S-Tag Monoclonal Antibody

What is the S-Tag epitope system and how does it compare to other tagging systems?

The S-Tag epitope system consists of a small 15-amino acid peptide tag (KETAAAKFERQHMDS) that can be fused to proteins of interest and detected using specific anti-S-Tag antibodies. Compared to other common tagging systems like FLAG (DYKDDDDK) or HA (YPYDVPDYA), the S-Tag offers several advantages, including its small size, the ability to purify tagged polypeptides without antibodies, and very low cross-reactivity with endogenous proteins in mammalian cells. This makes the S-peptide antibody an attractive alternative for many researchers working with protein expression and purification systems .

How can I incorporate the S-Tag into my protein of interest?

To incorporate the S-Tag into your protein of interest, you need to modify your protein expression vector with a nucleotide sequence encoding the S-peptide tag. The gene of interest should be cloned in-frame relative to the tag sequence, ensuring that upon expression, your protein will be synthesized as a fusion protein containing the S-peptide tag. This typically involves standard molecular cloning techniques such as PCR amplification with primers containing the S-Tag sequence, followed by restriction enzyme digestion and ligation into an appropriate expression vector. The tag can be positioned at either the N-terminus or C-terminus of your protein, depending on your experimental requirements and the protein's structural considerations .

What detection methods can be used with S-Tag labeled proteins?

S-Tag labeled proteins can be detected using several standard laboratory techniques. Western blotting is commonly employed using antibodies such as the MA1-981 monoclonal antibody or ab19321 polyclonal antibody, which specifically recognize the S-peptide epitope. Immunohistochemistry and immunofluorescence are also effective for visualizing S-tagged proteins in tissue sections or cultured cells. Additionally, ELISA (Enzyme-Linked Immunosorbent Assay) can be used for quantitative detection of S-tagged proteins in solution. The availability of different antibody formats (monoclonal and polyclonal) allows researchers to select the most appropriate reagent for their specific experimental conditions .

What cell types and expression systems are compatible with S-Tag technology?

S-Tag technology has been successfully implemented in various expression systems, including mammalian cell lines (such as K562 cells), bacterial systems (E. coli), and yeast cells. The versatility of the S-Tag system makes it applicable across diverse experimental models. When selecting an S-Tag antibody for your experiments, ensure it has the appropriate species reactivity for your expression system. The low endogenous expression of proteins that cross-react with the S-peptide in mammalian tissues makes this system particularly valuable for mammalian cell culture applications, reducing the likelihood of background signal .

How can I optimize S-Tag antibody performance in multiplexed immunoassays?

For multiplexed immunoassays using S-Tag antibodies alongside other antibodies, careful consideration of antibody compatibility is essential. To achieve optimal performance, select primary antibodies from different host species (e.g., mouse anti-S-Tag and rabbit anti-target protein) and use species-specific secondary antibodies conjugated to distinct fluorophores. This approach prevents cross-reactivity between detection systems. Alternatively, if using antibodies from the same host species, select antibodies of different isotypes or subtypes (e.g., mouse IgG1 vs. mouse IgG2a) and use subclass-specific secondary antibodies. For complex multiplexing, consider directly conjugated primary antibodies to eliminate secondary antibody cross-reactivity issues. Always perform preliminary single-staining controls to validate antibody performance before attempting multiplexed detection .

What are the biophysical considerations when using S-Tag for protein purification and characterization?

When using the S-Tag for protein purification and subsequent biophysical characterization, several factors should be considered. The small size (15 aa) of the S-Tag minimizes interference with protein folding and function, but its impact should still be validated for each specific protein. Consider performing parallel experiments with N-terminal and C-terminal tagged versions to identify any position-dependent effects. For biophysical analyses, evaluate whether the tag affects properties such as thermal stability, aggregation propensity, and colloidal stability. High-throughput developability assays can be employed to assess these parameters with small amounts of purified material (<1 mg). If the tag affects critical biophysical properties, consider removing it via engineered protease cleavage sites after initial purification steps .

How reliable is epitope accessibility in different structural contexts for S-Tag detection?

Epitope accessibility can vary significantly depending on the structural context of the fusion protein. The S-Tag may become partially or completely inaccessible when placed adjacent to structured domains or when the fusion protein adopts certain conformations. To address potential accessibility issues, consider incorporating flexible linker sequences (such as GSGSGS) between the S-Tag and your protein of interest. For membrane proteins or complex multi-domain proteins, perform epitope mapping experiments to determine optimal tag placement. Native versus denaturing detection conditions can also significantly impact epitope accessibility; while some antibodies like mAb G196 show strong recognition preference under specific conditions, S-Tag antibodies often retain functionality under both native and denaturing conditions, making them versatile for various application contexts .

What are the documented cross-reactivity concerns with S-Tag antibodies in different experimental systems?

Cross-reactivity concerns with S-Tag antibodies appear to be minimal compared to some other epitope tag systems. Database analysis similar to that performed for other epitope tags (such as the G196 tag system) typically reveals very few potential cross-reactive proteins. For example, a comparable analysis for the G196 tag found only 11 hits in the entire UniProtKB/Swiss-Prot database. For S-Tag antibodies, the specificity is generally high, with minimal cross-reactivity in mammalian cells. Nevertheless, it is prudent to include appropriate controls in your experiments, such as non-tagged versions of your expression constructs and wild-type (non-transfected) cells, to identify any potential cross-reactivity with endogenous proteins. This is particularly important when using the system in new cell types or organisms not previously validated .

How should I design expression constructs to maximize S-Tag functionality?

When designing expression constructs with the S-Tag, several factors can maximize functionality. First, consider tag placement—N-terminal tagging is often preferred for secreted proteins, while C-terminal tagging may be better for proteins where the N-terminus is critical for function. Include a flexible linker sequence (3-6 amino acids, typically glycine-serine repeats) between the tag and your protein to improve accessibility and reduce structural interference. For complex experimental designs, dual-tagging strategies can be advantageous; for example, combining S-Tag with FLAG or HA tags at opposite termini allows for orthogonal detection and purification options. For proteins expressed at low levels, consider codon optimization of both the tag and protein sequences for your expression system. Finally, include appropriate controls in your experimental design, such as untagged versions of your protein and empty vector controls .

What purification strategies are most effective for S-Tagged proteins?

S-Tagged proteins can be purified using several strategies, with affinity chromatography being the most common approach. S-protein resin, which contains immobilized S-protein (a ribonuclease S component that binds with high affinity to the S-peptide), provides a specific purification method that doesn't require antibodies. For higher purity requirements, consider a two-step purification process combining S-protein affinity chromatography with size exclusion or ion exchange chromatography. If using antibody-based purification, both monoclonal (e.g., MA1-981) and polyclonal (e.g., ab19321) anti-S-Tag antibodies can be immobilized on suitable matrices like Protein G or directly coupled to sepharose. For difficult-to-express or poorly soluble proteins, optimization of lysis conditions (buffer composition, detergents, salt concentration) can significantly improve purification yields while maintaining the S-Tag interaction integrity .

How can I effectively use S-Tag for protein-protein interaction studies?

S-Tag provides a valuable tool for protein-protein interaction studies through various experimental approaches. For co-immunoprecipitation (co-IP) experiments, use anti-S-Tag antibodies to pull down your S-tagged protein of interest along with its interaction partners. The small size of the S-Tag minimizes steric interference with protein-protein interactions compared to larger tags. For identifying novel interaction partners, S-Tag pull-downs followed by mass spectrometry analysis offer a powerful unbiased approach. When designing these experiments, consider including appropriate controls such as S-tagged unrelated proteins and beads-only conditions to identify non-specific interactions. For detecting interactions in living cells, the S-Tag can be combined with proximity-based labeling techniques like BioID or APEX by creating fusion constructs. Finally, for quantitative binding studies, surface plasmon resonance or isothermal titration calorimetry can be employed with purified S-tagged proteins to determine binding affinities and kinetics .

What considerations are important when using S-Tag antibodies for immunofluorescence microscopy?

When using S-Tag antibodies for immunofluorescence microscopy, several factors can optimize results. First, fixation method significantly impacts epitope accessibility—test multiple approaches (paraformaldehyde, methanol, acetone) to determine optimal conditions for your specific fusion protein. Permeabilization conditions should also be optimized; excessive permeabilization can lead to antigen loss, while insufficient permeabilization prevents antibody access to intracellular compartments. Blocking conditions should be stringent to minimize background (typically 5-10% serum or BSA with 0.1-0.3% Triton X-100). For co-localization studies, select antibodies from different host species to enable simultaneous detection with species-specific secondary antibodies. Signal amplification using biotinylated secondary antibodies and fluorescent streptavidin can enhance detection of low-abundance proteins. Finally, include important controls: untagged protein expression, secondary antibody-only controls, and competing peptide controls to confirm specificity .

How can I address weak or absent S-Tag antibody signal in Western blotting?

When encountering weak or absent S-Tag antibody signal in Western blotting, systematically troubleshoot the issue through several approaches. First, verify protein expression using alternative detection methods like direct fluorescence (if applicable) or an orthogonal tag if your construct contains multiple tags. Optimize protein extraction conditions—different lysis buffers (RIPA, NP-40, etc.) may be required depending on protein localization and solubility. For membrane proteins or hydrophobic proteins, consider adding appropriate detergents. If the signal remains weak, increase protein loading amount or concentrate your sample using precipitation methods (TCA or acetone). Testing different blocking agents can also help, as some proteins may interact non-specifically with certain blocking reagents. For transfer issues, try various membrane types (PVDF versus nitrocellulose) and transfer conditions (wet versus semi-dry). Finally, consider enhancing detection sensitivity using amplification systems such as biotin-streptavidin or tyramide signal amplification methods .

What strategies can resolve non-specific binding or high background with S-Tag antibodies?

To resolve non-specific binding or high background issues with S-Tag antibodies, implement several optimization strategies. First, increase the stringency of your washing steps by adding higher concentrations of detergent (0.1-0.5% Tween-20 or Triton X-100) to wash buffers. More frequent and longer washing steps can significantly reduce background. Optimize your blocking conditions by testing different blocking agents (BSA, non-fat milk, casein, commercial blocking buffers) and concentrations (3-10%). Titrate your primary antibody concentration to find the optimal signal-to-noise ratio—too concentrated antibody solutions often increase non-specific binding. For difficult samples, pre-adsorption of the antibody with non-specific proteins from the same species as your sample can reduce cross-reactivity. Adding reducing agents like DTT or β-mercaptoethanol to your sample buffer can help reduce non-specific bands caused by disulfide linkages in Western blotting. Finally, consider using more specific detection methods, such as monoclonal antibodies instead of polyclonal antibodies, which typically exhibit lower cross-reactivity .

How should I interpret conflicting results between different detection methods using S-Tag antibodies?

When faced with conflicting results between different detection methods using S-Tag antibodies, a systematic analytical approach is necessary. Create a comparison table of your results across methods, noting key variables such as sample preparation, antibody dilutions, and detection systems used. Consider that epitope accessibility varies significantly between methods—Western blotting detects denatured proteins, while immunofluorescence and immunoprecipitation often interact with native conformations. Antibody performance can differ dramatically between these contexts. For discrepancies between antibody-based and tag-based detection systems, consider that post-translational modifications or proteolytic processing may affect tag recognition. Confirm your findings using orthogonal approaches, such as mass spectrometry or functional assays, to validate protein identity and modification state. Consult published antibody validation studies for known limitations of specific antibodies. Finally, consider that cellular localization can impact detection efficiency—membrane-bound, nuclear, or aggregated proteins may require specialized extraction or detection protocols to ensure consistent results across methods .

What quantitative analysis approaches are most appropriate for S-Tag detection systems?

For quantitative analysis using S-Tag detection systems, several approaches can be implemented depending on the experimental context. For Western blot quantification, densitometry analysis using software like ImageJ should include normalization to appropriate loading controls and standard curves generated with known concentrations of purified S-tagged protein. For immunofluorescence quantification, consider mean fluorescence intensity measurements, fluorescence correlation spectroscopy for dynamic studies, or high-content imaging for population-level analyses. When comparing expression levels across different conditions, relative quantification is typically sufficient, but absolute quantification requires calibration with purified standards. For higher-throughput analyses, flow cytometry of cells expressing S-tagged fluorescent proteins can provide quantitative data across large cell populations. ELISA-based quantification offers another sensitive approach, especially for secreted proteins, with detection limits typically in the picogram to nanogram range. Finally, newer techniques like digital ELISA or single-molecule counting methods can push detection sensitivity into the femtomolar range for ultra-low abundance proteins .

How can S-Tag technology be integrated with high-throughput antibody developability assessments?

S-Tag technology can be effectively integrated into high-throughput antibody developability workflows by serving as a standardized detection system across diverse antibody candidates. For large-scale screening of antibody developability (100s-1000s of molecules), consistent tagging with S-peptide enables uniform purification and detection protocols, minimizing variability in downstream assays. Implement parallel processing in 96- or 384-well formats with S-tagged antibody variants expressed in appropriate mammalian cell lines, followed by standardized purification using S-protein affinity resins. This approach allows critical developability parameters to be assessed with minimal material (<1 mg), including tendency for self-interaction, aggregation propensity, thermal stability, and colloidal stability. Create comprehensive data tables comparing physicochemical properties across antibody candidates, enabling ranking based on developability profiles. Integration with efficient data management systems allows correlation between early-stage S-Tag-based biophysical assays and downstream manufacturing parameters, facilitating data-driven selection of candidates with optimal properties for further development .

How can S-Tag be utilized in advanced protein engineering applications?

S-Tag offers versatile applications in advanced protein engineering, particularly when developing novel therapeutic proteins or research tools. For directed evolution experiments, the S-Tag provides a consistent detection handle across diverse protein variants, enabling high-throughput screening via FACS, phage display, or yeast surface display. When engineering bi-specific or multi-specific proteins, S-Tag can serve as a standardized detection epitope across fusion proteins with different functional domains. For protein half-life extension strategies, S-Tag can be used as an internal control tag while testing various half-life extension moieties like Fc domains or albumin-binding peptides. In protein conjugation chemistry, terminal S-Tags with engineered unique amino acids (e.g., cysteine or non-natural amino acids) provide site-specific conjugation points for chemicals, drugs, or imaging agents. For biosensor development, S-Tag can provide a consistent immobilization strategy across different sensor protein variants. Finally, in cell-free protein synthesis systems, S-Tag enables rapid detection and quantification of synthesized proteins, facilitating optimization of cell-free expression conditions for difficult-to-express proteins .

What role can S-Tag play in monitoring protein degradation and turnover studies?

S-Tag provides valuable capabilities for monitoring protein degradation and turnover dynamics in cellular systems. For pulse-chase experiments, S-tagged proteins can be immunoprecipitated at various time points to measure degradation kinetics following synthesis inhibition with cycloheximide or after pulse-labeling with radioisotopes or click chemistry-compatible amino acids. When combined with ubiquitin detection methods, S-Tag immunoprecipitation enables tracking of ubiquitination patterns preceding degradation. For targeted degradation studies (e.g., evaluating PROTAC molecules), S-Tag provides a consistent detection method across different target proteins. In analyzing proteasomal versus lysosomal degradation pathways, S-tagged proteins can be monitored following treatment with pathway-specific inhibitors (e.g., MG132 for proteasome, bafilomycin A1 for lysosomes). For studying protein half-life across different cellular compartments, the S-Tag can be combined with compartment-specific targeting signals. Finally, when implementing live-cell degradation reporters, S-Tag can be fused with fluorescent proteins in tandem with degrons or other degradation signals to create biosensors for real-time degradation monitoring across different cellular conditions .

How is S-Tag technology evolving for single-cell and spatial biology applications?

S-Tag technology is being adapted for cutting-edge single-cell and spatial biology applications through several innovative approaches. For single-cell proteomics, S-tagged proteins combined with highly sensitive detection methods enable protein quantification at the single-cell level, allowing researchers to understand cell-to-cell variability in protein expression and localization. In spatial transcriptomics-proteomics integration, S-Tag antibodies are being combined with in situ hybridization techniques to simultaneously visualize protein localization and mRNA expression within tissue contexts. S-Tag detection is also being integrated with microfluidic platforms for high-throughput single-cell isolation and analysis of protein expression patterns. For more advanced spatial applications, multiplexed imaging techniques using cyclic immunofluorescence with S-Tag antibodies allow visualization of dozens of proteins in the same tissue section. Finally, ongoing antibody engineering efforts aim to produce smaller antibody formats (nanobodies or single-domain antibodies) against the S-Tag, potentially improving tissue penetration and reducing background in complex tissue environments. These developments collectively expand the utility of S-Tag systems for studying heterogeneous cell populations and complex tissue architectures .

What are the future prospects for combining S-Tag with emerging protein analysis technologies?

The integration of S-Tag technology with emerging protein analysis platforms promises to expand its utility across multiple research domains. Next-generation mass spectrometry approaches, including data-independent acquisition and targeted proteomics, can leverage S-tagged proteins as internal standards for absolute quantification across complex samples. The combination of S-Tag with proximity labeling technologies (BioID, APEX, TurboID) enables mapping of protein interaction networks in specific cellular compartments with high temporal resolution. For structural studies, techniques like cryo-electron tomography and integrative structural biology approaches can utilize S-Tag as consistent handles for protein identification within complex cellular contexts. In high-throughput functional genomics, S-Tag compatibility with CRISPR-based genetic screens facilitates rapid phenotypic assessment of genetic perturbations on protein expression, localization, and modification. Advanced biophysical techniques like super-resolution microscopy and single-molecule tracking can employ S-tagged proteins for studying nanoscale protein organization and dynamics in living cells. Finally, computational approaches integrating machine learning with S-Tag detection data are emerging to predict protein behavior based on sequence and structural information, potentially accelerating protein engineering and therapeutic development workflows .

Comparative Analysis of Common Epitope Tag Systems

S-Tag Antibody Performance in Various Applications

Permutation Analysis of Epitope Recognition

Based on similar epitope tag permutation studies, amino acid substitutions at critical positions within epitope tags can significantly impact antibody recognition. While specific permutation data for S-Tag is not provided in the search results, comparable analysis for the G196 epitope tag showed that:

• Position 1 (Asp) could be replaced by Glu, Gly, and Ser while maintaining antibody recognition

• Position 2 (Leu) could be substituted with hydrophobic amino acids (Ile, Met, Phe, Asn) but not Val

• Other positions showed varying degrees of tolerance for substitutions

This type of analysis reveals the molecular basis for antibody-epitope interactions and provides valuable insights for designing modified tags with enhanced properties or specialized functions while maintaining antibody recognition .

Developability Parameters for Epitope-Tagged Proteins