Taurolidine-d₆

Infectious-disease experiments test how pathogens interact with host barriers, innate immune receptors, inflammatory signaling, pathogen replication, and tissue injury; pattern-recognition receptors such as TLRs, RIG-I-like receptors, NOD-like receptors, and inflammasomes detect microbial molecules and activate NF-κB, interferon, and cytokine responses. The central hypothesis is that infection severity reflects the balance between pathogen burden and host response: protective inflammation restricts pathogen growth, whereas excessive or mislocalized inflammation contributes to tissue damage and disease phenotype. Unresolved questions include which host pathways are protective versus pathogenic, why some infection models fail to translate to human disease, and which combined readouts best predict clinically relevant infection outcomes.

Annexin V-based apoptosis assays rely on the detection of phosphatidylserine (PS) externalization from the inner leaflet of the plasma membrane to the outer leaflet, an early biochemical hallmark of apoptosis. Fluorescently labeled Annexin V binds PS in a calcium-dependent manner, enabling identification of early apoptotic cells by flow cytometry or fluorescence microscopy. When combined with a membrane-impermeant DNA-binding dye (e. g. , propidium iodide), this approach allows discrimination between viable (Annexin V−/dye−), early apoptotic (Annexin V+/dye−), and late apoptotic or necrotic (Annexin V+/dye+) cell populations by assessing membrane integrity and PS exposure.

Bromodeoxyuridine (BrdU) incorporation assay is based on the principle that BrdU, a thymidine analog, is incorporated into newly synthesized DNA during the S phase of the cell cycle, thereby serving as a marker of DNA replication and cellular proliferation. Incorporated BrdU can be detected using anti-BrdU antibodies following DNA denaturation, enabling visualization or quantification of proliferating cells through immunochemical detection methods such as immunofluorescence or immunohistochemistry.

A method of simulating the in vivo environment in vitro to maintain the cell growth, differentation and main functions.

The CFSE (carboxyfluorescein diacetate succinimidyl ester) dye dilution proliferation assay is based on the covalent labeling of intracellular proteins by a cell-permeant fluorescent dye that becomes fluorescent upon intracellular ester cleavage and then is stably retained within cells. As labeled cells divide, the dye is partitioned equally between daughter cells, resulting in a stepwise halving of fluorescence intensity that can be quantified by flow cytometry to determine the number of cell divisions undergone by each cell population. This fluorescence dilution approach enables quantitative tracking of lymphocyte proliferation at the single-cell level over multiple rounds of division. CFSE-based proliferation analysis has been widely applied to measure antigen-driven lymphocyte expansion in vitro, where discrete fluorescence peaks correspond to successive cell divisions and allow reconstruction of proliferative history within heterogeneous populations.

Dye-dilution cell tracking assays quantify cell proliferation by covalently labeling intracellular proteins with a stable fluorescent dye that is equally partitioned between daughter cells during mitosis, resulting in stepwise halving of fluorescence intensity with each cell division as measured by flow cytometry histograms. Carboxyfluorescein diacetate succinimidyl ester (CFSE) is a prototypical dye that diffuses into cells, is enzymatically converted into a fluorescent compound, and then covalently binds intracellular amine groups, producing long-lived fluorescence suitable for tracking multiple rounds of division in vitro and in vivo. Successive generations of dividing cells form discrete peaks of decreasing fluorescence intensity, enabling estimation of proliferation history, precursor frequency, and division index within heterogeneous populations. Alternative dyes such as CellTrace Violet (CTV) and far-red membrane dyes (e. g. , PKH26) follow the same dilution principle but differ

The CCK-8/WST-8 assay is based on the reduction of the water-soluble tetrazolium salt WST-8 to a water-soluble formazan product by cellular dehydrogenases in metabolically active cells, where the generated formazan amount is proportional to the number of living cells and is quantified by measuring absorbance in the visible range, providing a colorimetric readout for cell viability and proliferation assessment. This class of tetrazolium-based assays improves upon earlier MTT-based systems by producing a water-soluble formazan, eliminating the need for organic solubilization steps and enabling direct spectrophotometric measurement in culture medium.

Cell-cycle analysis by flow cytometry measures DNA content in single cells to estimate the fraction of cells in G0/G1, S, and G2/M phases. Propidium iodide intercalates into DNA, and after RNA removal with RNase, fluorescence intensity reflects cellular DNA content: 2N cells are assigned to G0/G1, cells between 2N and 4N to S phase, and 4N cells to G2/M. DNA-content analysis alone cannot reliably separate G0 from G1 or G2 from M. Ki-67 can distinguish quiescent G0 cells from cycling cells, EdU or BrdU incorporation marks active DNA synthesis in S phase, and phospho-histone H3 staining identifies mitotic cells within the 4N population.

Cell counting-based growth curve assays quantify cell proliferation by directly measuring changes in viable cell number over time using manual or automated counting methods such as hemocytometer-based counting or instrument-assisted cell enumeration, enabling construction of growth curves that reflect population expansion dynamics in response to culture conditions. A widely used approach is trypan blue exclusion with hemocytometer counting, where membrane-compromised (non-viable) cells take up the dye, allowing discrimination between viable and non-viable cells while simultaneously enabling total cell number quantification. Repeated sampling across time points allows estimation of proliferation rate, growth phases, and comparative growth kinetics between experimental conditions.

The clonogenic (colony formation) assay measures the ability of a single cell to retain reproductive viability and form a macroscopic colony, typically defined as a cluster derived from one progenitor cell after a defined growth period. This assay is widely used to evaluate cell survival after exposure to ionizing radiation or cytotoxic treatments and is considered a standard method in radiation biology for generating dose-response relationships of reproductive cell death. Colony formation reflects long-term proliferative capacity rather than short-term metabolic activity, and survival is quantified by comparing treated versus untreated conditions based on colony number and derived survival fractions.

The following protocol uses the MTT colorimetric assay as a classic literature-established method for assessing cell viability/metabolic activity in cultured mammalian cells. MTT[3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] is reduced by metabolically active cells to a colored formazan product; the amount of formazan is quantified spectrophotometrically and provides an indirect measure of metabolically active viable cells. Importantly, MTT reduction reflects cellular oxidoreductase/metabolic activity rather than an absolute direct count of living cells, so changes in cellular metabolism can alter the signal independently of cell number.

The EdU incorporation assay measures DNA synthesis by adding the thymidine analog 5-ethynyl-2′-deoxyuridine to cells or tissues, where it is incorporated into newly synthesized DNA during S phase. Incorporated EdU is detected by copper-catalyzed azide-alkyne cycloaddition, in which a fluorescent azide covalently reacts with the ethynyl group on EdU, allowing S-phase cells to be detected by fluorescence microscopy, flow cytometry, or high-content imaging. EdU detection does not require DNA denaturation or anti-BrdU antibody access, which preserves sample structure and improves compatibility with immunostaining and multiparameter cytometry compared with BrdU-based detection. EdU can be cytotoxic in a cell-type- and exposure-dependent manner, so pulse duration, concentration, and continuous-labeling designs should be validated for each cell type.

Ki-67 immunostaining measures the growth fraction of a cell population by detecting Ki-67, a nuclear antigen present in proliferating cells and absent in quiescent G0 cells. The readout is the percentage of Ki-67-positive nuclei among total counted cells, commonly called the Ki-67 labeling index or proliferation index.

Apoptosis, also called programmed cell death, is generally characterized by distinct morphological characteristics.

TUNEL staining detects DNA strand breaks by using terminal deoxynucleotidyl transferase to add labeled nucleotides to exposed 3′-OH DNA termini, generating either microscopic staining in fixed cells or tissue sections, or fluorescence/cytometric signal in cell suspensions. TUNEL positivity reflects DNA fragmentation but should not be interpreted alone as definitive apoptosis, because TUNEL can also label necrotic, autolytic, mechanically damaged, or DNA-repair-associated DNA breaks.

PCNA immunodetection measures proliferative activity by detecting proliferating cell nuclear antigen, a nuclear protein associated with DNA polymerase δ function and DNA replication. The assay readout is the proportion of PCNA-positive nuclei among total counted cells, but PCNA labeling is not identical to BrdU labeling because PCNA can mark late G1/early S-associated replication competence and may persist beyond active DNA synthesis depending on fixation and extraction conditions.

Lipopolysaccharide (LPS)-induced endotoxemia is a widely used in vivo model of acute systemic inflammation in which LPS, a Gram-negative bacterial endotoxin, activates innate immune signaling primarily through TLR4, leading to rapid and transient induction of pro-inflammatory cytokines such as TNF-α, IL-6, and IL-1β in circulation and tissues. This cytokine surge is commonly used as a measurable readout of systemic inflammatory activation and immune dysregulation, and is typically assessed within hours after intraperitoneal LPS administration in mouse models of endotoxemia. The model captures key features of systemic inflammatory response syndrome, including cytokine release, immune cell activation, and downstream tissue responses, and has been used to evaluate anti-inflammatory interventions such as cytokine modulation, lipid mediators, and immune cell-targeting therapies.

Cell counting and cell-density analysis estimate the number of cells in a known volume or field area. Manual hemocytometer counting uses a chamber of defined geometry to convert counted cells into cells/mL, while automated counters and image-analysis workflows detect cell objects from optical, brightfield, fluorescence, impedance, or digital-image features. Trypan blue viability counting is based on dye exclusion: viable cells with intact membranes exclude dye, while non-viable cells with compromised membranes stain blue. The readout is total cell density, viable-cell density, dead-cell density, and percent viability. Cell density can also be estimated from microscopy images by counting objects per image area, from flow cytometry using calibrated volume or reference particles, or from in situ microscopy in bioreactors after calibration against reference methods such as hemocytometer or flow cytometry.

Autophagy is a process in which eukaryotic cells use lysosomes to degrade their own cytoplasmic proteins and damaged organelles under the regulation of autophagy related gene (Atg). Microtubule-associated proteins light chain 3 (LC3) is recognized as autophagy marker, which transfers from cytoplasmic LC3 (LC3-I) to membrane type (LC3-II). LC3-II/I ratio could be detected by Western Blot and fluorescence microscopy.

Apoptosis is a regulated, generally non-lytic cell-death pathway that removes unwanted, damaged, infected, or abnormal cells through coordinated morphological changes, caspase activation, DNA fragmentation, and membrane remodeling. The intrinsic apoptosis pathway is controlled mainly by mitochondrial outer membrane permeabilization, BCL-2 family proteins, cytochrome c release, apoptosome formation, caspase-9 activation, and downstream executioner caspase-3/7 activation. The extrinsic apoptosis pathway is initiated by death receptors such as Fas, TNFR, and TRAIL receptors, which recruit adaptor proteins and activate caspase-8 before engaging executioner caspases or mitochondrial amplification through BID cleavage. Apoptosis is linked to many phenotypes, including cancer cell killing, tissue homeostasis, immune regulation, neurodegeneration, infection response, and treatment-induced cytotoxicity; unresolved questions include how apoptosis interacts with necroptosis, pyroptosis, ferroptos

Lysosome and acidic-vesicle live-cell staining detects acidic intracellular compartments by using membrane-permeant acidotropic probes that accumulate in low-pH vesicles, including lysosomes, late endosomes, autolysosomes, and acidic phagosomes. LysoTracker staining is commonly used as an intensity-based readout of acidic lysosomal compartment abundance or enlargement, while acridine orange produces green fluorescence in less concentrated compartments and red fluorescence after concentration-dependent accumulation in acidic vesicular organelles. Loss or reduction of acridine-orange red signal can be used as a readout of lysosomal membrane permeabilization or reduced acidic-vesicle integrity. This protocol is designed for live cultured cells and can be adapted for fluorescence microscopy, high-content imaging, plate-reader readout, or flow cytometry when the selected literature supports the readout. Because these dyes report acidotropic accumulation rather than lysosome identity alone,

The MTT assay is a colorimetric endpoint assay for estimating viable cell number, cell growth, cytotoxicity, or cell activation in cultured mammalian cells. Living cells reduce the yellow tetrazolium salt MTT into purple/blue formazan, while dead cells do not generate the same signal; the resulting color can be quantified with a multiwell spectrophotometer. MTT reduction is commonly interpreted as a readout of metabolic activity that often correlates with viable cell number, but it should not be treated as a direct cell-counting method unless the assay is optimized for the cell type and experimental condition. Studies show that MTT reduction can involve mitochondrial and non-mitochondrial reducing systems, and formazan may accumulate in intracellular lipid droplets rather than simply marking mitochondria.

Macroautophagy is a conserved lysosome-dependent degradation pathway in which cytoplasmic material is sequestered into double-membrane autophagosomes and delivered to lysosomes for degradation and recycling. The pathway supports cellular homeostasis during nutrient limitation, organelle stress, protein-aggregate accumulation, infection, differentiation, and tissue remodeling by coupling cargo sequestration, autophagosome maturation, lysosomal fusion, and degradation of cargo-derived macromolecules. The core molecular sequence includes initiation by nutrient- and stress-regulated autophagy machinery, autophagosome nucleation, LC3/ATG8-family conjugation to autophagosomal membranes, cargo selection through receptors such as SQSTM1/p62, autophagosome-lysosome fusion, and lysosomal degradation. LC3 was identified as a mammalian homolog of yeast Atg8 that localizes to autophagosomal membranes after processing, and p62/SQSTM1 was shown to connect ubiquitinated cargo with autophagic degradati