Effect of honokiol on erythrocytes

Toxicology in Vitro 27 (2013) 1737–1745

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Effect of honokiol on erythrocytes Mohanad Zbidah 1, Adrian Lupescu 1, Tabea Herrmann, Wenting Yang, Michael Foller, Kashif Jilani, Florian Lang ⇑ Department of Physiology, University of Tuebingen, Gmelinstraße 5, 72076 Tuebingen, Germany

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Article history: Received 6 December 2012 Accepted 3 May 2013 Available online 11 May 2013 Keywords: Phosphatidylserine Honokiol Calcium Cell volume Eryptosis

a b s t r a c t Honokiol ((3,5-di-(2-propenyl)-1,1-biphenyl-2,2-diol), a component of Magnolia officinalis, stimulates apoptosis and is thus considered for the treatment of malignancy. In analogy to apoptosis of nucleated cells, erythrocytes may enter eryptosis, a suicidal death characterized by cell shrinkage and by breakdown of cell membrane phosphatidylserine asymmetry with phosphatidylserine-exposure at the erythrocyte surface. Eryptosis may be triggered following increase of cytosolic Ca2+-activity ([Ca2+]i). The present study explored, whether honokiol elicits eryptosis. Cell volume has been estimated from forward scatter, phosphatidylserine-exposure from annexin V binding, hemolysis from hemoglobin release, [Ca2+]i from Fluo3-fluorescence, and ceramide from fluorescent antibodies. As a result, a 48 h exposure to honokiol was followed by a slight but significant increase of [Ca2+]i (15 lM), significant decrease of forward scatter (5 lM), significant increase of annexin-V-binding (5 lM) and significant increase of ceramide formation (15 lM). Honokiol further induced slight, but significant hemolysis. Honokiol (15 lM) induced annexin-V-binding was significantly blunted but not abrogated in the nominal absence of extracellular Ca2+. In conclusion, honokiol triggers suicidal erythrocyte death or eryptosis, an effect at least in part due to stimulation of Ca2+ entry and ceramide formation. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction The polyphenol honokiol [(3,5-di-(2-propenyl)-1,1-biphenyl2,2-diol], derived from Magnolia officinalis (Xu et al., 2011), has previously been shown to counteract inflammation, infection, anxiety, depression, stress, angiogenesis, oxidation and malignancy (Arora et al., 2012; Chen et al., 2011; Cheng et al., 2011; Lee et al., 2012; Lee et al., 2011; Li et al., 2012; Liu et al., 2011; Mannal et al., 2011; Ponnurangam et al., 2012; Shen et al., 2010; Steinmann et al., 2012; Wang et al., 2011; Weng et al., 2012; Xu et al., 2011). The anti-tumor activity of honokiol has been attributed to its ability to stimulate apoptosis (Arora et al., 2011; Chilampalli et al., 2011; He et al., 2011; Ishikawa et al., 2012; Jeong et al., 2012; Lin et al., 2012; Marin and Mansilla, 2010; Wang et al., 2010; Xu et al., 2011). On the other hand, honokiol has been shown to inhibit apoptosis (Hoi et al., 2010; Sheu et al., 2008; Tang et al., 2011). Mechanisms mediating effects of honokiol include death receptor pathways (Xu et al., 2011), mitochondria (Fried and Arbiser, 2009; Lin et al., 2012; Xu et al., 2011), caspases (Xu et al., 2011),

⇑ Corresponding author. Address: Physiologisches Institut, der Universität Tübingen, Gmelinstr. 5, D-72076 Tübingen, Germany. Tel.: +49 7071 29 72194; fax: +49 7071 29 5618. E-mail address: fl[email protected] (F. Lang). 1 Contributed equally and thus share first authorsip. 0887-2333/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.tiv.2013.05.003

apoptosis-related proteins (Arora et al., 2012), nuclear factor kappa B (NF-jB) (Arora et al., 2012), signal transducers and activator of transcription 3 (STAT3) (Arora et al., 2012; Rajendran et al., 2012; Yu et al., 2012), epidermal growth factor receptor (EGFR) (Arora et al., 2012; Leeman-Neill et al., 2010), mammalian target of rapamycin (m-TOR) (Arora et al., 2012), phospholipase D (Fried and Arbiser, 2009), Calpain (Liu et al., 2012), adhesion molecules (Jeong et al., 2012), 15-lipoxygenase-1 (Liu et al., 2010), cycloxygenase 2 (Liu et al., 2010), and peroxisome proliferator-activated receptor-gamma (Liu et al., 2010). Even though erythrocytes lack mitochondria and nuclei, key elements in the triggering and execution of apoptosis, they may undergo apoptosis-like suicidal death or eryptosis, which is characterized by breakdown of cell membrane phosphatidylserine asymmetry and cell shrinkage (Lang et al., 2008). Eryptosis may be elicited by increased cytosolic Ca2+ concentration ([Ca2+]i), which may result from Ca2+ entry through Ca2+-permeable cation channels (Foller et al., 2008c; Foller et al., 2009c). Ca2+ permeable erythrocyte cation channels are activated by oxidative stress (Brand et al., 2003). An increase of [Ca2+]i leads to activation of Ca2+-sensitive K+ channels (Brugnara et al., 1993) with subsequent cell shrinkage due to K+ exit, hyperpolarization, Cl exit and thus cellular KCl and water loss (Lang et al., 2003). An increase of [Ca2+]i further leads to breakdown of cell membrane phosphatidylserine asymmetry with subsequent phosphatidylserine exposure at the erythrocyte surface (Berg et al., 2001). Ca2+ sensitivity of

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phosphatidylserine translocation is increased by ceramide (Lang et al., 2010). Eryptosis is further triggered by energy depletion (Klarl et al., 2006) and caspase activation (Bhavsar et al., 2010; Foller et al., 2008b; Foller et al., 2009b; Lau et al., 2011; Maellaro et al., 2011). Several kinases participate in the regulation of eryptosis, such as AMP activated kinase AMPK (Foller et al., 2009c), cGMPdependent protein kinase (Foller et al., 2008a), Janus-activated kinase JAK3 (Bhavsar et al., 2011), casein kinase (Kucherenko et al., 2012; Zelenak et al., 2012a), p38 kinase (Gatidis et al., 2011), PAK2 kinase (Zelenak et al., 2011) as well as sorafenib (Lupescu et al., 2012d) and sunifinib (Shaik et al., 2012a) sensitive kinases. Eryptosis could be triggered by a wide variety of xenobiotics (Abed et al., 2012a; Abed et al., 2012b; Bottger et al., 2012; Felder et al., 2011; Firat et al., 2012; Ganesan et al., 2012; Gao et al., 2012; Ghashghaeinia et al., 2012; Ghashghaeinia et al., 2011; Jilani et al., 2012; Jilani et al., 2013; Kucherenko and Lang, 2012; Lang et al., 2011; Lang et al., 2012a; Lang et al., 2012b; Lang and Qadri, 2012; Lupescu et al., 2012a; Lupescu et al., 2012b; Lupescu et al., 2012c; Polak-Jonkisz and Purzyc, 2012; Qadri et al., 2011a; Qadri et al., 2011b; Qadri et al., 2011c; Qian et al., 2012; Shaik et al., 2012a; Shaik et al., 2012b; Vota et al., 2012; Weiss et al., 2012; Zappulla, 2008; Zbidah et al., 2012a; Zbidah et al., 2012b; Zelenak et al., 2012b) and is observed in several clinical disorders (Lang et al., 2008) including diabetes (Calderon-Salinas et al., 2011; Maellaro et al., 2011; Nicolay et al., 2006), renal insufficiency (Myssina et al., 2003), hemolytic uremic syndrome (Lang et al., 2006), sepsis (Kempe et al., 2007), malaria (Bobbala et al., 2010; Foller et al., 2009a; Koka et al., 2009; Lang et al., 2007; Siraskar et al., 2010), sickle cell disease (Lang et al., 2009), Wilson’s disease (Lang et al., 2007), iron deficiency (Kempe et al., 2006), malignancy (Qadri et al., 2012), phosphate depletion (Birka et al., 2004), and metabolic syndrome (Zappulla, 2008). The present study explored, whether honokiol influences [Ca2+]i, cell volume and phosphatidylserine abundance at the erythrocyte surface. The experiments revealed that treatment of human erythrocytes with honokiol increases cytosolic Ca2+ activity, leads to cell shrinkage and triggers the breakdown of phosphatidylserine asymmetry of the cell membrane. Thus, honokiol treatment stimulates eryptosis, the suicidal death of erythrocytes. 2. Materials and methods 2.1. Erythrocytes, solutions and chemicals Leukocyte-depleted erythrocytes were kindly provided by the blood bank of the University of Tübingen. The study is approved by the ethics committee of the University of Tübingen (184/ 2003V). Erythrocytes were incubated in vitro at a hematocrit of 0.4% in Ringer solution containing (in mM) 125 NaCl, 5 KCl, 1 MgSO4, 32 N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid (HEPES), 5 glucose, 1 CaCl2; pH 7.4 at 37 °C for 48 h. Where indicated, erythrocytes were exposed to honokiol (Enzo, Lörrach, Germany) at the indicated concentrations. Honokiol was dissolved in DMSO. The final concentration of DMSO in the samples did not exceed 0.15%. Ca2+ free Ringer solution was used to study the role of calcium entry in the effect of honokiol on breakdown of phosphatidylserine asymmetry. In Ca2+-free Ringer solution, 1 mM CaCl2 was substituted by 1 mM glycol-bis(2-aminoethylether)N,N,N0 ,N0 -tetraacetic acid (EGTA). The Ca2+ chelator EGTA was used to fully deplete extracellular Ca2+. Ca2+ ionophore ionomycin was used to increase cytosolic Ca2+ concentration. 2.2. FACS analysis of annexin-V-binding and forward scatter After incubation under the respective experimental condition, 50 ll cell suspension was washed in Ringer solution containing

5 mM CaCl2 and then stained with Annexin-V-FITC (1:200 dilution; ImmunoTools, Friesoythe, Germany) in this solution at 37 °C for 20 min under protection from light. In the following, the forward scatter (FSC) of the cells was determined, and annexin-V fluorescence intensity was measured with an excitation wavelength of 488 nm and an emission wavelength of 530 nm on a FACS Calibur (BD, Heidelberg, Germany). A total of 10,000–15,000 events were counted in each single experiment. Data acquisition and analysis were performed using the standard software CellQuest Pro (BD biosciences). For the analysis of annexin V fluorescence a marker was placed at the end of the first annexin V negative population and a batch analysis was run acquiring the percentage of annexin V positive erythrocytes using the CellQuest Pro software. For determination of forward scatter a batch analysis was performed to record the geometric mean of the cell population. 2.3. Determination of erythrocyte diameter For the determination of accurate erythrocyte diameter, the Flow Cytometer size calibration Kit (F13838) from Molecular Probes was used. Prior to sampling of the kit components, microspheres were suspended by vortex mixing and sonicating. The larger microspheres settled out within minutes. A mixed suspension of the six standard microspheres was prepared by adding one drop of each to approximately 2 mL of sheath fluid or buffered saline. Additionally, a subset of six separated suspensions with different diameters were prepared in the given experiment depending on the needed size markers. Suspended microspheres and blood samples were measured by flow cytometry. 2.4. Measurement of intracellular Ca2+ After incubation erythrocytes were washed in Ringer solution and then loaded with Fluo-3/AM (Biotium, Hayward, USA) in Ringer solution containing 5 mM CaCl2 and 2 lM Fluo-3/AM. The cells were incubated at 37 °C for 30 min and washed twice in Ringer solution containing 5 mM CaCl2. The Fluo-3/AM-loaded erythrocytes were resuspended in 200 ll Ringer. Then, Ca2+-dependent fluorescence intensity was measured with an excitation wavelength of 488 nm and an emission wavelength of 530 nm on a FACS Calibur (BD, Heidelberg, Germany). Cell Quest Pro software recorded the geometric mean of the samples using batch analysis. 2.5. Measurement of hemolysis For the determination of hemolysis the samples were centrifuged (3 min at 400 g, room temperature) after incubation, and the supernatants were harvested. As a measure of hemolysis, the hemoglobin (Hb) concentration of the supernatant was determined photometrically at 405 nm. The absorption of the supernatant of erythrocytes lysed in distilled water was defined as 100% hemolysis. 2.6. Confocal microscopy and immunofluorescence For the visualization of eryptotic erythrocytes, 4 ll of erythrocyte suspension, incubated under the respective experimental conditions, were stained with FITC-conjugated Annexin-V (1:100 dilution; ImmunoTools, Friesoythe, Germany) in 200 ll Ringer solution containing 5 mM CaCl2. Then the erythrocytes were washed twice and finally re-suspended in 50 ll of Ringer solution containing 5 mM CaCl2. Twenty microliter were mounted with Prolong Gold antifade reagent (Invitrogen, Darmstadt, Germany) onto a glass slide, covered with a coverslip, and images were subsequently taken on a Zeiss LSM 5 EXCITER confocal laser

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Fig. 1. Effect of honokiol on erythrocyte cytosolic Ca2+ concentration. (A) Original histogram of Fluo3 fluorescence in erythrocytes following exposure for 48 h to Ringer solution without () and with (+) presence of 15 lM honokiol. (B) Arithmetic means ± SEM (n = 15) of Fluo3 fluorescence (arbitrary units) in erythrocytes exposed for 48 h to Ringer solution without (white bar) or with (black bars) honokiol (1–15 lM). (p < 0.001) indicates significant difference from the absence of honokiol (ANOVA).

scanning microscope (Carl Zeiss MicroImaging, Oberkochen, Germany) with a water immersion Plan-Neofluar 63/1.3 NA DIC. 2.7. Determination of ceramide formation For the determination of ceramide, a monoclonal antibodybased assay was used. After incubation, cells were stained for 1 h at 37 °C with 1 lg/ml anti-ceramide antibody (clone MID 15B4, Alexis, Grünberg, Germany) in PBS containing 0.1% bovine serum

B Forward scatter (arb.units)

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albumin (BSA) at a dilution of 1:5. The samples were washed twice with PBS–BSA. Subsequently, the cells were stained for 30 min with polyclonal fluorescein-isothiocyanate (FITC)-conjugated goat anti-mouse IgG and IgM specific antibody (Pharmingen, Hamburg, Germany) diluted 1:50 in PBS–BSA. Unbound secondary antibody was removed by repeated washing with PBS–BSA. The samples were then analyzed by flow cytometric analysis. To rule out the effect of unspecific fluorescence emitted by the secondary antibody, fluorescence intensity was determined in the honokiol-treated

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Honokiol Fig. 2. Effect of honokiol on erythrocyte forward scatter and diameter. (A) Original histogram of forward scatter of erythrocytes following exposure for 48 h to Ringer solution without () and with (+) presence of 15 lM honokiol. (B) Arithmetic means ± SEM (n = 15) of the erythrocyte forward scatter (FSC) following incubation for 48 h to Ringer solution without (white bar) or with (black bars) honokiol (1–15 lM). (p < 0.001) indicates significant difference from the absence of honokiol (ANOVA). (C) Arithmetic means ± SEM (n = 4) of the erythrocyte diameter following incubation for 48 h to Ringer solution without (white bar) or with (black bar) honokiol (15 lM). Cell size was determined utilizing sizing beads. (p < 0.001) indicates significant difference from the absence of honokiol (t test).

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erythrocytes without primary antibody but in the presence or absence of secondary antibody. The mean fluorescence in honokioltreated erythrocytes was 20.1 ± 0.6 (a.u., n = 4) in the presence of secondary antibody and 20.1 ± 0.7 (a.u., n = 4) in the absence of secondary antibody.

fluorescence. Accordingly, only higher concentrations of honokiol significantly and appreciably increased cytosolic Ca2+ concentration. As a positive control we measured Fluo3 fluorescence in the presence of the calcium ionophore ionomycin. Erythrocytes treated with 1 lM of ionomycin significantly enhanced erythrocyte Fluo3 fluorescence from 21 ± 0.9 to 61 ± 2.0 (a.u., n = 4).

2.8. Statistics 3.2. Honokiol induced cell shrinkage Data are expressed as arithmetic means ± SEM. As indicated in the figure legends, statistical analysis was made by ANOVA with Tukey’s test as post-test and t test as appropriate using GraphPad InStat Version 3.06 (San Diego, CA, USA). n denotes the number of different erythrocyte specimens studied. Since different erythrocyte specimens used in distinct experiments are differently susceptible to triggers of eryptosis, the same erythrocyte specimens were been used for control and experimental conditions.

A hallmark of eryptosis is cell shrinkage. To estimate the effect of honokiol on the volume of erythrocytes, forward scatter was determined utilizing FACS analysis. As illustrated in Fig. 2, a 48 h exposure to honokiol was followed by a decrease of forward scatter, an effect reaching statistical significance at 5 lM honokiol concentration. Accordingly, honokiol decreased erythrocyte volume. To further define the effect of honokiol on cell volume, the accurate diameter of erythrocytes was determined with or without prior honokiol treatment. As shown in Fig 2C, honokiol (15 lM) treatment significantly decreased the erythrocyte diameter.

3. Results 3.1. Honokiol increased cytosolic Ca2+ activity

3.3. Honokiol induced loss of phosphatidylserine asymmetry of the cell membrane

Eryptosis is triggered by increase of cytosolic Ca2+ activity ([Ca2+]i). In order to determine [Ca2+]i of human erythrocytes, the erythrocytes were incubated in Ringer solution without or with honokiol (1–15 lM). The erythrocytes were subsequently loaded with Fluo3-AM and Fluo3 fluorescence determined in FACS analysis. As illustrated in Fig. 1, following a 48 h exposure of human erythrocytes up to 10 lM honokiol remained without significant effect on Fluo3 fluorescence. Only exposure to 15 lM honokiol was followed by a slight but significant increase of Fluo3

A

A further hallmark of eryptosis is breakdown of cell membrane phosphatidylserine asymmetry. Thus, phosphatidylserine abundance at the cell surface was determined. Phosphatidylserine exposing erythrocytes were identified by annexin-V-binding in FACS analysis. As shown in Fig. 3, a 48 h exposure to honokiol increased the percentage of annexin-V-binding erythrocytes, an effect reaching statistical significance at 5 lM honokiol

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Time (h) Fig. 3. Effect of honokiol on phosphatidylserine exposure and hemolysis. (A) Original histogram of annexin-V-binding of erythrocytes following exposure for 48 h to Ringer solution without () and with (+) presence of 15 lM honokiol. (B) Arithmetic means ± SEM (n = 15) of erythrocyte annexin-V-binding following incubation for 48 h to Ringer solution without (white bar) or with (black bars) presence of honokiol (1–15 lM). For comparison, arithmetic means ± SEM (n = 4) of the percentage of hemolysis is shown as grey bars. (p < 0.05), (p < 0.001), indicate significant differences from the absence of honokiol (ANOVA). (C) Arithmetic means ± SEM (n = 4) of erythrocyte annexin-Vbinding following incubation for 6–48 h to Ringer solution without (white triangles) or with (black squares) honokiol (15 lM). (p < 0.05), (p < 0.001) indicates significant difference from the absence of honokiol (t test).

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0 µM Honokiol

15 µM Honokiol

Fig. 4. Confocal images of phosphatidylserine exposing erythrocytes with or without honokiol treatment. Light microscopy (upper panels) and confocal microscopy of FITCdependent fluorescence (lower panels) of human erythrocytes stained with FITC-conjugated Annexin-V-Fluos following a 48 h incubation in Ringer solution without (left panels) and with (right panels) 15 lM honokiol.

3.4. Honokiol induced hemolysis Access of fluorescent annexin V to phosphatidylserine could result from permeabilization of erythrocytes by hemolysis. Thus, the percentage of hemolysed erythrocytes was quantified by determination of hemoglobin release into the supernatant. As illustrated in Fig. 3, exposure of erythrocytes for 48 h to honokiol increased the hemoglobin concentration in the supernatant, an effect, however, affecting only a relatively small percentage of erythrocytes (Fig. 3 B). 3.5. Ca2+ sensitivity of honokiol induced phosphatidylserine translocation Further experiments aimed to define the mechanisms accounting for the breakdown of phosphatidylserine asymmetry of the cell membrane following honokiol treatment. In order to quantify the importance of extracellular Ca2+, erythrocytes were exposed to 15 lM honokiol for 48 h in the absence of 1 mM Ca2+ and presence of the Ca2+ chelator EGTA (1 mM). As illustrated in Fig. 5, the effect of honokiol on annexin-V-binding was significantly decreased in the nominal absence of Ca2+. However, removal of extracellular

9

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PS-exposing erythrocytes (%)

concentration. Accordingly, honokiol exposure was followed by breakdown of phosphatidylserine asymmetry of the cell membrane with phosphatidylserine exposure at the cell surface. As honokiol was dissolved in DMSO, the effect of the solvent on phosphatidylserine abundance was studied. Exposure of erythrocytes to 0.15% DMSO did not significantly modify phosphatidylserine abundance. In the absence of DMSO the phosphatidylserine abundance was 2.6 ± 0.5% and in the presence of DMSO 2.8 ± 0.4% (n = 4). The time course of annexin V binding following honokiol treatment is shown in Fig. 3C. In order to visualize annexin V binding erythrocytes, confocal imaging of erythrocytes was done without and with prior honokiol treatment (Fig. 4).

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Fig. 5. Effect of Ca2+ withdrawal on honokiol-induced annexin-V-binding. Arithmetic means ± SEM (n = 4) of the percentage of annexin-V-binding erythrocytes after a 48 h treatment with Ringer solution without (white bars) or with (black bars) 15 lM honokiol in the presence (left bars, + Calcium) and absence (right bars, Calcium) of Ca2+. (p < 0.001) indicates significant difference from the absence of honokiol (ANOVA) ##(p < 0.01) indicates significant difference from the respective values in the presence of Ca2+.

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###

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80 70 60 50

following honokiol treatment. Ca2+ sensitivity of phosphatdidylserine translocation is enhanced by ceramide, which thus triggers phosphatidylserine translocation even without increase of cytosolic Ca2+ activity. Thus, additional experiments were performed to test, whether honokiol treatment increases formation of ceramide. Ceramide abundance at the cell surface was determined utilizing FITC-labeled anti-ceramide antibodies. As illustrated in Fig. 7, honokiol significantly increased ceramide-dependent fluorescence.

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Fig. 6. Effect of ionomycin on honokiol-induced annexin-V-binding. Arithmetic means ± SEM (n = 4) of the percentage of annexin-V-binding erythrocytes after a 1 h treatment with Ringer solution without (white bar) or with (black bars) 15 lM honokiol in the absence (left bars) and presence (right bars) of ionomycin (1 lM). ⁄, (p < 0.001) indicate significant difference from the absence of honokiol (ANOVA) ### (p < 0.001) indicates significant difference from the respective values in the absence of ionomycin.

Ca2+ did not abrogate the effect of honokiol on phosphatidylserine abundance. In order to test, whether honokiol modified the breakdown of phosphatidylserine asymmetry by increase of cytosolic Ca2+ activity, the effect of the Ca2+ ionophore ionomycin on phosphatidylserine abundance was tested in the absence and presence of honokiol. As shown in Fig. 6, the phosphatidylserine abundance was significantly higher in erythrocytes treated with both, 15 lM honokiol and 1 lM ionomycin than in erythrocytes treated with honokiol or ionomycin alone.

3.6. Honokiol induced ceramide formation Removal of extracellular Ca2+ did not fully abrogate the breakdown of phosphatidylserine asymmetry of the cell membrane

The present observations reveal the ability of honokiol to elicit suicidal death of erythrocytes. Exposure of human erythrocytes to honokiol resulted in erythrocyte shrinkage and erythrocyte membrane scrambling. The concentrations required (5 lM) were similar to those (10 lM) triggering apoptosis of tumor cells (Jeong et al., 2012). Lower concentrations of honokiol (0.125–1 lM) were shown to counteract apoptosis (Sheu et al., 2008). Following in vivo administration of 250 mg/kg plasma honokiol concentration increased up to 1 g/l ( 4 mM) (Chen et al., 2004). The therapeutic plasma concentration was approximately 10 mg/l ( 40 lM) (Chen et al., 2004; Jiang et al., 2008). Thus, the concentrations shown to trigger eryptosis are indeed relevant in vivo. Honokiol induced erythrocyte shrinkage may have resulted from activation of Ca2+ sensitive K+ channels (Bookchin et al., 1987; Brugnara et al., 1993) with subsequent K+ exit, cell membrane hyperpolarisation, Cl exit and thus cellular loss of KCl with osmotically obliged water (Lang et al., 2003). Honokiol induced phosphatidylserne translocation required the presence of extracellular Ca2+. Cytosolic Ca2+ activity ([Ca2+]i) may be increased in erythrocytes by activating Ca2+ permeable non-selective cation channels, which involve the transient receptor potential channel TRPC6 (Foller et al., 2008c). The channels are activated by oxidative stress (Brand et al., 2003). However, somewhat surprisingly, in the present study high concentrations (15 lM) of honkiol were required to appreciably increase [Ca2+]i. Thus, at lower honokiol concentrations, the effect of honokiol on phosphatidylserine translocation may require the presence of Ca2+ but not necessarily an increase of [Ca2+]i. Ca2+ sensitivity of phosphatidylserine translocation could be enhanced by ceramide (Lang et al., 2010). As a matter of fact, honokiol treatment resulted in an increase of ceramide formation. Ceramide

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Fig. 7. Effect of honokiol on ceramide formation. (A) Original histogram of anti-ceramide FITC-fluorescence in erythrocytes following exposure for 48 h to Ringer solution without () and with (+) presence of 15 lM honokiol. (B) Arithmetic means ± SEM (n = 4) of ceramide abundance after a 48 h incubation in Ringer solution without (white bar) or with (black bar) honokiol (15 lM). (p < 0.05) indicates significant difference from control (absence of honokiol) (t test).

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is known to trigger breakdown of cell membrane phosphatidylserine asymmetry in erythrocytes (Lang et al., 2010). In vivo, eryptosis is followed by clearance of eryptotic erythrocytes from circulating blood (Lang et al., 2008). The accelerated loss of erythrocytes may be compensated by enhanced formation of new erythrocytes and thus remain without appreciable effect on blood count. As soon as eryptosis exceeds the formation of new erythrocytes, anemia develops (Lang et al., 2008). Eryptosis may further compromise microcirculation because phosphatidylserine exposing erythrocytes adhere to endothelial CXCL16/SR-PSO of the vascular wall (Borst et al., 2012). At least in theory, the adhering erythrocytes may interfere with blood flow (Andrews and Low, 1999; Borst et al., 2012; Closse et al., 1999; Gallagher et al., 2003; Pandolfi et al., 2007; Wood et al., 1996). Moreover, phosphatidylserine exposure may foster blood clotting and thus thrombosis (Andrews and Low, 1999; Chung et al., 2007; Zwaal et al., 2005). In conclusion, honokiol triggers breakdown of cell membrane phosphatidylserine asymmetry and cell shrinkage and thus suicidal death of human erythrocytes. The effect is in large part dependent on the presence of extracellular Ca2+. Acknowledgements The authors acknowledge the meticulous preparation of the manuscript by Ali Soleimanpour. The study was supported by the Deutsche Forschungsgemeinschaft. References Abed, M., Towhid, S.T., Mia, S., Pakladok, T., Alesutan, I., Borst, O., Gawaz, M., Gulbins, E., Lang, F., 2012a. Sphingomyelinase-induced adhesion of eryptotic erythrocytes to endothelial cells. Am. J. Physiol. Cell Physiol. 303, C991–C999. Abed, M., Towhid, S.T., Shaik, N., Lang, F., 2012b. Stimulation of suicidal death of erythrocytes by rifampicin. Toxicology 302, 123–128. Andrews, D.A., Low, P.S., 1999. Role of red blood cells in thrombosis. Curr. Opin. Hematol. 6, 76–82. Arora, S., Bhardwaj, A., Srivastava, S.K., Singh, S., McClellan, S., Wang, B., Singh, A.P., 2011. Honokiol arrests cell cycle, induces apoptosis, and potentiates the cytotoxic effect of gemcitabine in human pancreatic cancer cells. PLoS One 6, e21573. Arora, S., Singh, S., Piazza, G.A., Contreras, C.M., Panyam, J., Singh, A.P., 2012. Honokiol: a novel natural agent for cancer prevention and therapy. Curr. Mol. Med. 12, 1244–1252. Berg, C.P., Engels, I.H., Rothbart, A., Lauber, K., Renz, A., Schlosser, S.F., SchulzeOsthoff, K., Wesselborg, S., 2001. Human mature red blood cells express caspase-3 and caspase-8, but are devoid of mitochondrial regulators of apoptosis. Cell Death. Differ. 8, 1197–1206. Bhavsar, S.K., Bobbala, D., Xuan, N.T., Foller, M., Lang, F., 2010. Stimulation of suicidal erythrocyte death by alpha-lipoic acid. Cell. Physiol. Biochem. 26, 859– 868. Bhavsar, S.K., Gu, S., Bobbala, D., Lang, F., 2011. Janus kinase 3 is expressed in erythrocytes, phosphorylated upon energy depletion and involved in the regulation of suicidal erythrocyte death. Cell. Physiol. Biochem. 27, 547– 556. Birka, C., Lang, P.A., Kempe, D.S., Hoefling, L., Tanneur, V., Duranton, C., Nammi, S., Henke, G., Myssina, S., Krikov, M., Huber, S.M., Wieder, T., Lang, F., 2004. Enhanced susceptibility to erythrocyte ‘‘apoptosis’’ following phosphate depletion. Pflugers Arch. 448, 471–477. Bobbala, D., Alesutan, I., Foller, M., Huber, S.M., Lang, F., 2010. Effect of anandamide in plasmodium Berghei-infected mice. Cell. Physiol. Biochem. 26, 355–362. Bookchin, R.M., Ortiz, O.E., Lew, V.L., 1987. Activation of calcium-dependent potassium channels in deoxygenated sickled red cells. Prog. Clin. Biol. Res. 240, 193–200. Borst, O., Abed, M., Alesutan, I., Towhid, S.T., Qadri, S.M., Foller, M., Gawaz, M., Lang, F., 2012. Dynamic adhesion of eryptotic erythrocytes to endothelial cells via CXCL16/SR-PSOX. Am. J. Physiol. Cell Physiol. 302, C644–C651. Bottger, E., Multhoff, G., Kun, J.F., Esen, M., 2012. Plasmodium falciparum-infected erythrocytes induce granzyme B by NK cells through expression of host-Hsp70. PLoS One 7, e33774. Brand, V.B., Sandu, C.D., Duranton, C., Tanneur, V., Lang, K.S., Huber, S.M., Lang, F., 2003. Dependence of Plasmodium falciparum in vitro growth on the cation permeability of the human host erythrocyte. Cell. Physiol. Biochem. 13, 347– 356. Brugnara, C., de Franceschi, L., Alper, S.L., 1993. Inhibition of Ca(2+)-dependent K+ transport and cell dehydration in sickle erythrocytes by clotrimazole and other imidazole derivatives. J. Clin. Invest. 92, 520–526.

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