RPA2 Monoclonal Antibody

What is the biological function of RPA2 in DNA metabolism?

RPA2 functions as a critical component of the heterotrimeric replication protein A complex (RPA/RP-A) that binds and stabilizes single-stranded DNA intermediates formed during DNA replication or upon DNA stress. It prevents DNA reannealing while simultaneously recruiting and activating various proteins and complexes involved in DNA metabolism. This dual function allows RPA2 to play essential roles in both DNA replication and cellular response to DNA damage. In damage response pathways specifically, the RPA complex controls DNA repair and checkpoint activation through recruitment of ATRIP, which activates the ATR kinase—a master regulator of DNA damage response . Additionally, RPA2 facilitates recruitment of double-strand break repair factors RAD51 and RAD52 to chromatin in response to DNA damage and interacts with nucleotide excision repair proteins like XPA and XPG .

How does RPA2 differ from other subunits in the RPA complex?

RPA2 (also known as RPA32) is the 32 kDa middle subunit of the heterotrimeric RPA complex, distinguishing it from the larger RPA1 (70 kDa) and smaller RPA3 (14 kDa) subunits. While all three subunits contribute to the complex's ability to bind single-stranded DNA, RPA2 specifically undergoes extensive phosphorylation in response to DNA damage, serving as a critical regulatory mechanism. The phosphorylation state of RPA2 modulates the activity of the entire RPA complex and influences its interactions with various DNA repair proteins . This positions RPA2 as a key signal transduction component within the complex, translating DNA damage detection into appropriate repair pathway activation. The 32 kDa molecular weight of RPA2 makes it readily identifiable on western blots, as confirmed by validation studies showing specific band detection at approximately 32 kDa .

Why are RPA2 monoclonal antibodies important tools in DNA damage research?

RPA2 monoclonal antibodies serve as critical tools in DNA damage research because they enable precise detection of RPA2 localization, quantification, and post-translational modifications in response to genotoxic stress. Since RPA2 undergoes extensive phosphorylation following DNA damage, antibodies targeting total RPA2 or specific phosphorylated forms provide valuable insights into damage response activation and progression. These antibodies facilitate the study of replication stress, DNA break repair kinetics, and checkpoint activation through various techniques including western blotting, immunofluorescence, and flow cytometry . The availability of highly specific monoclonal antibodies with validated reactivity across multiple species (human, mouse, rat) enables comparative studies between model organisms and human samples, accelerating translational research efforts . Additionally, the ability to immunoprecipitate RPA2 allows researchers to identify novel interaction partners and characterize protein complexes forming during different stages of the DNA damage response .

What criteria should researchers consider when selecting an appropriate RPA2 monoclonal antibody?

Researchers should evaluate multiple critical parameters when selecting an RPA2 monoclonal antibody for their specific experimental needs. First, consider the host species and clone type—both mouse monoclonal (such as clone 9H8 or 3B2E9) and rabbit monoclonal (like EPR2877Y) antibodies are available with distinct properties . Species cross-reactivity is equally important, with some antibodies recognizing only human RPA2, while others detect mouse and rat homologs as well . The intended application dictates antibody selection, as performance varies across techniques—validation data demonstrates that certain clones excel in western blotting but may show limited efficacy in immunofluorescence or flow cytometry . For phosphorylation studies, researchers must verify whether the antibody recognizes total RPA2 or specific phosphorylated epitopes, as this distinction is crucial for monitoring DNA damage response. Additionally, examine validation data showing antibody specificity, including western blots revealing appropriate molecular weight (32 kDa) bands and immunofluorescence/flow cytometry results confirming nuclear localization patterns . Finally, citation records provide valuable insights into antibody reliability across diverse experimental systems and conditions .

How can researchers validate the specificity of an RPA2 monoclonal antibody in their experimental system?

Validation of RPA2 monoclonal antibody specificity requires a multi-faceted approach. Begin with western blot analysis using positive control lysates from cell lines known to express RPA2 (such as U20S, HEK293, Jurkat, or K562 cells) alongside RPA2-depleted samples generated through siRNA or CRISPR-Cas9 knockout . A specific antibody should detect a single band at the expected molecular weight (32 kDa) that disappears in knockdown/knockout samples. For immunostaining applications, compare staining patterns between wild-type and RPA2-depleted cells, expecting predominantly nuclear localization that intensifies following DNA damage induction with agents like hydroxyurea or UV radiation . Peptide competition assays provide additional validation by demonstrating signal reduction when the antibody is pre-incubated with excess RPA2 peptide. For phospho-specific antibodies, treatment with lambda phosphatase should eliminate signal. Flow cytometry validation should include appropriate isotype controls and unstained samples as demonstrated in the Boster Bio validation data using JK cells . Finally, immunoprecipitation followed by mass spectrometry can confirm antibody specificity by identifying RPA2 and known interaction partners in the immunoprecipitated complex .

What are the common pitfalls in RPA2 antibody-based experiments and how can they be avoided?

Several common pitfalls can compromise RPA2 antibody experiments. First, cross-reactivity with other RPA subunits or unrelated proteins may occur, particularly with polyclonal antibodies. This can be mitigated by using well-characterized monoclonal antibodies like 9H8, 3B2E9, or EPR2877Y with demonstrated specificity . Second, cell fixation conditions significantly impact epitope accessibility, especially for nuclear proteins like RPA2. For immunofluorescence, researchers should optimize fixation methods (4% paraformaldehyde with permeabilization is effective as shown in validation data) and include antigen retrieval steps for tissue sections. Third, RPA2 phosphorylation status changes rapidly during cell cycle progression and following DNA damage, potentially confounding results if experimental timing is inconsistent. Researchers should synchronize cells and precisely control treatment durations. Fourth, antibody batch variability can introduce inconsistencies—performing lot-specific validation is advisable. Finally, detection sensitivity varies across applications; western blotting typically requires 1:1000 dilution while immunoprecipitation works optimally at 1:50 . To avoid these pitfalls, researchers should include appropriate positive controls (cell lines with confirmed RPA2 expression), negative controls (knockdown/knockout samples), and technical controls (isotype antibodies, secondary-only staining) in all experiments, while meticulously documenting experimental conditions to ensure reproducibility.

How should researchers optimize western blotting protocols for RPA2 detection?

Optimizing western blotting for RPA2 detection requires attention to several technical parameters. Sample preparation should include protease and phosphatase inhibitors to preserve RPA2 integrity and phosphorylation status. Validation data shows optimal results using 30 μg of total protein per lane under reducing conditions . Select an appropriate gel percentage; 5-20% gradient SDS-PAGE gels provide excellent resolution for the 32 kDa RPA2 protein . For protein transfer, nitrocellulose membranes with medium pore size (0.45 μm) perform well with transfer conditions of 150 mA for 50-90 minutes . Blocking with 5% non-fat milk in TBS for 1.5 hours at room temperature effectively minimizes background signal . Antibody concentrations require careful titration, with validated dilutions ranging from 0.5 μg/mL (clone 3B2E9) to 1:1000 (CST antibody) . Overnight incubation at 4°C maximizes signal specificity, followed by thorough washing with TBS containing 0.1% Tween (three washes, 5 minutes each) . For detection, HRP-conjugated secondary antibodies at 1:10,000 dilution with enhanced chemiluminescence provide excellent sensitivity . Include appropriate controls: positive controls (cell lines with known RPA2 expression such as U20S, HEK293, Jurkat, and K562), negative controls (RPA2-depleted samples), and loading controls (housekeeping proteins like GAPDH or β-actin). When studying RPA2 phosphorylation, additional controls including lambda phosphatase treatment help validate phospho-specific signals.

What is the optimal methodology for detecting RPA2 in fixed cells and tissues via immunofluorescence and immunohistochemistry?

For optimal immunofluorescence detection of RPA2 in fixed cells, begin with appropriate fixation protocols. Validated methods include 4% paraformaldehyde fixation for 15-20 minutes at room temperature, followed by permeabilization using 0.2% Triton X-100 in PBS . For tissue sections, enzyme-based antigen retrieval has proven effective, as demonstrated in validation studies using IHC enzyme antigen retrieval reagent . Blocking with 10% normal serum from the secondary antibody host species (e.g., goat serum) for 1 hour effectively minimizes non-specific binding . Antibody incubation conditions vary by clone: EPR2877Y (Abcam) and 3B2E9 (Boster) have validated protocols using 5 μg/mL antibody concentration with overnight incubation at 4°C . Secondary antibody selection should match the primary host species, with fluorophore-conjugated anti-mouse or anti-rabbit IgG (e.g., DyLight 594-conjugated goat anti-mouse IgG) at 1:100-1:200 dilution for 30 minutes at 37°C . Nuclear counterstaining with DAPI and cytoskeletal visualization with phalloidin-iFluor 488 provide contextual information for RPA2 localization . For co-localization studies with other DNA repair proteins like γH2AX or 53BP1, sequential staining may be necessary to avoid cross-reactivity. Confocal microscopy with appropriate filter sets optimized for the specific fluorophores offers superior resolution of nuclear foci formation after DNA damage. For quantitative analysis, establish consistent image acquisition parameters and analyze at least 100-200 cells per condition using automated foci counting software.

How can researchers effectively use RPA2 antibodies in flow cytometry to analyze cell cycle-dependent changes or damage responses?

Flow cytometry with RPA2 antibodies enables quantitative analysis of cell cycle-dependent changes and DNA damage responses across large cell populations. Begin by optimizing cell fixation and permeabilization—validation data demonstrates effective protocols using 4% paraformaldehyde fixation followed by permeabilization buffer treatment . For intracellular staining, block cells with 10% normal goat serum to reduce background, then incubate with RPA2 primary antibody (1 μg per 1×10^6 cells is effective for clone 3B2E9) . Fluorophore-conjugated secondary antibodies (e.g., DyLight 488-conjugated goat anti-mouse IgG) at appropriate dilution (1:100-1:200) provide good signal-to-noise ratio with 30-minute incubation at 20°C . Include comprehensive controls: isotype control antibodies (matching the primary antibody class, e.g., mouse IgG at 1 μg per 1×10^6 cells), unlabeled samples (no primary or secondary antibody), and single-color controls for multiparameter experiments . For cell cycle analysis, co-stain with DNA content markers (propidium iodide or DAPI) and use bivariate plotting to visualize RPA2 levels across cell cycle phases. When analyzing DNA damage responses, include timepoint series after damage induction and consider co-staining with γH2AX to correlate RPA2 recruitment with double-strand break formation. For phospho-specific RPA2 detection, additional validation steps including phosphatase treatment controls are essential. Data analysis should include both percentage of positive cells and mean fluorescence intensity measurements to capture both population shifts and per-cell protein level changes.

How should researchers interpret discrepancies between RPA2 detection methods across different experimental platforms?

When encountering discrepancies in RPA2 detection across different experimental platforms, researchers should systematically evaluate several factors. First, epitope accessibility varies significantly between techniques: denatured proteins in SDS-PAGE expose different epitopes than partially-native proteins in immunofluorescence or flow cytometry. For instance, clone 9H8 (Abcam) performs optimally in western blotting but may show limited reactivity in immunoprecipitation . Second, examine fixation and extraction methods—nuclear proteins like RPA2 may be incompletely extracted in certain lysis buffers, or their epitopes masked by overfixation with crosslinking agents. Third, consider detection sensitivity thresholds; western blotting can detect low abundance proteins that may be below detection limits in immunofluorescence. Fourth, post-translational modifications (particularly phosphorylation) can alter antibody recognition, explaining why total RPA2 levels may appear unchanged in western blotting while phospho-specific antibodies show dramatic changes after DNA damage. Fifth, cell cycle variation significantly impacts RPA2 expression and localization, with S-phase cells showing elevated levels that may skew population-based assays differently than single-cell techniques . To resolve discrepancies, employ orthogonal validation approaches: confirm protein identity with mass spectrometry, validate cellular localization with fractionation followed by western blotting, and verify antibody specificity with genetic approaches (siRNA, CRISPR/Cas9 knockout controls). Data integration across techniques often provides complementary insights rather than contradictions when properly interpreted.

What are the most common technical challenges when detecting phosphorylated forms of RPA2, and how can they be addressed?

Detection of phosphorylated RPA2 presents several technical challenges. First, phosphorylation events are often transient and can be rapidly reversed by cellular phosphatases during sample preparation. This challenge can be mitigated by immediate sample processing in buffers containing phosphatase inhibitor cocktails (sodium fluoride, sodium orthovanadate, β-glycerophosphate) and maintaining samples at 4°C throughout processing . Second, antibody specificity for particular phosphorylation sites varies considerably—cross-reactivity between different phospho-sites occurs frequently. Researchers should validate phospho-specific antibodies using phosphatase treatment controls and phospho-mimetic/phospho-null RPA2 mutants when possible. Third, the stoichiometry of RPA2 phosphorylation is often low, making detection challenging. Signal amplification through enhanced chemiluminescence systems for western blotting or tyramide signal amplification for immunostaining can improve sensitivity . Fourth, phosphorylation patterns are highly dynamic and cell cycle-dependent; synchronize cell populations and establish precise timepoints after damage induction for reproducible results. Fifth, multiple phosphorylation events often occur simultaneously, creating complex migration patterns on SDS-PAGE gels—hyperphosphorylated RPA2 typically appears as slower-migrating bands above the main 32 kDa band . Phos-tag™ acrylamide gels enhance separation of different phosphorylated species. Finally, context-specific phosphorylation varies with damage type; compare phosphorylation patterns across different genotoxic agents (UV, hydroxyurea, ionizing radiation) to comprehensively characterize the phosphorylation response.

How can researchers distinguish between true RPA2 foci and non-specific staining in nuclear immunofluorescence experiments?

Distinguishing true RPA2 foci from non-specific staining in nuclear immunofluorescence requires rigorous controls and analysis protocols. True RPA2 foci exhibit distinct characteristics: they typically appear as discrete nuclear puncta that increase in number and intensity following DNA damage induction, are absent in RPA2-depleted cells, and colocalize with other replication or repair markers (like PCNA for replication foci or γH2AX for damage sites) . To differentiate from artifacts, implement these validation approaches: First, include biological controls—compare untreated versus damage-induced samples (UV irradiation or hydroxyurea treatment significantly increases RPA2 foci) and wildtype versus RPA2-knockdown cells . Second, employ technical controls including secondary-only staining, isotype control antibodies, and peptide competition assays to identify non-specific binding . Third, use confocal microscopy with appropriate z-sectioning (0.5-1 μm steps) to distinguish true nuclear foci from background or cytoplasmic staining. Fourth, quantify foci characteristics objectively—true RPA2 foci typically range from 0.5-2 μm in diameter with signal intensity significantly above background, while non-specific specks tend to be smaller, dimmer, and randomly distributed. Fifth, perform co-localization studies with established replication/repair markers (EdU for replication foci, γH2AX for damage sites) as true RPA2 foci show high Pearson's correlation coefficients (>0.6) with these markers in appropriate contexts . Finally, evaluate temporal dynamics—true damage-induced RPA2 foci follow predictable formation and resolution kinetics after genotoxic stress.

How can researchers utilize RPA2 antibodies to investigate replication fork dynamics and replication stress responses?

RPA2 antibodies offer powerful tools for investigating replication fork dynamics and stress responses through several advanced approaches. Chromatin immunoprecipitation (ChIP) using RPA2 antibodies (particularly EPR2877Y and CST #52448 which demonstrate high immunoprecipitation efficiency) enables mapping of RPA-bound regions at stalled or collapsed replication forks . This can be combined with sequencing (ChIP-seq) to generate genome-wide profiles of replication stress sites. For spatiotemporal analysis of replication dynamics, immunofluorescence pulse-labeling experiments combining RPA2 staining with thymidine analogs (EdU/BrdU) allow discrimination between normal replication forks and stressed forks based on colocalization patterns . To investigate protein recruitment kinetics at individual replication forks, proximity ligation assays (PLA) using RPA2 antibodies paired with antibodies against fork protection factors (like BRCA1/2, FANCD2, or SMARCAL1) provide quantitative measurements of protein-protein interactions during stress responses . Advanced microscopy techniques including super-resolution imaging (STORM, PALM) with RPA2 antibodies achieve 20-30 nm resolution of fork structures when combined with DNA fiber analysis. For mechanistic studies, immunoprecipitation of RPA2 followed by mass spectrometry enables identification of stress-specific interaction partners, while phospho-specific RPA2 antibodies allow monitoring of ATR/ATM-dependent signaling at forks . Live-cell imaging approaches using cell lines expressing fluorescently-tagged RPA2 complemented with validation by immunofluorescence using RPA2 antibodies provide dynamic visualization of fork progression and stalling events.

What novel insights can be gained by combining RPA2 antibody-based detection with other DNA damage response markers?

Combining RPA2 antibody detection with other DNA damage response (DDR) markers reveals sophisticated insights into repair pathway choice and coordination. Multiplex immunofluorescence using RPA2 antibodies together with pathway-specific markers enables characterization of repair mechanism selection at individual damage sites . For example, co-staining with RPA2 and RAD51 distinguishes resected DSBs undergoing homologous recombination (dual-positive) from those committed to non-homologous end joining (RPA2-negative/53BP1-positive). Temporal analysis of repair factor recruitment using synchronized RPA2 co-staining with γH2AX, 53BP1, BRCA1, and RAD51 establishes the precise choreography of repair events and identifies potential bottlenecks in repair progression. Flow cytometry combining RPA2 with cell cycle markers (such as cyclin proteins or EdU incorporation) and other DDR proteins quantifies cell cycle-specific damage responses across large populations . Proximity ligation assays between RPA2 and various interaction partners (like ATRIP, ETAA1, or SMARCAL1) quantify stimulus-specific protein-protein interactions in situ, revealing regulatory mechanisms that conventional co-immunoprecipitation might miss . ChIP-sequential approaches (sequential immunoprecipitation with different antibodies) using RPA2 antibodies followed by other repair factors maps the handover of damaged DNA between different protein complexes. Integration of these multi-parameter datasets using machine learning approaches can identify unique damage response signatures associated with different genotoxic agents or genetic backgrounds, potentially revealing novel therapeutic vulnerabilities in cancer cells with DDR defects.

How can researchers employ RPA2 antibodies to investigate the relationship between replication stress and genomic instability in cancer models?

RPA2 antibodies provide versatile tools for investigating links between replication stress and genomic instability in cancer models through multiple experimental approaches. Immunohistochemical analysis of human tumor tissue microarrays using validated RPA2 antibodies (such as EPR2877Y) with established protocols enables correlation between RPA2 expression/phosphorylation patterns and clinical outcomes or genetic backgrounds . For preclinical models, tissue-specific analysis of replication stress can be achieved through immunofluorescence co-staining of RPA2 with γH2AX in tumor sections from patient-derived xenografts or genetically engineered mouse models . To investigate mechanisms of oncogene-induced replication stress, researchers can perform western blot analysis of RPA2 phosphorylation status following induced expression of oncogenes like MYC, RAS, or cyclin E, using antibodies with validated reactivity across human and mouse species . Flow cytometry with RPA2 and DNA content markers quantifies cell cycle-specific replication stress levels in heterogeneous tumor populations and can be combined with cell sorting to isolate subpopulations for further genomic analysis . For therapeutic applications, high-throughput immunofluorescence screening using RPA2 antibodies can identify compounds that exacerbate replication stress specifically in cancer cells. ChIP-seq approaches with RPA2 antibodies map genome-wide replication stress hotspots that may correspond to sites of potential genomic instability, while sequential chromatin immunoprecipitation (re-ChIP) with antibodies against RPA2 and DNA break markers identifies regions simultaneously experiencing replication stress and DNA breakage. Integration of these RPA2-based approaches with genomic analyses (such as whole-genome sequencing to identify structural variations) establishes direct mechanistic links between sites of persistent replication stress and resultant genomic alterations.