Phase Ib open-label, multicenter study of pixatimod, an activator of TLR9, in combination with nivolumab in subjects with microsatellite-stable metastatic colorectal cancer, metastatic pancreatic ductal adenocarcinoma and other solid tumors.

BACKGROUND: Pixatimod is a unique activator of the Toll-like Receptor 9 pathway. This phase I trial evaluated safety, efficacy and pharmacodynamics of pixatimod and PD-1 inhibitor nivolumab in immunologically cold cancers. METHODS: 3+3 dose escalation with microsatellite stable metastatic colorectal cancer (MSS mCRC) and metastatic pancreatic ductal adenocarcinoma (mPDAC) expansion cohorts. Participants received pixatimod once weekly as a 1-hour intravenous infusion plus nivolumab every 2 weeks. Objectives included assessment of safety, antitumor activity, pharmacodynamics, and pharmacokinetic profile. RESULTS: Fifty-eight participants started treatment. The maximum tolerated dose of pixatimod was 25 mg in combination with 240 mg nivolumab, which was used in the expansion phases of the study. Twenty-one grade 3-5 treatment-related adverse events were reported in 12 participants (21%); one participant receiving 50 mg pixatimod/nivolumab had a treatment-related grade 5 AE. The grade 3/4 rate in the MSS mCRC cohort (n=33) was 12%. There were no responders in the mPDAC cohort (n=18). In the MSS mCRC cohort, 25 participants were evaluable (initial postbaseline assessment scans >6 weeks); of these, three participants had confirmed partial responses (PR) and eight had stable disease (SD) for at least 9 weeks. Clinical benefit (PR+SD) was associated with lower Pan-Immune-Inflammation Value and plasma IL-6 but increased IP-10 and IP-10/IL-8 ratio. In an MSS mCRC participant with PR as best response, increased infiltration of T cells, dendritic cells, and to a lesser extent NK cells, were evident 5 weeks post-treatment. CONCLUSIONS: Pixatimod is well tolerated at 25 mg in combination with nivolumab. The efficacy signal and pharmacodynamic changes in MSS mCRC warrants further investigation. TRIAL REGISTRATION NUMBER: NCT05061017.

Immune contexture analysis in immuno-oncology: applications and challenges of multiplex fluorescent immunohistochemistry.

The tumor microenvironment is an integral player in cancer initiation, tumor progression, response and resistance to anti-cancer therapy. Understanding the complex interactions of tumor immune architecture (referred to as 'immune contexture') has therefore become increasingly desirable to guide our approach to patient selection, clinical trial design, combination therapies, and patient management. Quantitative image analysis based on multiplexed fluorescence immunohistochemistry and deep learning technologies are rapidly developing to enable researchers to interrogate complex information from the tumor microenvironment and find predictive insights into treatment response. Herein, we discuss current developments in multiplexed fluorescence immunohistochemistry for immune contexture analysis, and their application in immuno-oncology, and discuss challenges to effectively use this technology in clinical settings. We also present a multiplexed image analysis workflow to analyse fluorescence multiplexed stained tumor sections using the Vectra Automated Digital Pathology System together with FCS express flow cytometry software. The benefit of this strategy is that the spectral unmixing accurately generates and analyses complex arrays of multiple biomarkers, which can be helpful for diagnosis, risk stratification, and guiding clinical management of oncology patients.

Human peripheral blood DNAM-1neg NK cells are a terminally differentiated subset with limited effector functions.

Natural killer (NK) cells are a heterogeneous population of innate lymphocytes whose potent anticancer properties make them ideal candidates for cellular therapeutic application. However, our lack of understanding of the role of NK cell diversity in antitumor responses has hindered advances in this area. In this study, we describe a new CD56dim NK cell subset characterized by the lack of expression of DNAX accessory molecule-1 (DNAM-1). Compared with CD56bright and CD56dimDNAM-1pos NK cell subsets, CD56dimDNAM-1neg NK cells displayed reduced motility, poor proliferation, lower production of interferon-γ, and limited killing capacities. Soluble factors secreted by CD56dimDNAM-1neg NK cells impaired CD56dimDNAM-1pos NK cell-mediated killing, indicating a potential inhibitory role for the CD56dimDNAM-1neg NK cell subset. Transcriptome analysis revealed that CD56dimDNAM-1neg NK cells constitute a new mature NK cell subset with a specific gene signature. Upon in vitro cytokine stimulation, CD56dimDNAM-1neg NK cells were found to differentiate from CD56dimDNAM-1pos NK cells. Finally, we report a dysregulation of NK cell subsets in the blood of patients diagnosed with Hodgkin lymphoma and diffuse large B-cell lymphoma, characterized by decreased CD56dimDNAM-1pos/CD56dimDNAM-1neg NK cell ratios and reduced cytotoxic activity of CD56dimDNAM-1pos NK cells. Altogether, our data offer a better understanding of human peripheral blood NK cell populations and have important clinical implications for the design of NK cell-targeting therapies.

Human papillomavirus E7 oncoprotein expression by keratinocytes alters the cytotoxic mechanisms used by CD8 T cells.

Cervical cancer is a malignant transformation of keratinocytes initiated by the E7 oncoprotein of human papillomavirus (HPV). These tumors are characterized by keratinocyte hyperproliferation and are often infiltrated with activated CD8 T cells. HPV infection confers changes to gain immunological advantage to promote chronic infection, and these persist with malignant transformation. We investigated the relative importance of the many redundant mechanisms of cytotoxicity used by CD8 T cells to kill keratinocytes expressing HPV E7 oncoprotein using extended-duration time-lapse microscopy that allows examination of cell-to-cell interactions during killing. E7 expression by keratinocytes increased susceptibility to cell-mediated killing. However, while killing of non-transgenic keratinocytes was traditional, perforin-mediated, and caspase-dependent, E7-expression favored killing by perforin-independent, caspase-independent mechanisms. The roles of perforin, TNFα, IFNγ, Fas/FasL and PD1/PD-L1 were graded according to target cell survival to produce a hierarchy of killing mechanisms utilized in killing E7-expressing cells. TNFα was essential for perforin-mediated killing of E7-expressing cells, but not perforin-independent killing. IFNγ facilitated killing by Fas/FasL interaction, especially in the absence of perforin. Additionally, expression of E7 offered protection from killing by up regulation of PD-L1, Fas and FasL expression on keratinocytes promoting fight-back by target cells, resulting in effector cell death. This study shows that keratinocytes expressing E7 are highly susceptible to killing by CD8 T cells, but utilizing different armamentarium. Down-regulation of CD8 T cell cytotoxicity in HPV-related tumors may be due to suppression by E7-expressing keratinocytes. Immunotherapy for HPV-related cancers may be improved by suppression of PD-L1, or by suppression of FasL.

Interferon-γ derived from cytotoxic lymphocytes directly enhances their motility and cytotoxicity.

Interferon gamma (IFNγ) is a key moderator of cell-mediated immunity with diverse, mainly pro-inflammatory actions on immunocytes and target tissue. Recent studies have shown it may enhance anti-tumor and antiviral effects of CD8 T cells. Here we investigate the mechanisms by which IFNγ mediates CD8 T-cell cytotoxic function. We show that in vivo, antigen-specific CD8 T cells that produce INFγ are necessary to effect rejection of skin grafts expressing OVA as a transgene in keratinocytes. The ability of CD8 T cells to produce IFNγ enhanced their ability to migrate to the site of antigen-presenting skin cells. By in vivo imaging, we show that CTL motility, particularly speed, during graft rejection was enhanced by locally available IFNγ. We then used a reductionist two-cell model of CTL effectors and keratinocyte targets to investigate the effects of locally available (paracrine) and CTL-producing (autocrine) IFNγ on the motility behavior and killing ability of the CTL. Using live-cell imaging by prolonged time-lapse microscopy of primary effector CD8 T cells and antigen-expressing primary keratinocyte targets, we show that CD8 T-cell cytotoxic function and motility is enhanced by locally available IFNγ. Conversely, deprivation of either autocrine or paracrine IFNγ, or blockade of IFNγ signaling to CTL markedly reduced their cytotoxic function, their kinematics, and effector cell survival. We conclude that in vitro and in vivo, autocrine production of IFNγ by CTL enhances their motility and promotes killing of primary target keratinocytes. The absolute need for local IFNγ to enable cytotoxic CD8 T-cell function is of significance for immunotherapy for chronic viral infection and for cancer.

Analysis of Cytochrome c Release by Immunocytochemistry.

Cytochrome c is normally localized between the inner and outer membranes of mitochondria in healthy cells. However, during apoptosis, it is released into the cytoplasm, where it binds to apoptotic protease activating factor. Caspase-9 is then recruited and activated by this complex in a process known as the induced proximity model. Release of cytochrome c from mitochondria is therefore a critical event in apoptosis and various protocols are available for its measurement. Cytochrome c in mitochondria has a punctate localization pattern in the cell and its translocation to the cytoplasm results in a diffuse distribution. This is visually striking and easily observed by immunocytochemistry. This protocol describes the use of immunocytochemistry to assay cytochrome c release during apoptosis.

Measuring Mitochondrial Transmembrane Potential by TMRE Staining.

Adenosine triphosphate (ATP) is the main source of energy for metabolism. Mitochondria provide the majority of this ATP by a process known as oxidative phosphorylation. This process involves active transfer of positively charged protons across the mitochondrial inner membrane resulting in a net internal negative charge, known as the mitochondrial transmembrane potential (ΔΨm). The proton gradient is then used by ATP synthase to produce ATP by fusing adenosine diphosphate and free phosphate. The net negative charge across a healthy mitochondrion is maintained at approximately -180 mV, which can be detected by staining cells with positively charged dyes such as tetramethylrhodamine ethyl ester (TMRE). TMRE emits a red fluorescence that can be detected by flow cytometry or fluorescence microscopy and the level of TMRE fluorescence in stained cells can be used to determine whether mitochondria in a cell have high or low ΔΨm. Cytochrome c is essential for producing ΔΨm because it promotes the pumping the protons into the mitochondrial intermembrane space as it shuttles electrons from Complex III to Complex IV along the electron transport chain. Cytochrome c is released from the mitochondrial intermembrane space into the cytosol during apoptosis. This impairs its ability to shuttle electrons between Complex III and Complex IV and results in rapid dissipation of ΔΨm. Loss of ΔΨm is therefore closely associated with cytochrome c release during apoptosis and is often used as a surrogate marker for cytochrome c release in cells.

Dead Cert: Measuring Cell Death.

Many cells in the body die at specific times to facilitate healthy development or because they have become old, damaged, or infected. Defects in cells that result in their inappropriate survival or untimely death can negatively impact development or contribute to a variety of human pathologies, including cancer, AIDS, autoimmune disorders, and chronic infection. Cell death may also occur following exposure to environmental toxins or cytotoxic chemicals. Although this is often harmful, it can be beneficial in some cases, such as in the treatment of cancer. The ability to objectively measure cell death in a laboratory setting is therefore essential to understanding and investigating the causes and treatments of many human diseases and disorders. Often, it is sufficient to know the extent of cell death in a sample; however, the mechanism of death may also have implications for disease progression, treatment, and the outcomes of experimental investigations. There are a myriad of assays available for measuring the known forms of cell death, including apoptosis, necrosis, autophagy, necroptosis, anoikis, and pyroptosis. Here, we introduce a range of assays for measuring cell death in cultured cells, and we outline basic techniques for distinguishing healthy cells from apoptotic or necrotic cells-the two most common forms of cell death. We also provide personal insight into where these assays may be useful and how they may or may not be used to distinguish apoptotic cell death from other death modalities.

Quantitation of Apoptosis and Necrosis by Annexin V Binding, Propidium Iodide Uptake, and Flow Cytometry.

The surface of healthy cells is composed of lipids that are asymmetrically distributed on the inner and outer leaflet of the plasma membrane. One of these lipids, phosphatidylserine (PS), is normally restricted to the inner leaflet of the plasma membrane and is, therefore, only exposed to the cell cytoplasm. However, during apoptosis lipid asymmetry is lost and PS becomes exposed on the outer leaflet of the plasma membrane. Annexin V, a 36-kDa calcium-binding protein, binds to PS; therefore, fluorescently labeled Annexin V can be used to detect PS that is exposed on the outside of apoptotic cells. Annexin V can also stain necrotic cells because these cells have ruptured membranes that permit Annexin V to access the entire plasma membrane. However, apoptotic cells can be distinguished from necrotic cells by co-staining with propidium iodide (PI) because PI enters necrotic cells but is excluded from apoptotic cells. This protocol describes Annexin V binding and PI uptake followed by flow cytometry to detect and quantify apoptotic and necrotic cells.

Detecting Cleaved Caspase-3 in Apoptotic Cells by Flow Cytometry.

Apoptosis is orchestrated by caspases, a family of cysteine proteases that cleave their substrates on the carboxy-terminal side of specific aspartic acid residues. These proteases are generally present in healthy cells as inactive zymogens, but when stimulated they undergo autolytic cleavage to become fully active. They subsequently cleave their substrates at one or two specific sites, which can result in activation, inactivation, relocalization, or remodeling of the substrate. Consequently, many of the cleaved fragments remain intact during apoptosis and can be detected using substrate-specific antibodies. These fragments are most commonly detected by western blotting, which resolves proteins and their fragments based on molecular mass. However, antibodies that only recognize cleaved fragments can be used to specifically label cells in which caspase cleavage has occurred. It is then possible to quantify these cells by flow cytometry. A number of antibodies that specifically recognize caspase-cleaved fragments have been generated, including antibodies that recognize the cleaved form of caspase-3. This caspase is responsible for the majority of proteolysis during apoptosis, and detection of cleaved caspase-3 is therefore considered a reliable marker for cells that are dying, or have died by apoptosis. This protocol outlines the quantification of apoptosis by flow cytometric detection of cleaved caspase-3.

Detection of DNA Fragmentation in Apoptotic Cells by TUNEL.

Degradation of DNA into oligonucleosomal-sized fragments is a unique event in apoptosis that is orchestrated by caspase-activated DNase. Traditionally, this event is observed by resolving cellular DNA by gel electrophoresis, which results in a characteristic "ladder" pattern. However, this technique is time-consuming and cannot be used to quantitate the number of apoptotic cells in a sample. Terminal dUTP nick-end labeling (TUNEL) of fragmented DNA allows researchers to identify DNA fragmentation at the single-cell level. This method involves the specific addition of fluorescently labeled UTP to the 3'-end of the DNA fragments by terminal deoxynucleotidyl transferase. The TUNEL assay is both fast and sensitive. Here, we describe a protocol in which cells are treated with TUNEL reagent and counterstained with Hoechst 33342. In contrast to TUNEL, which only stains apoptotic cells, Hoechst 33342 stains the DNA of all cells.

Measuring the DNA Content of Cells in Apoptosis and at Different Cell-Cycle Stages by Propidium Iodide Staining and Flow Cytometry.

All cells are created from preexisting cells. This involves complete duplication of the parent cell to create two daughter cells by a process known as the cell cycle. For this process to be successful, the DNA of the parent cell must be faithfully replicated so that each daughter cell receives a full copy of the genetic information. During the cell cycle, the DNA content of the parent cell increases as new DNA is synthesized (S phase). When there are two full copies of the DNA (G2/M phase), the cell splits to form two new cells (G0/G1 phase). As such, cells in different stages of the cell cycle have different DNA contents. The cell cycle is tightly regulated to safeguard the integrity of the cell and any cell that is defective or unable to complete the cell cycle is programmed to die by apoptosis. When this occurs, the DNA is fragmented into oligonucleosomal-sized fragments that are disposed of when the dead cell is removed by phagocytosis. Consequently apoptotic cells have reduced DNA content compared with living cells. This can be measured by staining cells with propidium iodide (PI), a fluorescent molecule that intercalates with DNA at a specific ratio. The level of PI fluorescence in a cell is, therefore, directly proportional to the DNA content of that cell. This protocol describes the use of PI staining to determine the percentage of cells in each phase of the cell cycle and the percentage of apoptotic cells in a sample.

Morphological Analysis of Cell Death by Cytospinning Followed by Rapid Staining.

Identifying and characterizing different forms of cell death can be facilitated by staining internal cellular structures with dyes such as hematoxylin and eosin (H&E). These dyes stain the nucleus and cytoplasm, respectively, and optimized reagents (e.g., Rapi-Diff, Rapid Stain, or Quick Dip) are commonly used in pathology laboratories. Fixing and staining adherent cells with these optimized reagents is a straightforward procedure, but apoptotic cells may detach from the culture plate and be washed away during the fixing and staining procedure. To prevent the loss of apoptotic cells, cells can be gently centrifuged onto glass slides by cytospinning before fixing and staining. In addition to apoptotic cells, this procedure can be used on cells in suspension, or adherent cells that have been trypsinized and removed from the culture dish. This protocol describes cytospinning followed by Rapi-Diff staining for morphological analysis of cell death.

Analyzing Cell Death by Nuclear Staining with Hoechst 33342.

The nuclei of healthy cells are generally spherical, and the DNA is evenly distributed. During apoptosis the DNA becomes condensed, but this process does not occur during necrosis. Nuclear condensation can therefore be used to distinguish apoptotic cells from healthy cells or necrotic cells. Dyes that bind to DNA, such as Hoechst 33342 or 4',6-diamidino-2-phenylindole (DAPI), can be used to observe nuclear condensation. These dyes fluoresce at 461 nm when excited by ultraviolet light and can therefore be visualized using conventional fluorescent microscopes equipped with light sources that emit light at ∼350 nm and filter sets that permit the transmission of light at ∼460 nm. This protocol describes staining and visualization of cells stained with Hoechst 33342, but it can be adapted for staining with DAPI or other dyes.

Measuring Survival of Adherent Cells with the Colony-Forming Assay.

Measuring cell death with colorimetric or fluorimetric dyes such as trypan blue and propidium iodide (PI) can provide an accurate measure of the number of dead cells in a population at a specific time; however, these assays cannot be used to distinguish cells that are dying or marked for future death. In many cases it is essential to measure the proliferative capacity of treated cells to provide an indirect measurement of cell death. This can be achieved using the colony-forming assay described here. This protocol specifically applies to measurement of HeLa cells but can be used for most adherent cell lines with limited motility.

Measuring Survival of Hematopoietic Cancer Cells with the Colony-Forming Assay in Soft Agar.

Colony-forming assays measure the ability of cells in culture to grow and divide into groups. Any cell that has the potential to form a colony may also have the potential to cause cancer or relapse in vivo. Colony-forming assays also provide an indirect measurement of cell death because any cell that is dead or dying will not continue to proliferate. The proliferative capacity of adherent cells such as fibroblasts can be determined by growing cells at low density on culture dishes and counting the number of distinct groups that form over time. Cells that grow in suspension, such as hematopoietic cells, cannot be assayed this way because the cells move freely in the media. Assays to determine the colony-forming ability of hematopoietic cells must therefore be performed in solid matrices that restrict large-scale movement of the cells. One such matrix is soft agar. This protocol describes the use of soft agar to compare the colony-forming ability of untreated hematopoietic cells to the colony-forming ability of hematopoietic cells that have been treated with a cytotoxic agent.

Triggering Death of Adherent Cells with Ultraviolet Radiation.

Ultraviolet (UV) radiation is a convenient stimulus for triggering cell death that is available in most laboratories. We use a Stratalinker UV cross-linker because it is a safe, cheap, reliable, consistent, and easily controlled source of UV irradiation. This protocol describes using a Stratalinker to trigger UV-induced death of HeLa cells.

Triggering Apoptosis in Hematopoietic Cells with Cytotoxic Drugs.

Cytotoxic agents are commonly added to cultured cells in the laboratory to investigate their efficacy, mechanism of action, and therapeutic potential. Most of these agents trigger cell death by apoptosis, which is also the most common form of cell death during development, aging, homeostasis, and eradication of disease. Treatment of cells with cytotoxic agents is therefore useful for investigating basic mechanisms of cell death in the human body. Actinomycin D, a cytotoxic agent isolated from Streptomyces, induces apoptosis in a variety of cell lines including the histiocytic lymphoma cell line U937. Treatment of U937 cells with actinomycin D provides an ideal model of drug-induced apoptosis that can also be used as a positive control for comparison with other treatments.

Measuring Cell Death by Trypan Blue Uptake and Light Microscopy.

Trypan blue is a colorimetric dye that stains dead cells with a blue color easily observed using light microscopy at low resolution. The staining procedure is rapid and cells can be analyzed within minutes. The number of live (unstained) and dead (blue) cells can be counted using a hemocytometer on a basic upright microscope. Trypan blue staining is therefore a convenient assay for rapidly determining the overall viability of cells in a culture before commencing scientific experimentation, or for quantitating cell death following treatment with any cytotoxic stimuli.

Measuring Cell Death by Propidium Iodide Uptake and Flow Cytometry.

Propidium iodide (PI) is a small fluorescent molecule that binds to DNA but cannot passively traverse into cells that possess an intact plasma membrane. PI uptake versus exclusion can be used to discriminate dead cells, in which plasma membranes become permeable regardless of the mechanism of death, from live cells with intact membranes. PI is excited by wavelengths between 400 and 600 nm and emits light between 600 and 700 nm, and is therefore compatible with lasers and photodetectors commonly available in flow cytometers. This protocol for PI staining can be used to quantitate cell death in most modern research facilities and universities.