoresis combined with quantification 1. Principle: Dimers can maintain conformational differences compared to monomers under non-reducing and low-denaturing conditions. 2. Method: Separate using non-reducing SDS-PAGE or Native-PAGE; after staining, use image analysis software (such as ImageJ) to perform grayscale integration of the monomer and dimer bands, calculating the dimer ratio; it is essential to ensure that the sample amount is within the linear range to avoid false positives caused by aggregation or excessive loading. 3. Applicability: Simple and intuitive, but may underestimate weakly bound or dynamically balanced dimers. 2. SEC (Size-Exclusion Chromatography)/ HPLC 1. Principle: Separation based on molecular weight can distinguish monomer and dimer peaks. 2. Method: Quantification of peak area through a standard curve (external standard protein or purified target protein of known concentration); it is recommended to combine with MALS (multi-angle light scattering) or UV detection to improve accuracy. 3. Applicability: Can provide a more accurate ratio and detect drug-induced dimerization transitions. 3. Crosslinking combined with mass spectrometry (Crosslinking-MS) or chemical crosslinking + Western 1. Principle: Drugs may alter the dynamics of dimerization, and specific dimers can be detected after crosslinking. 2. Method: Use mild crosslinkers (such as.....
Key considerations include the following points: 1. Protein Integrity 1. Since it has been boiled (usually containing SDS and reducing agents), most of the natural conformation and enzymatic activity in the sample have been destroyed, and theoretically, significant degradation will not occur. 2. However, prolonged exposure at room temperature may lead to the aggregation of some easily precipitating or insoluble components, especially when containing high concentrations of SDS or urea, which may precipitate upon cooling. 2. Microbial Contamination and Buffer System 1. If there are no antimicrobial agents in the buffer (besides high concentrations of SDS), bacteria may still grow when left at room temperature for a long time, leading to sample turbidity or degradation. 2. The pH of buffer systems like Tris may change slightly at room temperature, but usually, the impact is minimal. 3. The subsequent use determines whether it can continue to be used 1. If only for SDS-PAGE/Western blot: it can basically continue to be used, but it is recommended to centrifuge to remove any possible precipitate and reboil if necessary. 2. If enzyme experiments or mass spectrometry (especially quantitative mass spectrometry) are needed: it is not recommended to use directly due to the high risk of contamination and modification (oxidation, deamidation), which may affect data reliability. Suggested handling: First, centrifuge to check for any obvious precipitate or turbidity; if only for WB, it can be used directly or reboiled; if for mass spectrometry or functional experiments, it is recommended to reprepare or at least do a control to assess the impact.
• After predicting peptides using docking software, what should be used to validate the binding?
Typically, after predicting the binding mode of a peptide and target protein through molecular docking, experimental methods are required to validate the binding affinity and specificity. Common methods can be divided into two categories: 1. Biochemical or biophysical methods (direct measurement of binding) 1. Surface plasmon resonance (SPR) can obtain real-time binding kinetics parameters (ka, kd) and equilibrium dissociation constant (KD), suitable for validating affinity and kinetic characteristics. 2. Isothermal titration calorimetry (ITC) can directly determine thermodynamic parameters (ΔG, ΔH, ΔS) and affinity, suitable for high-affinity systems. 3. MicroScale Thermophoresis (MST) has low sample requirements and can quickly assess the binding constant. 4. Fluorescence polarization (FP) or FRET can be used for screening the binding of small molecules/peptides with proteins, but labeling is required. 2. Cellular or functional level validation (indirect evidence) 1. Co-immunoprecipitation (Co-IP) or pull-down experiments validate whether peptides and proteins form complexes in cells or in vitro. 2. Functional assays (signal pathway activation/inhibition, enzyme activity inhibition, etc.) validate whether the binding has biological effects. It is recommended that in the initial screening phase, MST or SPR be used to assess the actual affinity of the peptide to the target, comparing it to the docking prediction results. High-priority candidates can then be accurately measured for thermodynamic parameters using ITC. If the binding is stable and has biological function, then physiological relevance can be validated through cellular functional experiments.
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