免疫学

Biomaterials are designed to interact with biological systems to replace, support, enhance, or monitor their function. However, there are challenges associated with traditional biomaterials’ development due to the lack of underlying theory governing cell response to materials’ chemistry. This leads to the time-consuming process of testing different materials plus the adverse reactions in the body such as cytotoxicity and foreign body response. High-throughput screening (HTS) offers a solution to these challenges by enabling rapid and simultaneous testing of a large number of materials to determine their bio-interactions and biocompatibility. Secreted proteins regulate many physiological functions and determine the success of implanted biomaterials through directing cell behaviour. However, the majority of biomaterials’ HTS platforms are suitable for microscopic analyses of cell behaviour and not for investigating non-adherent cells or measuring cell secretions. Here, we describe a multi-well platform adaptable to robotic printing of polymers and suitable for secretome profiling of both adherent and non-adherent cells. We detail the platform's development steps, encompassing the preparation of individual cell culture chambers, polymer printing, and the culture environment, as well as examples to demonstrate surface chemical characterisation and biological assessments of secreted mediators. Such platforms will no doubt facilitate the discovery of novel biomaterials and broaden their scope by adapting wider arrays of cell types and incorporating assessments of both secretome and cell-bound interactions.

Key features

• Detailed protocols for preparation of substrate for contact printing of acrylate-based polymers including O₂ plasma etching, functionalisation process, and Poly(2-hydroxyethyl methacrylate) (pHEMA) dip coating.

• Preparations of 7 mm × 7 mm polymers employing pin printing system.

• Provision of confined area for each polymer using ProPlate^® multi-well chambers.

• Compatibility of this platform was validated using adherent cells [primary human monocyte–derived macrophages (MDMs)) and non-adherent cells (primary human monocyte–derived dendritic cells (moDCs)].

• Examples of the adaptability of the platform for secretome analysis including five different cytokines using enzyme-linked immunosorbent assay (ELISA, DuoSet^®).

Graphical overview

CD8⁺CD28⁻ T suppressor cells (Ts) have been documented to promote immune tolerance by suppressing effector T cell responses to alloantigens following transplantation. The suppressive function of T cells has been defined as the inhibitory effect of Ts on the proliferation rate of effector T cells. ³H-thymidine is a classical immunological technique for assaying T cell proliferation but this approach has drawbacks such as the inconvenience of working with radioactive materials. Labeling T cells with CFSE allows relatively easy tracking of generations of proliferated cells. In this report, we utilized antigen presenting cells (APCs) and T cells matched for human leukocyte antigen (HLA) class I or class II to study CD8⁺CD28^- T cell suppression generated in vitro by this novel approach of combining allogeneic APCs and γc cytokines. The expanded CD8⁺CD28^- T cells were isolated (purity 95%) and evaluated for their suppressive capacity in mixed lymphocyte reactions using CD4⁺ T cells as responders. Here, we present our adapted protocol for assaying the Ts allospecific suppression of CFSE-labeled responder T cells.

Functional and mechanistic studies of CD4⁺ T cell lineages rely on robust methods of in vitro T cell polarization. Here, we report an optimized protocol for in vitro differentiation of a mouse non-pathogenic T helper 17 (T_H17) cell lineage. Most of the previously established protocols require irradiated splenocytes as artificial antigen presenting cells (APC) for TCR activation. The protocol described here employs plate-bound antibodies and a T_H17-polarizing cytokine cocktail to activate and differentiate naïve CD4⁺ T (T_nai) cells, reflecting a simple and robust protocol for in vitro T_H17n differentiation. Using T cells that are genetically engineered with an IL-17 reporter, this protocol may enable the rapid production of a pure population of IL17-expressing CD4⁺ T cells for system biology studies and high-throughput functional screening.

Activating the STING (stimulator of interferon genes) signaling pathway via administration of STING agonist cyclic GMP-AMP (cGAMP) has shown great promise in cancer immunotherapy. While state-of-the-art approaches have predominantly focused on the encapsulation of cGAMP into liposomes or polymersomes for cellular delivery, we discovered that the recombinant STING protein lacking the transmembrane domain (STINGΔTM) could be used as a functional carrier for cGAMP delivery and elicit type I IFN expression in STING-deficient cell lines. Using this approach, we generated anti-tumoral immunity in mouse melanoma and colon cancer models, providing a potential translatable platform for STING agonist-based immunotherapy. Here, we report the detailed in vitro STING activation protocols with cGAMP-STINGΔTM complex to assist researchers in further development of this approach. This protocol can also be easily expanded to other applications related to STING activation, such as control of various types of infections.

For many infectious diseases T cells are an important part of naturally acquired protective immune responses, and inducing these by vaccination has been the aim of much research. Here, we describe a protocol for the analysis of vaccine-induced antigen-specific immune responses. For this purpose, cells of whole spleens obtained from vaccinated BALB/c mice were ex vivo stimulated with the antigen incorporated in the vaccine. Evaluation and characterization of vaccine-induced adaptive T cell responses was performed by assaying spleen cell proliferation through radioactive ³[H]-thymidine incorporation and multiplex cytokine analysis of IL-2, IFN-γ and TNFα in supernatants from spleen cell suspensions. This protocol can be very useful as a starting point for assessing vaccine-induced memory T cell populations in pre-clinical studies.

We describe here a method to visualize concentration fields of cytokines around cytokine-secreting cells. The main challenge is that physiological cytokine concentrations can be very low, in the pico-molar range. Since it is currently impossible to measure such concentrations directly, we rely on cell’s response to the cytokines–the phosphorylation of a transcription factor–that can be visualized through antibody staining. Our devices aim at mimicking conditions in dense tissues, such as lymph nodes. A small number of secreting cells is deposited on a polylysine-coated glass and covered by multiple layers of cytokine-consuming. The cells are left to communicate for 1 h, after which the top layers are removed and the bottom layer of cells is antibody labeled for the response to cytokines. Then a cross-section of cytokine fields can be visualized by standard fluorescence microscopy. This manuscript summarized our method to quantify the extent of cytokine-mediated cell-to-cell communications in dense collection of cells in vitro.

Long-lived T-cell–mediated immunity requires persistence of memory T cells in an antigen-free environment while also maintaining a heightened capacity to recall effector functions. Such antigen-independent homeostatic proliferation is mediated in part by the common gamma-chain cytokines IL-7 and IL-15. To further explore the mechanisms governing maintenance of effector functions in long-lived memory T cells during antigen-independent proliferation, human naïve and memory CD8 T cells can be sorted from peripheral blood mononuclear cells (PBMCs), labeled with the proliferation-tracking dye carboxyfluorescein succinimidyl ester (CFSE), and then purified based on their levels of cell division. This allows investigators to assess differences in the desired molecular target in cells that have undergone cytokine-driven proliferation. We provide here a protocol for assessing epigenetic programs in divided and undivided human naïve and memory CD8 T cells following 7 days in culture with IL-7 and IL-15 to illustrate how this approach can shed light on the mechanism(s) that governs the preservation of effector functions during homeostasis of long-lived memory CD8 T cells.

Interferon-gamma (IFN-γ) is crucial for immunity against intracellular pathogens and for tumor control. It is produced predominantly by natural killer (NK) and natural killer T cells (NKT) as well as by antigen-specific Th1 CD4⁺ and CD8⁺ effector T cells. When investigating immune responses against pathogens and cancer cells, measuring antigen-specific cytokine-responses by cells of adaptive immunity offers an advantage over total non-specific cytokine responses. Significantly, the measurement of antigen-specific IFN-γ responses against pathogens or cancer cells, when compared to a treatment group, provides a quantitative measure of how well the treatment works. Measuring antigen-specific IFN-γ responses involves culture of the cells being considered (CD4⁺ or CD8⁺ T cells) with antigen presenting cells (APC) and a specific peptide from the target pathogen or cancer cell compared to control cultures without a peptide. After a suitable timeframe, the cytokine released is measured by an ELISPOT assay. The difference in the number of cells secreting IFN-γ, with and without peptide, is a measure of antigen-specific IFN-γ responses. This assay can be applied to other cytokines such as IL-10.

Production of cytokines plays an important role in the immune response. Cytokines are involved in many different pathways including the induction of many anti-viral proteins by IFN gamma, the induction of T cell proliferation by IL-2 and the inhibition of viral gene expression and replication by TNF alpha. Cytokines are not preformed factors but are rapidly produced and secreted in response to cellular activation. Intracellular cytokine detection by flow cytometry has emerged as the premier technique for studying cytokine production at the single-cell level. It detects the production and accumulation of cytokines within the endoplasmic reticulum after cell stimulation, allowing direct TH1 versus TH2 determination. It can also be used in combination with other flow cytometry protocols for immunophenotyping using cell surface markers or with MHC multimers to detect an antigen specific response, making it an extremely flexible and versatile method. This capability, combined with the high throughput nature of the instrumentation, gives intracellular cytokine staining an enormous advantage over existing single-cell techniques such as ELISPOT, limiting dilution, and T cell cloning. The principle steps of intracellular cytokine staining is as follows:
1. Cells are activated for a few hours using either a specific peptide or a non-specific activation cocktail;
2. An inhibitor of protein transport (e.g. Brefeldin A) is added to retain the cytokines within the cell;
3. Next, EDTA is added to remove adherent cells from the activation vessel;
4. After washing, antibodies to cell surface markers can be added to the cells;
5. The cells are then fixed in paraformaldehyde and permeabilized;
6. The anti-cytokine antibody is added and the cells can be analyzed by flow cytometer.

In this protocol, we use a CyTOF^TM mass cytometry to collect single-cell data on a large number of cytokines/chemokines as well as cell-surface proteins that characterize T cells and other immune cells. The current selected mass window in AW 103-203 includes the lanthanides used for most antibody labeling, along with iridium and rhodium for DNA intercalators. The output data are in the format as .txt and .fcs files, which is compatible with many analysis programs. This protocol could be adapted to include tetramers into the staining panel, but we have not optimized for that purpose.

The principal steps of intracellular cytokine staining are as follows: First, cells are activated for a few hours using either a specific peptide or a non-specific activation cocktail. An inhibitor of protein transport (e.g. Brefeldin A) is added to retain the cytokines within the cell. Next, EDTA is added to remove adherent cells from the activation vessel. After washing, antibodies to cell surface markers are added to the cells. The cells are then fixed in paraformaldehyde and permeabilized. We use a gentle detergent, saponin, as the permealization buffer because it is less destructive to surface and intracellular epitopes compared to harsh detergents or methanol. After permeabilization, the metal-conjugated anti-cytokine antibodies are added into the cell suspension. The stained cells are then sequentially introduced into the mass cytometry for signal intensity analysis.