CHR28 Antibody

What is CD28 and what cellular functions does it mediate?

CD28 is a 44 kDa homodimeric surface glycoprotein expressed by thymocytes, mature T cells, and plasma cells. Structurally, it belongs to the immunoglobulin superfamily and functions as a critical costimulatory receptor for T cell activation . CD28 acts as a ligand for CD80 (B7-1) and CD86 (B7-2) molecules expressed on antigen-presenting cells, providing the essential "second signal" required for complete T cell activation . This costimulatory pathway significantly augments interleukin-2 (IL-2) production, IL-2 receptor expression, and enhances the cytotoxicity of CD3-activated T cells . Beyond its costimulatory role, CD28 prevents T cells from entering an anergic (hyporesponsive) state and protects them from premature apoptotic cell death, making it essential for maintaining functional T cell responses in both physiological and pathological conditions .

How do CD28 antibodies differ in their mechanisms of action?

CD28 antibodies can be classified based on their binding epitopes and functional consequences. Conventional agonistic antibodies require TCR/CD3 engagement (signal 1) to provide costimulation, mimicking the natural CD28-CD80/CD86 interaction. In contrast, superagonistic antibodies like TGN1412 can activate T cells independently of TCR stimulation by clustering CD28 molecules, leading to potentially dangerous cytokine release syndrome . Non-superagonistic antibodies, such as the recently developed E1P2, bind to CD28 without causing T cell activation in the absence of TCR/CD3 stimulation . The binding epitope plays a crucial role in determining these functional properties - superagonistic antibodies typically bind to the lateral epitope of CD28, while non-superagonistic antibodies like E1P2 bind close to the apex of CD28, similar to where natural ligands CD80/CD86 bind . This epitope distinction explains why some CD28 antibodies require TCR costimulation while others can activate T cells independently.

What experimental considerations are important when using CD28 antibodies for flow cytometry?

When using CD28 antibodies for flow cytometric analysis, researchers should consider several technical aspects. First, titration is essential - pre-titrated antibodies like CD28.2 can be used at approximately 5 μL (0.25 μg) per test, with a test defined as the amount of antibody needed to stain a cell sample in a final volume of 100 μL . Cell concentration is another critical factor; while this may vary empirically, typical ranges are between 10^5 to 10^8 cells per test . For optimal performance with PE-conjugated CD28 antibodies, researchers should select appropriate excitation (488-561 nm) and emission (578 nm) parameters compatible with blue, green, or yellow-green lasers . Post-acquisition analysis should account for CD28's differential expression patterns across T cell subsets, with varying expression levels between immature and mature T cells. For instance, CD28 expression is highest on immature CD3-, CD8+, CD4+8+, and CD4-8- thymocytes that express α/β and γ/δ TCR, while mature CD4+ and CD8+ α/β TCR+ thymocytes express two- to four-fold lower levels .

How can phage display technology be used to develop novel CD28 antibodies?

Phage display technology represents a powerful approach for generating fully human antibodies with specific binding properties. For CD28 antibody development, this methodology can be strategically designed to favor binders toward particular epitopes. As demonstrated in the development of E1P2, researchers cloned and expressed the extracellular domain (ECD) of human CD28 with an Fc tag for homo-dimerization and an AviTag™ for site-specific biotinylation . The biotin moiety was strategically attached to the membrane-proximal part of CD28, near the C"D loop (the epitope of most superagonistic antibodies), to bias selection toward binders targeting the apex of CD28 .

Following protein production and quality control through size exclusion chromatography (SEC) and SDS-PAGE, the biotinylated antigen is captured on streptavidin-coated magnetic beads and exposed to a synthetic phage display single-chain variable fragment (scFv) library . To isolate CD28-specific binders and exclude Fc-binding clones, the selection process includes a parallel ELISA screening against human IgG1 . Successful clones can be reformatted into complete antibodies (e.g., IgG4 format) and expressed in mammalian cell lines for further characterization . This methodology enables epitope-biased selection that can help avoid isolating potentially dangerous superagonistic antibodies while favoring those with properties more similar to natural ligand interactions.

What assays should be used to evaluate potential superagonistic properties of CD28 antibodies?

Given the historical safety concerns with superagonistic CD28 antibodies like TGN1412, rigorous testing for potential superagonistic properties is essential for any new CD28 antibody. A comprehensive evaluation should include in vitro cellular assays using human peripheral blood mononuclear cells (PBMCs) from multiple donors to account for inter-individual variability . In these assays, the candidate antibody should be immobilized (wet-coated) on plates, followed by addition of freshly isolated human PBMCs .

Key parameters to measure include: T cell proliferation, cytokine secretion (particularly IL-2, IFN-γ, and TNF-α), and expression of early (CD69) and late (CD25) T cell activation markers, with special attention to CD4+ T cells, which are most susceptible to superagonistic activation . A true superagonistic antibody would induce these activation markers without concomitant TCR/CD3 stimulation, while conventional costimulatory antibodies would not . Additionally, potential species differences should be evaluated if the antibody cross-reacts with mouse CD28, as seen with E1P2 . For antibodies advancing to preclinical development, in vivo safety studies in humanized NSG mice represent a valuable model to assess the risk of cytokine release syndrome before human trials . These multi-faceted approaches can help identify potentially dangerous superagonistic properties early in development.

How can researchers optimize CD28 antibody-based costimulation in combination with CD3 bispecific antibodies for cancer immunotherapy?

The combination of CD3 bispecific T-cell engagers with CD28 costimulation represents a promising strategy to enhance anti-tumor immune responses. When optimizing such combinations, researchers should consider several experimental parameters. First, the epitope and binding characteristics of the CD28 antibody are critical—non-superagonistic antibodies that bind to the apex of CD28 (similar to natural ligands) tend to provide safer costimulation . Second, the ratio between CD3 and CD28 stimulation requires titration to identify optimal activation conditions without excessive cytokine release .

In experimental designs, researchers can evaluate the efficacy of combined CD3/CD28 targeting using in vitro tumor cell killing assays with human PBMCs as effector cells and tumor cell lines as targets . Key readouts should include: tumor cell death (e.g., by flow cytometry), T cell proliferation, cytokine production profiles (assessing both efficacy-related cytokines like IFN-γ and potential toxicity markers like IL-6 and TNF-α), and markers of T cell exhaustion to determine if CD28 costimulation prevents early T cell dysfunction . For formats, researchers can test various configurations including separate antibodies, bispecific constructs combining both CD3 and CD28 targeting, and tumor-targeted trispecific molecules that localize both signals to the tumor microenvironment . These optimization approaches can help identify combinations that maximize anti-tumor activity while minimizing systemic toxicity risks.

What lessons can be learned from the TGN1412 clinical trial that inform safer CD28 antibody development?

The 2006 TeGenero clinical trial of the superagonistic anti-CD28 antibody TGN1412 resulted in severe cytokine release syndrome in all six healthy volunteers, highlighting critical considerations for CD28-targeting therapeutic development . Several key lessons emerged from this event that continue to guide antibody development. First, binding epitope is crucial—TGN1412 bound to a lateral epitope of CD28, enabling antibody-mediated receptor clustering and T cell activation independent of TCR engagement . Newer approaches favor antibodies that bind to the apex of CD28, similar to natural ligands, requiring TCR costimulation for activity .

Second, preclinical testing strategies must be reevaluated. Despite extensive preclinical testing, the TGN1412 toxicity was not predicted in animal models . This reveals limitations in conventional toxicology models and underscores the need for human cell-based assays and potentially humanized mouse models that better recapitulate human immune responses . Third, initial dosing strategies for first-in-human trials of immunomodulatory agents should implement extremely conservative dosing—starting several orders of magnitude below the anticipated effective dose . Finally, the incident highlighted the importance of understanding mechanism of action thoroughly before clinical testing, particularly for novel immunomodulatory agents targeting costimulatory pathways . These lessons continue to shape the regulatory landscape and development strategies for immunomodulatory antibodies targeting T cell activation pathways.

What strategies can improve the safety profile of CD28-targeting therapeutic antibodies?

Multiple engineering approaches can enhance the safety profile of CD28-targeting antibodies. First, epitope-directed antibody discovery, as employed in the development of E1P2, can favor binding to regions of CD28 that require TCR/CD3 co-engagement for T cell activation . This approach uses strategic antigen presentation during phage display selections to bias toward non-superagonistic binders . Second, conditional activation systems can be designed where CD28 stimulation occurs only in the tumor microenvironment. This can be achieved through bispecific antibodies targeting both CD28 and tumor-associated antigens, ensuring costimulation is localized to tumor sites .

Third, antibody format and valency modifications can reduce the risk of receptor clustering. Monovalent CD28-binding formats or those with reduced crosslinking potential may provide costimulation without triggering superagonistic effects . Fourth, Fc engineering approaches can modulate effector functions and half-life. Modifications to reduce Fc receptor binding or complement activation can minimize unwanted inflammatory responses, while maintaining the desired costimulatory function . Finally, comprehensive in vitro screening using human PBMCs from multiple donors is essential, as there can be significant inter-individual variability in responses to CD28 stimulation . The combination of these approaches can help develop CD28-targeting therapeutics with improved safety profiles while maintaining efficacy in enhancing anti-tumor immune responses.

How do recent advances in CD28 antibody development contribute to next-generation cancer immunotherapy approaches?

Recent advances in CD28 antibody development are expanding the potential of cancer immunotherapy beyond current limitations. First, the development of non-superagonistic CD28 antibodies like E1P2 that enhance T cell activation only in the presence of TCR/CD3 stimulation addresses a key safety limitation in this field . These antibodies can potentially boost the efficacy of existing immunotherapies, including CD3 bispecific T-cell engagers, which have shown remarkable clinical outcomes against several hematological malignancies but may lead to insufficient T-cell activation and early exhaustion without proper costimulation .

Second, the cross-reactivity with both human and mouse CD28 demonstrated by newer antibodies like E1P2 enables more predictive preclinical testing in immunocompetent mouse models, potentially avoiding the translational failures seen with previous CD28-targeting agents . Third, epitope mapping techniques have revealed critical distinctions between superagonistic and costimulatory binding sites, enabling rational design of safer therapeutic candidates . Fourth, the combination of CD28 stimulation with other immunotherapy modalities represents a promising approach to overcome resistance mechanisms in solid tumors, particularly those with immunosuppressive microenvironments .

Finally, these advances contribute to the development of multi-specific antibody formats that can integrate tumor targeting, T cell recruitment via CD3, and costimulation via CD28 into single molecules, potentially offering superior efficacy and safety profiles compared to current immunotherapeutic approaches . Collectively, these developments may address key limitations of current cancer immunotherapies, including T cell exhaustion, poor infiltration into solid tumors, and resistance mechanisms in the tumor microenvironment.

What are the optimal protocols for evaluating CD28 antibody binding characteristics?

Comprehensive evaluation of CD28 antibody binding characteristics requires multiple complementary approaches. For initial binding assessment, enzyme-linked immunosorbent assay (ELISA) provides a quantitative measure of antibody-antigen interaction using recombinant CD28 proteins. As demonstrated with E1P2 antibody characterization, apparent affinity (EC50) can be determined in both scFv format (showing 31 nM affinity) and IgG4 format (showing improved affinity of 2.7 nM to human CD28) . When assessing cross-reactivity with orthologs, parallel ELISA against human and mouse CD28 can reveal species specificity, as seen with E1P2 binding to both species while TGN1412 bound only human CD28 .

Flow cytometry using primary T cells represents a crucial validation step, confirming binding to native CD28 as expressed on cell surfaces. Serial dilution experiments allow determination of binding EC50 values, with E1P2 and TGN1412 showing EC50 values of 4.9 nM and 2.5 nM respectively on human T cells . To ensure specificity, negative control cell lines lacking CD28 expression should be included to exclude non-specific binding . For more detailed epitope characterization, competition assays with known ligands (CD80/CD86) or other CD28 antibodies can reveal whether the novel antibody binds to overlapping or distinct epitopes . Advanced techniques like hydrogen-deuterium exchange mass spectrometry (HDX-MS) can provide detailed mapping of conformational epitopes, revealing critical differences between superagonistic and non-superagonistic antibodies . These multi-faceted approaches provide comprehensive binding profiles essential for understanding the functional consequences of CD28-antibody interactions.

What cellular assays best demonstrate the functional differences between superagonistic and conventional CD28 antibodies?

To distinguish between superagonistic and conventional CD28 antibodies, researchers should implement a systematic set of cellular assays focusing on T cell activation parameters. The primary differential characteristic is the ability of superagonistic antibodies to activate T cells without TCR/CD3 co-engagement . This can be evaluated through an immobilized antibody assay where CD28 antibodies are wet-coated on plates and incubated with freshly isolated human PBMCs in the absence of anti-CD3 or other TCR stimuli . In this context, superagonistic antibodies like TGN1412 will induce T cell proliferation (measurable by CFSE dilution or thymidine incorporation), while conventional antibodies like E1P2 will not .

Cytokine secretion profiling represents another critical readout, with superagonistic antibodies inducing significant release of IL-2, IFN-γ, and TNF-α without TCR stimulation . Flow cytometric analysis of activation markers provides additional discrimination, with superagonistic antibodies upregulating early activation marker CD69 and late activation marker CD25, particularly on CD4+ T cells . To confirm costimulatory activity of non-superagonistic antibodies, parallel assays should be performed with suboptimal anti-CD3 stimulation, where conventional CD28 antibodies should enhance T cell activation compared to anti-CD3 alone . Finally, to account for donor variability, these assays should be performed using PBMCs from multiple donors (at least 3-5), as differential responses have been observed across individuals . This comprehensive approach enables clear discrimination between potentially dangerous superagonistic antibodies and conventional costimulatory CD28 antibodies suitable for therapeutic development.

How should researchers design experiments to evaluate the efficacy of CD28 antibodies in combination with other immunotherapeutic agents?

When evaluating CD28 antibodies in combination with other immunotherapeutic agents, researchers should implement a systematic experimental design addressing multiple efficacy and safety parameters. For in vitro studies, tumor cell killing assays using human PBMCs as effector cells and relevant tumor cell lines as targets provide a foundational readout. The experimental setup should include: 1) target cells alone (negative control), 2) target cells with PBMCs but no antibody, 3) target cells with PBMCs and CD3 bispecific antibody alone, 4) target cells with PBMCs and CD28 antibody alone, and 5) the combination of CD3 and CD28 antibodies at various ratios . Cell killing can be monitored using flow cytometry with viability dyes or real-time impedance-based systems that provide kinetic readouts .

Beyond cytotoxicity, researchers should assess T cell proliferation (using CFSE dilution), activation marker expression (CD25, CD69), and exhaustion marker expression (PD-1, TIM-3, LAG-3) to determine if CD28 costimulation prevents early T cell dysfunction . Cytokine profiling should include both efficacy markers (IL-2, IFN-γ) and potential toxicity indicators (IL-6, TNF-α) . For in vivo studies, immunodeficient mouse models engrafted with human tumor cells and human immune components (e.g., PBMCs or CD34+ hematopoietic stem cells) allow evaluation of efficacy while monitoring for potential cytokine release syndrome . Treatment groups should include single agents and combinations, with careful dose-ranging to identify synergistic versus additive effects. These comprehensive approaches enable identification of optimal combinations that maximize anti-tumor activity while maintaining an acceptable safety profile.

How can researchers address variability in CD28 antibody experiments using primary human T cells?

Primary human T cells exhibit significant donor-to-donor variability in CD28 expression levels and functional responses, presenting challenges for consistent experimental results. To address this variability, researchers should implement several methodological approaches. First, experiments should include multiple donors (minimum 3-5, ideally more for critical studies) to capture the range of potential responses . When possible, donors should be characterized for factors known to affect T cell responsiveness, including age, sex, and prior immune exposures, as these can significantly impact CD28 expression and function .

Second, within-donor normalization can reduce variability—responses to experimental CD28 antibodies can be expressed as a percentage of the response to a standardized positive control (e.g., anti-CD3/CD28 beads or PHA stimulation) . Third, cryopreservation effects must be considered, as frozen PBMCs may exhibit altered responsiveness compared to fresh cells. When feasible, key experiments should be validated using both fresh and frozen cells from the same donors . Fourth, detailed phenotyping of T cell subsets (naive, memory, effector) should be performed, as CD28 expression varies significantly across these populations, potentially explaining differential responses .

Finally, statistical approaches should account for this biological variability—mixed effects models that incorporate donor as a random effect can provide more appropriate analysis than standard t-tests or ANOVAs. By implementing these strategies, researchers can better account for and interpret the inherent variability in CD28 antibody experiments using primary human T cells, leading to more reproducible and translatable findings.

What are the common pitfalls in epitope mapping of CD28 antibodies and how can they be overcome?

Epitope mapping of CD28 antibodies presents several challenges due to the conformational nature of many CD28 epitopes and their potential overlap with binding sites for natural ligands. One common pitfall is reliance on linear peptide arrays or truncation mutants, which often fail to capture conformational epitopes that depend on the three-dimensional structure of CD28 . To overcome this limitation, researchers should employ techniques like hydrogen-deuterium exchange mass spectrometry (HDX-MS), which can identify conformational epitopes by measuring changes in solvent accessibility upon antibody binding .

Another challenge arises from the homodimeric nature of CD28, which can complicate interpretation of binding data. Single-molecule techniques or careful analysis of binding stoichiometry can help distinguish whether an antibody binds to one or both subunits of the CD28 dimer . Epitope binning experiments using competition assays with well-characterized antibodies or natural ligands (CD80/CD86) can provide valuable information about epitope relationships but may give ambiguous results due to allosteric effects where binding at one site influences accessibility at distant sites .

How should contradictory data between in vitro and in vivo CD28 antibody studies be interpreted?

Discrepancies between in vitro and in vivo CD28 antibody studies present significant challenges for data interpretation and therapeutic development decisions. The TGN1412 experience demonstrates this problem acutely—despite promising preclinical testing, including in non-human primates, the antibody caused severe cytokine release syndrome in humans . When faced with such contradictions, researchers should systematically analyze potential sources of discrepancy.

First, species differences in CD28 expression patterns, density, and signaling pathways must be considered. While an antibody may cross-react with mouse CD28, differences in downstream signaling components or T cell subset distributions may lead to different functional outcomes . Second, the three-dimensional organization and dynamics of the immune synapse differ significantly between simplified in vitro systems and the complex in vivo environment, affecting how CD28 clustering and signaling occur . Third, the presence of regulatory feedback mechanisms in vivo that are absent in vitro can substantially modify outcomes—regulatory T cells, anti-inflammatory cytokines, and homeostatic control mechanisms may dampen responses in vivo that appear robust in vitro .

Fourth, pharmacokinetic and biodistribution factors play crucial roles in vivo but are not captured in vitro. The distribution of antibodies to lymphoid tissues and the duration of exposure significantly impact outcomes . When interpreting contradictory data, researchers should prioritize models that most closely recapitulate human physiology. Humanized mouse models, ex vivo human tissue assays, and advanced in vitro systems incorporating multiple cell types (e.g., T cells, APCs, and endothelial cells) can bridge this gap . Ultimately, conservative translation to first-in-human studies with very low starting doses represents the most prudent approach when confronted with contradictory preclinical data for novel immunomodulatory antibodies.