Flow Cytometry and Light Scattering Technique in Evaluation of Nutraceuticals

C H A P T E R

24 Flow Cytometry and Light Scattering Technique in Evaluation of Nutraceuticals Igor Mindukshev, Igor Kudryavtsev, Maria Serebriakova, Andrey Trulioff, Stepan Gambaryan, Julia Sudnitsyna, Denis Khmelevskoy, Natalia Voitenko, Pavel Avdonin, Richard Jenkins and Nikolay Goncharov

INTRODUCTION AND BACKGROUND Modern automated particle- and cell-counting instruments use optical methods (light scatter), impedancebased methods based on the Coulter principle (changes in electrical current induced by blood cells flowing through an electrically charged opening), or a combination of both (Green and Wachsmann-Hogiu, 2015). They are used for a variety of applications, including routine cell culture, hematological analysis, and industrial controls (Dittami et  al., 2012). A limitation with existing Coulter technologies is the lack of metrics on the overall health of cell samples. Consequently, additional techniques must often be used in conjunction with Coulter counting to assess cell viability. This extends experimental setup time and cost because the traditional methods of viability assessment require cell staining and use of more elaborate and expensive equipment, such as a flow cytometer. In contrast to light scattering, which cannot distinguish damaged cells from debris, fluorescent dyes used in membrane integrity and mitochondrial polarization assays are capable of labeling and discriminating all cells in suspension (Reardon et al., 2014). Nevertheless, there is an advantage, besides low cost, of the light scattering technique: the capability for online recording of the dynamic changes of suspended viable cells in response to receptor agonists, toxic agents, and pharmaceuticals. A decade ago, authors of this chapter developed Nutraceuticals. DOI: http://dx.doi.org/10.1016/B978-0-12-802147-7.00024-3

a method for simultaneous monitoring of changes in shape and aggregation of platelets (Mindukshev et  al., 2005a). The signal of light scattering alterations at angles less than 6 degrees was shown to be caused by platelet aggregation dynamics (aggregation, disaggregation, coagulation), whereas over a range of larger angles (6–15 degrees) cell shape changes also contributed to the cell spherization and pseudopodia formation. Aggregation kinetics in saline solution under turbulent flow showed second-order kinetics in relation to initial cell concentration. The rate constant depended on stirring conditions and on calcium concentration in the medium. In subsequent studies, the low-angle light scattering technique was applied to studies of tumor cells and erythrocytes (Zinchenko et al., 2006; Mindukshev et al., 2007). Numerous investigations on toxic agents and pharmaceuticals have used flow cytometry or light scattering techniques (Mindukshev et al., 2005a, 2006; Goncharov et al., 2009; Radilov et al., 2009; Kudryavtsev et al., 2014; Geisler et al., 2014; Prado et al., 2015), although there are not many examples of the combined application of both technical approaches. Also, compared with pharmaceuticals and toxic agents, the toxic and mechanistic properties of nutraceuticals have not been so extensively studied. For example, the PubMed database contains nearly three times more papers with key words “flow cytometry toxic agents” than with key words “flow cytometry nutraceuticals” or “flow cytometry supplements.” Nevertheless,

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there are many common points and correlations between the techniques; therefore, convergence seems to be inevitable. For example, changes of mitochondrial membrane potential lead to morphological and volume changes in cells (Zinchenko et al., 2006). There are continuing concerns over the toxicity and the purity of many plant extracts used as nutraceuticals or supplements, as well as over the simple methods used to evaluate them. Cells affected with nutraceuticals/supplements can be easily analyzed using flow cytometry and light scattering, and sometimes this is performed together with electron microscopy, Western blotting, and immunohistochemical analysis for cell size and mitochondrial membrane potential, as well as cell cycle profile (LeBlanc et al., 2007; Buenz, 2008; Nielsen et al., 2008; Attilio et al., 2010; Yang et al., 2012). This chapter briefly describes the methodologies of low-angle light scattering and flow cytometry in relation to mechanisms of cell death and survival and presents an overview of some cellular studies of nutraceuticals involving light scattering and/or flow cytometry techniques.

Methodology of Research Mechanisms of Cell Death Studies of various cellular death pathways—such as apoptosis, necroptosis, autophagy, and some others— can aid our understanding of the mechanistic properties of toxic or therapeutic effects of chemical agents and are important in the development of therapeutic strategies. Apoptosis and necrosis are the two major modes of cell death. Initially they were thought to constitute mutually exclusive cellular states, although recent findings have revealed a balanced interplay between them. DNA fragmentation is one of the principal characteristics of apoptosis that is used in flow cytometry for the identification of apoptotic cells. The fragmentation is a result of activation of caspase-3, caspasse-6, and caspase-7, which initiates several mechanisms of fragmentation (Fan et al., 2005). The first one is bound to impairment of DNA separation due to inactivation by caspase-3 of poly(ADP-ribose) polymerase-1 (PARP-1) and topoisomerase I, which both participate in DNA recombination. Another mechanism is activation of DNases. One more substrate of caspases is histone H1, which blocks boarding of endonucleases on DNA fragments between separate nucleosomes (Kitazumi and Tsukahara, 2011). Mitochondrial damage is a universal sign of apoptosis specific to eukaryotic organisms (Abdelwahid et al., 2011). As a result, a series of proapoptotic factors are released to the cytoplasm: cytochrome C, AIF (apoptosis-inducing factor), Smac/DIABLO, endonuclease G, and procaspases 2, 3, and 9 (Orrenius et  al., 2007).

This is coupled with changes of the membrane potential (ΔΨm). Finally, the principal sign of apoptosis is phosphatidylserine exposure to the extracellular space due to impairment of ATP-dependent maintenance of the cell membrane lipid distribution (Verhoven et al., 1995). Also, exposure of phosphatidylserine can be initiated by scramblase, which is activated by Ca2+ at an early stages of apoptosis. Necrosis can occur in a regulated caspase-independent manner and shares characteristics with apoptosis. Several death initiator and effector molecules, signaling pathways, and subcellular sites have been identified as key mediators in both processes, either by constituting common modules or by functioning as a switch allowing cells to decide which route to take. As for autophagy, this is a predominantly cytoprotective process that has been linked to both types of cell death, serving either a prosurvival or a pro-death function (Nikoletopoulou et al., 2013). Autophagy is a central mechanism mediating the lysosomal degradation of cytoplasmic components. The central autophagic organelle is the autophagosome, a double membrane-bound vacuole that sequesters the cytoplasmic material destined to disposal. The ultimate destiny of the autophagosome is to fuse with a lysosome, resulting in the degradation of the autophagic cargo (Dupont et al., 2014). Dysregulation in autophagy is associated with a diverse range of pathologies. Both natural and synthetic compounds regulate the interplay between apoptosis, autophagy, and necroptosis, stimulating common molecular mediators and sharing common organelles (Radogna et al., 2015). Recently, ferroptosis was described as a nonapoptotic form of cell death that may facilitate the selective elimination of some tumor cells or may be activated in specific pathological states (Dixon et al., 2012). Ferroptosis is dependent on intracellular iron, but not other metals, and is morphologically, biochemically, and genetically distinct from apoptosis, necrosis, and autophagy. Cells transformed with oncogenic Ras have increased iron content relative to their normal cell counterparts through upregulation of transferrin receptor 1 and downregulation of ferritin heavy chain 1 and ferritin light chain (Yang and Stockwell, 2008). Ferrostatin-1 and deferoxamine are inhibitors of ferroptosis in cancer cells and glutamate-induced cell death in organotypic rat brain slices, suggesting similarities between these two processes. Erastin (as well as glutamate) inhibits cystine uptake by the cystine/glutamate antiporter, ultimately leading to iron-dependent oxidative death (Dixon et al., 2012). Numerous overlapping mechanisms using thiol compounds exist to decompose hydrogen peroxide to water to bypass the generation of hydroxyl radicals. Hydrogen peroxide is now recognized as a signaling molecule whose main regulator is the peroxiredoxin/ sulfiredoxin systems, which are at least partially under

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the control of the Nrf2/Keap1 system. Ironically, the persistent activation of antioxidant systems via genetic alterations in Nrf2 and Keap1 contributes to carcinogenesis. Some cancers hijack the Nrf2/Keap1 system with mutation in these genes, which persistently activate antioxidant systems in the cancer cells; overexpression of CD44v(8-11) stabilizes the cystine/glutamate antiporter to increase cysteine and GSH levels in cancer stem cells (Toyokuni, 2014). Pyroptosis, a highly pyrogenic inflammatory form of cell death, develops due to formation of inflammasomes, which are cytosolic multiprotein platforms assembled in response to invading pathogens and other danger signals. Typically, inflammasome complexes contain a sensor protein, an adaptor protein, and a zymogen (procaspase-1). The assembly results in processing of inactive procaspase-1 into an active cysteine-protease enzyme (caspase-1), which subsequently activates the proinflammatory cytokines IL-1β and IL-18 (Vanaja et  al., 2015). Although these processes intend to protect the body from insults, prolonged or exacerbated inflammatory responses associated with inflammasome activation are related to a growing number of diseases. Recently, inflammasome activation and autophagy were shown to be linked and to mutually influence each other (Martins et al., 2015). Eryptosis, the suicidal erythrocyte death characterized by cell shrinkage and cell membrane scrambling, is stimulated by Ca2+ entry through Ca2+-permeable PGE2-activated cation channels by ceramide, caspases, calpain, complement, hyperosmotic shock, energy depletion, oxidative stress, and deranged activity of several kinases (e.g., AMPK, GK, PAK2, CK1α, JAK3, PKC, p38-MAPK). Eryptosis is triggered by intoxication, malignancy, hepatic failure, diabetes, chronic renal insufficiency, hemolytic uremic syndrome, phosphate depletion, malaria, iron deficiency, sickle cell anemia, thalassemia, glucose-6-phosphate dehydrogenase deficiency, and many other factors or pathological states. Eryptosis may precede and protect against hemolysis, but by the same token it may result in anemia and deranged microcirculation (Lang and Lang, 2015).

Flow Cytometry in Assessment of Cellular Viability This section briefly outlines the commonly used methods in flow cytometry that are based on the assessment of phosphatidylserine exposure, mitochondrial transmembrane potential, and DNA accessibility and/or fragmentation. It should be noted here that methods of revealing the DNA fragmentation are rather laborious, greatly depend on professional skills of the personnel, and require costly reagents. Therefore, other methods based on staining of DNA have wide applications. Regarding the apoptotic cells, they bear an incomplete

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or hypodiploid number of DNA (less than 2N) forming the so-called sub-G1 peak. A distribution of cells according to fluorescence intensity after staining of DNA is caused by the stoichiometric character of the staining, and this kind of binding is a distinguishing feature for all the probes of this group. Besides apoptotic studies, these probes are widely used to investigate proliferative activity of various cells. Moreover, within the frame of apoptotic studies they are used together with other probes oriented at revealing asymmetry of the cell membrane, changes of mitochondrial membrane potential, and activity of caspases (Zhao et  al., 2009; Wlodkowic et al., 2012). YO-PRO-1 and PO-PRO-1 are DNA-binding probes that are used together with PI to stoichiometrically bind to nucleic acids (i.e., RNA in the cytoplasm and DNA in the nucleus) (Idziorek et  al., 1995). Being excited at 488 nm, YO-PRO-1 emits fluorescence in the green region (near 509 nm) and PI emits fluorescence in the red region (near 617 nm) of the spectrum. However, the principal difference between them is the way in which they penetrate the cells. YO-PRO-1 enters cells through purinergic receptors (P2RX7), which are ligand-dependent ionic channels (Glisic-Milosavljevic et  al., 2005; Stokes et  al., 2006). In the living cells, YO-PRO-1 would not accumulate because of almost inactive channels and very low transporting ability of the membrane. The channels become activated at early stages of apoptosis development, together with disturbance of plasma membrane asymmetry. To reveal the later stages, cells should be stained with DNA-binding PI, which does not need special transporters and is able to penetrate into cells and nuclei only via damaged/fragmented membranes; this process takes place during late stages of apoptosis or necrotic cell death. Thus, the living cells would not be stained at all, the cells at early stages of apoptosis would be only YO-PRO-1-positive, and the cells at later stages would be effectively stained by both probes. The SYTO family of dyes, capable of measuring apoptosis with flow cytometry techniques, was first announced in 1995 (Frey, 1995). Presently, there are four main groups of cyanide stains distinguished by excitation wavelengths. Dyes of the SYTO blue group (SYTO 40–45) have an excitation maximum at 419–452 nm, with fluorescent emission maximum at 445–484 nm. Dyes of the SYTO green group (SYTO 11–16 and 20–25) have an excitation maximum at 483–521 nm and fluorescent emission maximum at 500–556 nm. Two other groups are SYTO orange (SYTO 80–86) and SYTO red (SYTO 17 and 59–64), with corresponding excitation maxima at 528– 567 and 598–654 nm and fluorescent emission maxima at 544–583 and 620–680 nm. Distinguishing features of SYTO dyes are their ability to spontaneously penetrate the lipid bilayers and bind with nucleic acids (DNA and RNA) in cytoplasm, nucleus, and mitochondria.

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This interaction is accompanied by multiple (up to 40 times) enhancements of fluorescence. Another advantage is an absence of visible toxic effects, which was demonstrated in long-term experiments (up to 72 h) with cells (Wlodkowic et  al., 2007). The exact mechanism of interaction of SYTO dyes with nucleic acids is not known (Wlodkowic et al., 2012), but their principal difference from other DNA dyes is low interaction and decrease of fluorescence in apoptotic cells. Thus, live cells are designated as SYTObright, whereas dying cells are SYTOdim. Interestingly, there may be a correlation between the fluorescence intensity of SYTO dyes and lipophilic cationic mitochondrial probes (e.g., JC-1 or TMRM) (Wlodkowic et  al., 2008), which is indicative of an analogy in their mechanism of interaction due to positive-charged groups within molecules and dependence of interaction and fluorescence on ΔΨm. Methods of studying mitochondrial membrane potential ΔΨm with flow cytometry are based on use of cationic lipophilic dyes, which are also known as “mitochondrial probes.” These dyes spontaneously penetrate through lipid bilayers and accumulate in areas with high concentrations of protons (i.e., under inner mitochondrial membrane). This is accompanied by changes of fluorescence. If proton concentration is low (decrease of ΔΨm), then accumulation of dye and its fluorescence is also low. Most probes are related to three principal groups: rhodamine derivatives, such as rhodamine 123 (Rh123) and ethyl (TMRE) and methyl (TMRM) ethers of tetramethylrhodamine; carbocyanine derivatives, such as 5,5′-6,6′-tetrachloro-1,1′,3,3′-tetraethylben zimidazolylcarbocyanine iodide (JC-1), 3,3′-dimethyl-αnaphthoxacarbocyanine iodide (JC-9), and 3,3′-dihexiloxacarbocyanine iodide DiOC6(3); and MitoTracker dyes (Cottet-Rousselle et al., 2011). To investigate asymmetry of the apoptotic cell membranes, namely to reveal phosphatidylserine, conjugates of annexin V with fluorochromes (usually FITS or PE) are used. In the presence of Ca2+ this protein can conjugate with negatively charged phospholipids, with the greatest affinity to phosphatidylserine (KD = 5 × 10−10 M). The changes of mitochondrial membrane potential and activation of caspase-3 precede the onset of phosphatidyl serine in the outer layer, and the latter precedes reorganization of cytoskeleton and the cell membrane integrity. Therefore, there are many combinations of cell staining with Annexin V and other probes for revealing both early and late phases of apoptosis (Wlodkowic et  al., 2011). The cell membrane of healthy cells is impermeable to DNA-binding stains, so they are negative with both Annexin-FITC and propidium iodide (AnnV-FITC–PI–). At early apoptosis, phosphatidylserine is exposed on the cell surface, making it possible to conjugate with AnnV-FITC in the presence of Ca2+ ions. This exposure coincides with chromatin condensation and DNA

fragmentation, although the membrane is not yet permeable to DNA-binding stains. On a histogram, the early apoptotic cells are AnnV-FITC-positive and PI-negative (AnnV-FITC+ PI–); dead cells or those at late stages of apoptosis would be AnnV-FITC-positive and PI-positive (AnnV-FITC+ PI+). The same characteristics of fluorescence would also have necrotic cells, although these cells could be AnnV-FITC-negative (AnnV-FITC–PI+).

LIGHT SCATTERING TECHNIQUE IN STUDIES OF PLATELETS AND RED BLOOD CELLS Laser particle size analyzers available from commercial sources (e.g., Malvern, Beckman, Horiba, Fritch, and others) are widely used for size distribution analysis of natural and artificial particles. However, until now these devices have not been routinely applied to biological systems. Based on the technology used in these analyzers, the authors of this chapter created a new device (LaSca, Biomedsystems Ltd., Russia) and developed original software for the registration of the kinetics of changes in cell volume, morphology, and aggregation (Mindukshev et  al., 2005b, 2010, 2012). Quantitative characterization of cell volume can only be achieved under single scatter conditions. For this, the cell density must be adjusted to yield a transmittance of at least 50% of the transmittance of the blank cuvette (Van de Hulst, 1981). Cell volume and cell shape undergo dynamic changes in many physiological and pathological processes. Active volume regulation to protect cells from lysis and apoptosis and to maintain optimal concentration of intracellular components is one of the most basic and essential cellular functions. Several methods are applied for measurement of cell volume changes evoked by different stimuli. Changes in volume of attached cells can be accurately measured by the 3D reconstruction of cell shape based on cell images acquired from two perpendicular directions (Boudreault and Grygorczyk, 2004). A laser light scattering method was introduced and used for volume change registration of attached cells (McManus et  al., 1993) and cells in suspension (Faggio et al., 2011). However, no quantitative or time-resolved data regarding cell volume have yet been presented by these methods. Recently, for analysis of cell volume in suspension, which is especially important for blood cells, the Cell Lab Quanta SC Flow analyzer was introduced by Beckman and has been used for analysis of cellular apoptosis (Bortner et  al., 2007). However, flow cytometry methods are not suitable for online registration of fast changes of cell volumes. For demonstrating the feasibility of a method for characterization of cell volume regulation, the authors of this chapter used osmotic stress-induced volume changes in

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FIGURE 24.1  Application of low-angle light scattering method for registration of HL-60 cell volume changes. (A) Original recordings (representative of four independent experiments) of light scattering intensity (LSI) measured at 1° and 12° of HL-60 cells incubated in media with different osmolarity. (B) Light microscopic images of HL-60 cells incubated for 5 min in media with the indicated osmolarity. The mean diameter calculated from 60 individual images of control cells (302 mOsm) was 11.2 ± 0.6 µm, in 495 mOsm it was 8.1 ± 0.3 µm, and in 79 mOsm it was 14.5 ± 0.5 µm, which corresponds to the calculated mean cell volumes (MCV; data from Figure 24.1D) 654 ± 52 fl, 531 ± 96 fl, and 1,240 ± 93 fl, correspondingly. (C) Histogram of cell volume distribution in media of the osmolarity indicated. (D) Calculated relations of MCV with the osmolarity of the media. (E) Dependence of relative LSI (normalized to LSI in isotonic media) in indicated LSI angles on changes of MCV.

HL-60 cells (Figure 24.1). The cell density range applicable for HL-60 cells corresponds to 120–130 cells/µL. To measure cell volume distribution, light scattering was used to monitor 28 channels ranging from 1° to 12°. For simplification of the graph, changes of light scattering intensity (LSI) are presented only for 1° and 12° (Figure 24.1A). In hypertonic conditions, LSI decreased at 1° but increased at 12°. In hypotonic conditions, the changes in LSI are inversed for 1° and 12°. It should be noted that cells are highly tolerable to phase changes of osmolarity, whereas acute changes from iso-osmotic to hypo-osmotic (45 mOsm) solution lead to immediate cell lysis, which is reflected in the strong decrease of LSI (Figure 24.1A). The registered scatter intensity correlates with changes of cell size observed by light microscopy (Figure 24.1B). Cell volume distributions in hyperosmotic, iso-osmotic, and hypo-osmotic solutions were constructed according to the changes of LSI. Based on data of cell volume

distribution, the mean cell volume (MCV) was calculated (Figure 24.1D). Because the relation between LSI and cell volume is nonlinear (Figure 24.1E), all data registered at 28 channels are necessary for calculation of cell volume distribution and MCV. The method shows very high sensitivity, and it allows measurements and calculations of acute MCV changes of as little as 1%, making this method especially applicable for receptor-mediated kinetic analysis of fast and reversible cell volume changes. With this method, one can also demonstrate the kinetics of erythrocyte transformations (Figure 24.2). To maintain single scattering conditions for erythrocytes, the cell density was adjusted to 500–600 cells/µL. Under hypoosmotic conditions, the LSI at 1° increased but decreased at 12° (Figure 24.2A). Cell volume distribution for these conditions was calculated according to the changes of LSI (Figure 24.2B). The maximal increase of erythrocyte MCV was achieved at 130 mOsm from 91± 6 fL (control, isotonic

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FIGURE 24.2  Application of low-angle light scattering method to the registration of erythrocyte transformations. (A) Original recordings (representative of four independent experiments) of light scattering intensity (LSI) measured at 1° and 12° of erythrocytes incubated in iso-osmotic and hypo-osmotic media. (B) Histogram of cell volume distribution in media of the osmolarity indicated. (C) Calculated relation of MCV to the osmolarity of the media (red line indicates process of erythrocytes hemolysis). (D) Original recordings (representative of four independent experiments) of light scattering intensity (LSI) measured at 1° and 12° of erythrocytes incubated with 2 µM of calcium ionophore (A23187). (E) Histogram of cell volume distribution after stimulation with A23187 at the time indicated. (F) Calculated relation of MCV to the time of incubation with A23187. (G) Light microscopic images of control erythrocytes (i), incubated 5 min in hypo-osmotic (180 mOsm) solution (ii), and after 5 min (iii) and 30 min (iv) of incubation with A23187.

conditions) to 152 ± 8 fL. Lower osmolarity caused a fast and irreversible lysis reaction (Figure 24.2C). Under hypoosmotic conditions, erythrocytes change their shape from discoid to spherical (Figure 24.2G(i, ii)). Calcium increase induced by the calcium ionophore (A23187) causes erythrocyte shrinkage. With respect to light scatter intensity, the changes in erythrocyte shape on intracellular calcium increase elicit a biphasic signal (Figure 24.2D), reflecting a biphasic change of MCV (Figure 24.2F) as calculated from the data on cell volume distribution (Figure 24.2E). The initial increase of MCV corresponds to the transformation of erythrocytes from discoid to a spherical shape, and the subsequent decrease of MCV corresponds to cell shrinkage (Figure 24.2F and G(iii, iv)). Several methods, including light scattering analysis (Turcu et al., 2006), are used for investigation of erythrocyte transformations under different conditions. The fast and rather easy registration of erythrocyte transformations is one of the advantages of the method.

Next, this method was applied for registration of blood platelet shape change and aggregation. During the aggregation process, platelets first undergo a change from an oblate spheroid shape to a spherical shape, and then to a complex shape characterized by cell protrusions (shape change) linking the cells to a microaggregate and, in a later step, to a macroscopic clot. The widely used Born aggregometry method, which is based on multiple scattering, registers changes in the transmission of the whole sample, thus providing relatively low sensitivity and limited information about net changes in cell shape or volume; this is overcome with our technique. The theoretical basis of single scattering is more elaborate than for multiple scattering and allows the combined analysis of cell size, volume, and distribution, together with shape change and aggregation. Also, the sensitivity of our method is considerably increased due to the fact that the scattered light is observed against a virtually dark background. It is widely recognized that particularly

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FIGURE 24.3  Application of low-angle light scattering method for measuring platelet aggregation and shape change. (A) Light scatter intensity (LSI) measured at 12 different scatter angles of ADP (500 nM) stimulated human platelet suspension. For better resolution, the ordinate scale is enlarged five times for the 5°–8° curves and 10 times for the 9°–12° curves. According to the predicted changes in LSI during aggregation, the LSI increases in the range of 1°–5° and decreases in the range of 8°–12°. (B) Characterization of platelet shape change by LSI. LSI presented as % increase from basal (100%) measured at seven different scatter angles with ADP-stimulated (90 nM) human platelet suspension. The maximal increase in LSI is observed at 12°. (C) Original recording of LSI measured at 1° and 12° of ADP-stimulated human platelets with the concentrations indicated. I(1) and I(12) correspond to LSI at 1° or 12°. (D) Dependence of platelet activation (flow cytometry analysis of platelet P-selectin CD62P, surface exposure, and α2bβ3 integrin activation (PAC-1 binding)) and initial velocity (Uia) of aggregation from ADP concentrations. LSI is given in arbitrary units of the intensity of the scattered light. Flow cytometry data presented as mean fluorescence. For flow cytometry analysis, platelets were taken directly from the LaSca cuvette 2 min after stimulation.

mild platelet function disorders demand a highly sensitive analysis not provided by conventional aggregometry methods (Watson et  al., 2010). To establish and verify this novel method, authors of this chapter monitored ADP-induced platelet shape change and aggregation. The maximum platelet density allowing for single light scattering was determined to be 10,000 platelets/µL; therefore, platelet reach plasma (PRP) used in the experiments was diluted with buffer accordingly. For verification of the underlying theory, a model for the platelet aggregation processes was established. The model predictions were experimentally proven for the aggregation of human platelet suspensions. Angle-dependent scatter intensities were measured in a range from 1° to 12° using human platelet suspensions stimulated with 500 nM ADP. The experimental data were in good accordance with the calculated scatter curves. As predicted by the model, the

intensity of the scattered light increased between 1° and 5° and decreased between 9° and 12° (Figure 24.3A) during ADP-stimulated platelet aggregation. In the range between 5° and 8°, the changes in scatter intensity are less pronounced and are thus not suited for aggregation monitoring. From the scatter curves it is evident that changes in scatter intensity are maximal at 1°, making this angle particularly suitable for aggregation measurement. Platelet shape change is reflected by an increase of scatter intensity at 8° to 12°, with a maximal increase at 12° (Figure 24.3B). Consequently, a scatter angle of 1° was chosen for the detection of platelet aggregation and 12° was chosen for the monitoring of platelet shape change. Maximal shape change was induced by 90 nM ADP, a concentration not sufficient to induce platelet aggregation (Figure 24.3B). For quantitative characterization of platelet aggregation response, different ADP concentrations

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were applied, showing a dose-dependent increase of aggregation velocity and amplitude with a maximum response at 1 µM of ADP (Figure 24.3C). Verification of the observations with the light scattering method was accomplished with measurement of the established platelet activation markers P-selectin expression and activated integrin αIIbβ3 in flow cytometry (Figure 24.3D). These data show a good correlation of light scatter intensity changes with common activation markers (Figure 24.3D). Platelet aggregation is characterized by the initial velocity (Via) of aggregation, which was calculated from the first 60-second interval after stimulation and the maximal amplitude [ΔI(1)], calculated as differences between the maximal scatter intensity and the initial scatter intensity prior to stimulation. The aggregation velocity was used as an important parameter for pharmacological experiments. The maximal aggregation velocity (Vmax) was calculated for each experiment. For a direct comparison of different experiments, the relative initial aggregation velocity, designated as Uia, was calculated as Uia=Via/ Vmax, and the normalized maximal aggregation velocity (Umax) was also calculated. The normalized initial aggregation velocity Uia according to the Hill equation shows the expected sigmoid dose dependence on ADP concentration (Figure 24.3d). The parameters for this equation are obtained from a Schild plot. The value for the Hill coefficient (2.53± 0.29) reflects the observed cooperativity of P2Y1 and P2Y12 receptor activation in platelets (Gachet, 2006). ADP concentrations in routinely used clinical platelet aggregation tests are in the range of 5–10 µM, which gives a maximal aggregation response. The ЕС50 for ADPinduced platelet aggregation calculated from our data was 307± 17 nM, with a maximum response to 1 µM. Because nucleotide-induced platelet activation is mediated by three different purinergic receptors (Geiger et al., 1998), the study was performed for detailed determination of the individual contribution of these receptors with established specific inhibitors (Figure 24.4). Inhibition of the P2Y1 receptor with MRS significantly increased the EC50 for ADP but did not affect Umax (Figure 24.4A). P2Y12 receptor inhibition with ARC significantly decreased Umax, leaving the EC50 unchanged (Figure 24.4B). These data underline the hypothesized function of the P2Y1 receptor, which is necessary for the initiation of platelet aggregation, and the P2Y12 receptor for completion of ADP-induced aggregation (Gachet, 2006). However, as was described for P2Y1 (Léon et  al., 1999) and P2Y12 (Cattaneo et al., 2003) knockout mouse models, lack of either receptor still allows for a small aggregation response. The similar results were obtained in light scatter experiments (Mindukshev et al., 2012). For platelet shape change, EC50 for ADP was 47± 7 nM, with a maximal response at 90 nM. This ADP response could be completely blocked by a specific P2Y1 receptor inhibitor, without any effects of P2Y12 and P2X1 receptor inhibitors

(Figure 24.4C). ADP stimulation of shape change can be described by the Langmuir equation (Hill coefficient 1.1± 0.3). P2X1 receptor is a well-characterized nonselective ligand-gated cation channel activated by extracellular ATP. It was shown that the P2X1 receptor in platelets is activated at high concentrations (10 µM and more) of nonhydrolyzed ATP analogs (Rolf and Mahaut-Smith, 2002), whereas the EC50 for this receptor expressed in oocytes is in the nanomolar range (Rettinger and Schmalzing, 2003). One can directly demonstrate the P2X1 receptor–mediated activation of platelet shape change by ATP (Figure 24.4D). The present technique offers quantitative characterization of P2Y1-mediated and P2X1-mediated platelet shape changes. This method could be used for sensitive detection of receptor defects or drug action. The low sample consumption makes this method ideally suited for investigation of mouse platelets and for transgenic mouse models in particular (Mindukshev et al., 2012).

Studies of Nutraceuticals with the Help of Light Scattering and Flow Cytometry Flow cytometry and/or light scattering have been applied to investigate the following: the effects of omega-3 fatty acid supplementation on platelet and endothelial activation (Mackay et al., 2012); ● effects of conjugated linoleic acid or in combination with omega-3 on metabolic characteristics in muscle cells (Vaughan et al., 2012); ● inter-relationships among the hypothalamuspituitary-adrenal stress axis and T-cell maturation in both zinc deficiency and responses during zinc repletion (Blewett and Taylor, 2012); ● protective effects of gallic acid on endothelial cell death and the mechanisms involved (Kam et al., 2014); ● ability of quercetin and apigenin to modulate platelet activation and aggregation (Del Turco et al., 2015); ● the potential of epigallocatechin gallate to suppress cell proliferation of SSC-4 human oral squamous cell carcinoma (Irimie et al., 2015) and human nonsmallcell lung cancer A549 cells (Sonoda et al., 2014); ● curcumin-induced apoptotic cell death of NCCIT human embryonic carcinoma cells (Yun et al., 2015); ● toxic effects of garlic (Allium sativum) on pathogenic and beneficial bacteria (Booyens and Thantsha, 2014); ● effects of sodium 2-propenylthiosulfate, allicin, and diallyl trisulfide (organosulfane sulfur compound from garlic) in cancer cell cytotoxicity (Sabelli et al., 2008; Chandra-Kuntal et al., 2013; Tao et al., 2013) and in many other investigations. ●

Some work over the past decade was aimed at the development of methodology for cytophysiological screening of

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FIGURE 24.4  Quantitative characterization of P2 receptor–induced platelet aggregation and shape change. Diluted human platelet suspension was analyzed by light scattering method for ADP-induced aggregation (A, B), ADP- (C), and ATP-induced shape change (D). Dependence of initial velocity (Uia) of aggregation from ADP concentrations in the absence (ADP) or presence of different concentrations of P2Y1 receptor antagonist (+MRS2179) (A) and P2Y12 receptor antagonist (+ARC69931) (B). Inhibition of P2Y1 receptor significantly increases EC50, without changes of Umax, whereas inhibition of P2Y12 receptor significantly decreases Umax. (C, D) Characterization of platelet shape changes induced by different ADP (C) and ATP (D) concentrations calculated as percent increase of LSI at 12° from the basal signal level (∆ I(12), %). The P2Y1 receptor can be described by the Langmuir equation (h = 1.1 ± 0.3), whereas P2X1 receptor can be described as the sigmoid shape of curves in normal axes and Hill coefficient higher than unity (h = 2.6 ± 0.5).

antidote substances and/or those enhancing therapeutic effects of antineoplastic and other drugs. Flow cytometry, together with low-angle light scattering techniques, and some other methods of cell viability assessment were valuable instruments for this research (Mindukshev et al., 2005a, 2006, 2007; Zinchenko et  al., 2006, 2007; Radilov et  al., 2009; Kudryavtsev et  al., 2014; Prokofieva and Goncharov, 2014; Goncharov et al., 2009, 2015a). Development of cancer is dependent on different cells (cancer and stem cancer cells, endothelial cells, blood platelets, red blood cells, neutrophyles, and stromal cells) contributing to tumor growth and forming a vicious cycle (Szabo et  al., 2010; Qu et  al., 2013). To break the cycle, an effective therapeutic complex should be multi-targeted (i.e., simultaneously directed at different molecular and cellular targets). Nutraceuticals seem to be complex and are applied as adjuvant and/or neoadjuvant therapy. The essence of an anticancer screening algorithm is treating different tumor cells and other cells (endothelial, platelets, red blood cells, etc.) in vitro with various concentrations of camptothecin (positive control) or other cytotoxic drugs

with known molecular targets (rotenone, iodoacetate, etc.) and then with nutraceuticals, such as garlic oil, diallyl disulfide (DADS), green tea extract, epigallocatechin gallate, or others, followed by testing of their combinations to reveal if there was amplification, attenuation, or no combined effect at all. This is a rather simple and effective approach, allowing the opportunity to reveal or suggest a “personal” response of the cells participating in the tumor growth. Further analysis of the possible impact on the cells relating to effective concentrations of the substances tested renders it possible to suggest an effective combination to test it in vivo. Such an algorithm is now undergoing development with nutraceuticals; the first positive results were obtained with fluoroacetate applied in combination with cyclophosphamide, which proved to be no less effective, and even a bit more effective, than the combination of metformin with cyclophosphamide (Anikin et al., 2013, 2014). Figures 24.5–24.8 show some primary results obtained with flow cytometry, and those obtained with the low-angle light scattering technique within the frame of complex anticancer screening experiments are shown in Figure 24.9.

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FIGURE 24.5  Dissipation of mitochondrial transmembrane potential and plasma membrane permeability assessment. Analysis by concurrent staining with lipophilic cationic carboxycyanine dye DiOC6(3) and propidium iodide (PI). THP-1 cells (human monocytic leukemia cell line) were either unexposed (A) or exposed to camptothecin 5 μM (B) or to DADS 100 μM (C). Viable cells (V) accumulate DiOC6(3) in mitochondria and are impermeable to PI. THP-1 cells during early apoptosis (Ap) have impaired mitochondrial transmembrane potential and an impermeable cell membrane (decreased intensity of DIOC6(3) and PI-negative). Ap/N cells exhibit loss of mitochondrial transmembrane potential (low DiOC6(3) fluorescence) and become permeable to PI.

FIGURE 24.6  Analysis of plasma membrane permeability with cyanine dyes. THP-1 cells either unexposed (A) or exposed to DADS 100 μM (B) or DADS 33 μM (C). Live cells (V) remain impermeable to both dyes (YO-PRO-1neg/PIneg). Early apoptotic cells (Ap) are characterized by cell membrane permeability for YO-PRO-1 due to activation of P2X7 channels (YO-PRO-1pos/PIneg). THP-1 cells in late stages of apoptosis and necrotic cells (Ap/N) lose cell membrane integrity and become permeable to both YO-PRO-1 and PI.

FIGURE 24.7  Detection of early and late stages of apoptosis by concurrent staining with annexinV-FITC and DRAQ7. THP-1 cells unexposed

(A), exposed to camptothecin (B), or exposed to DADS (33 μM) (C) were treated with annexin V-FITC conjugate and DRAQ7, and their green and far-red fluorescence were measured by flow cytometry. Viable (V) cells comprise a double-negative population in the lower left corner. At an early stage of apoptosis (Ap), annexin V-FITC interacts with phosphatidylserine molecules at the outer lipid bilayer of cell membranes, but the membrane does not lose its integrity, so the cells are DRAQ7-negative. At a late stage of apoptosis (Ap/N), the cells bind annexin V-FITC and are stained with DRAQ7, which binds readily to nuclear DNA and reports cell death by strong far-red fluorescence.

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FIGURE 24.8  Detection of apoptosis by SYTO 16 dye and DNA-binding dye DRAQ7. The cells were treated with both dyes for 15 min in

final concentrations of 250 nM and 3 μM, respectively, then washed with PBS supplemented with 2% of fetal calf serum (FCS), centrifuged for 7 min (330 g), resuspended in fresh PBS, and analyzed on flow cytometer. THP-1 cells were either unexposed (A) or exposed to camptothecin 2.5 μM (B) or to DADS 33 μM (C) for 24 h. Live cells (V) are SYTO 16bright and DRAQ7-negative. Early apoptotic cells (Ap) are SYTO 16dim but still DRAQ7-negative. Late apoptotic/necrotic cells (Ap/N) are SYTO 16low and DRAQ7bright.

FIGURE 24.9  Parameters of hemolysis of human red blood cells measured by low-angle light scattering technique at the angle of 1.5°. (A) Maximal rate (velocity) of hemolysis. (B) Quantity of hemolyzed cells (%). Red blood cells taken from venous blood of a healthy individual were washed with saline solution 1 and treated with tert-butyl-hydroperoxide (tBH), garlic oil (GO), or GO followed by tBH (with 1.5-minute interval) for 30, 60, 120, 180, or 1,500 minutes. Two saline solutions were used in the experiments (mM): (1) NaCl 140, KCl 2, HEPES 10, MgCl2 1, glucose 5, and EGTA 2; and (2) NH4Cl 140, KCl 2, HEPES 10, and glucose 5. RBCs were washed and incubated in saline solution 1, and at each time point hemolysis was initiated with saline solution 2 (140 mM NH4Cl). Decrease of maximum velocity of hemolysis (A) and decreased percent of hemolysis (B) after exposure to tBH indicate enhanced rigidity of erythrocytes upon development of eryptosis; 0.1 mM GO shows a protective effect if added to the cells 1.5 minute before 1.5 mM tBH. *P < 0.05; #P < 0.3.

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CONCLUDING REMARKS AND FUTURE DIRECTIONS The search for drugs with low toxicity, together with the development of technical and methodological approaches for their testing in vitro, is an urgent task for experimental biology and medicine. Discriminating between the toxic, nontoxic, and protective effects and understanding how cell death or cell viability is regulated upon exposure to cytotoxic agents and/or nutraceuticals may provide insight into how these compounds can be combined and manipulated in the treatment of disease. Most exogenous compounds, including toxic agents, pharma compounds, and nutraceuticals (with low specificity), have multiple cellular and molecular targets. For example, some thiols and disulfides are used as nutraceuticals in complex cardiovascular or cancer therapy (Ried and Fakler, 2014; Cerella et  al., 2011), and at certain concentrations they can be toxic to endothelial cells (Prokofieva and Goncharov, 2014) and red blood cells (Munday, 2012; Alzoubi et  al., 2015), with ROS being the most likely mediators of their toxic effects. However, there are different cellular and molecular sources of ROS with respect to their participation in pathogenesis of diseases (Goncharov et  al., 2015b), and revealing the impact of each of them at different stages of pathogenesis is a major challenge. Therefore, the methodologies of experimental and preclinical studies should be revised and expanded to bring into play more in vitro models and fewer in vivo or invasive experiments; even more so, ethical and financial considerations favor these scientific endeavors. Intuitively, there should be a strong correlation between cellular responses in vitro and the final efficacy of the therapy, although this correlation has yet to be proven. Thus, the development of tools for interventions aimed at improving human health require a comprehensive knowledge of the cellular reactions to positive or negative stimuli, correlating these in vitro and/or ex vivo obtained data with final in vivo therapeutic or toxic effects. Despite many studies on separate cell lines with flow cytometry or light scattering, there are very few examples of successful development of such a complex methodology. Dr. F. Lang and collaborators have made much progress in establishing mechanisms of eryptosis evoked by different agents, including nutraceuticals, through flow cytometry and light scattering studies (based just on forward scatter) (Alzoubi et  al., 2015; Peter et  al., 2015). Importantly, they also applied some other biochemical techniques to reveal the molecular mechanisms (Lang et al., 2015; Lang and Lang, 2015). In a search for alternative ways of monitoring functional potential and effectiveness of organosulfur compounds and polyphenols within the frame of anticancer therapy development, experiments are being conducted by our group with flow cytometry and the low-angle light

scattering technique. Some primary results demonstrating characteristic elements of the approach are given in this chapter, although the final scientific results are the matter of special publications and/or a patent pending. Low-angle light scattering analysis of erythrocytes, platelets, and other cells allows very sensitive and simultaneous quantitative online registration of changes in cell volume, shape change, and cell–cell interactions. This ensures detection of subtle changes in cell volume and shape changes when compared to other techniques such as Born aggregometry. However, flow cytometry has greater potential to enhance sensitivity for measuring the small particles of sub-micrometer size, primarily due to fluorescence triggering. For example, approximately 50 times more Annexin-5 (Anx5) positive extracellular vesicles can be detected by flow cytometry, with the limit of detection of the fluorescence triggering method estimated at approximately 1,000–2,500 Anx5 molecules (Arraud et al., 2015). Nevertheless, label-free and rapid classification of cells can have a wide range of applications in experimental biology. A new robust method of polarization diffraction imaging flow cytometry (p-DIFC) has been developed recently for achieving this goal (Feng et  al., 2014). This method has the capacity to discriminate cells of high similarity in their morphology with “fingerprint” features extracted from the diffraction images, which may be attributed to subtle but statistically significant differences in the nucleus-to-cell volume ratio in some cells. It was noted recently that future directions of in vitro instrumentation for cell screening research should include the addition of new parameters and the development of point-of-care instrumentation (Green and Wachsmann-Hogiu, 2015). The low-angle light scattering technique is a simple but unique supplement to the methodology of investigating natural supplements such as nutraceuticals. Taken together, these supplements would greatly add to human health and knowledge.

Acknowledgments This work has been supported by the Russian Foundation for Basic Research Grants 14-04-00951, 13-04-01728, and 15-04-02438, and the Russian Science Foundation Grant 14-15-01004 (section “Mechanisms of Cell Death”).

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