Role of intracellular Ca2+ on the formation of microvesicle in human red blood cells - Nguyen Duc Bach1

TAP CHI SINH HOC 2015, 37(1se): 67-74 DOI: 10.15625/0866-7160/v37n1se. ROLE OF INTRACELLULAR CA2+ ON THE FORMATION OF MICROVESICLE IN HUMAN RED BLOOD CELLS Nguyen Duc Bach1*, Nguyen Huu Duc1, Pham Kim Dang1 Luu Thao Linh1, Ly Thi BichThuy2, Ingolf Bernhardt3 1Vietnam National University of Agriculture, *ndbach@gmail.com 2Institute of Biotechnology 3Saarland University, Germany ABSTRACT: Release of vesicles from various cell types has been seen as a kind of intercellular communication. So far three types of vesicles have been described including exosomes (40-100 nm), microvesicles (100-1,000 nm) and apoptotic bodies (1-4 mm). Although the mechanisms for the formation of vesicles are unclear at the moment, microvesicles (MVs) have been reported as a mean of transport for lipids, growth factors, protein, RNA and signal molecules among cells. In human, red blood cells (RBCs) are the most common type of blood cell which deliver oxygen to the body tissues. The formation and release of MVs from RBCs have been reported for years. However, the mechanism for the formation and function of MVs released from RBCs has not well understood. In this study, conditions for the formation and release of MVs from human (RBCs) have been investigated. By using fluorescence label techniques, fluorescence microscope and flow cytometer. The results showed that phorbol-12-myristat-13-acetate (PMA), lysophosphatidic acid (LPA) and calcium inonophore (A23187) induced the increase of intracellular Ca2+. Increase of intracellular of Ca2+ led to the formation, shedding and release of MVs in human RBCs. The formation of MVs was also coupled with the externalization of phosphatidyl serine (PS). The kinetics of the formation of MVs was investigated by using annexin V-FITC. The obtained results showed that increase intracellular Ca2+ led to cell shrinkage, formation and release of MVs in human RBCs. Keywords: Fluorescence microscope, intracellular Ca2+, microvesicles, phosphatidyl serine, red blood cell. INTRODUCTION Release of vesicles is a process observed in many eukaryotic cell types. Based on their size, 3 types of vesicles have been classified, exosomes (40 -100 nm), microvesicles (MVs) or membrane particles (0.1-1.0 µm) and apoptotic bodies (1.0 -4.0 mm) [9, 15, 18]. Exosomes have endogenous origins, which are released from membrane structures of organels in cytosol into the extracellular environment. So far, exosomes have been found in almost cell types including blood plasma, urine and milk [10, 19, 25]. Apoptosis is considered as a component of various processes including normal cell turnover, proper development and functioning of the immune system, hormone-dependent atrophy, embryonic development and chemical-induced cell death [3, 8,16]. Apoptotic body can be seen as large parts of dead cell covered by membrane. Unlike exosomes and apoptotic bodies, MVs are membrane derivated structures which are formed and shed from cellular membrane [9, 12]. In blood plasma, the size of MVs has been reported in the range of 100-400 nm [23]. Recently, MVs have proved as a mediate intercellular communication through the carry and transfer of cellular information including RNAs (both coding and non-coding RNAs), proteins and other signal molecules to recipient cells [20, 22]. Therefore, MVs may involve in the regulation of normal physiological as well as pathological processes. It has been demonstrated that an increase of intracellular Ca2+ is a very important factor leading to the activation of the Gardos channel, scramblase, calcium-dependent protease calpain, inhibition of amino phospholipid translocase and ceramide formation [21, 26]. The consequences of these processes are cell shrinkage, phosphatidyl serine (PS) exposure and the breakdown of cell membrane structure to form MVs [12, 14]. The release and formation of MVs from platelet has been described by the increase of intracellular Ca2+ [1, 17]. Recently, the release of MVs in human RBCs has been investigated and they are thought to involve the formation of the blood clots and phagocytosis of apoptotic cells [3, 4]. However, the mechanism for the formation and release of MVs as well as their roles has not been well described. In this study, conditions for the formation of MVs have been investigated by using calcium inonophore A32187, lysophosphatidic acid (LPA). MATERIALS AND METHODS Red blood cell preparation Human venous blood was drawn from healthy donors. EDTA was used as anticoagulant. Whole blood was centrifuged at 2,000g for 5 min at room temperature. The plasma was removed by aspiration. Subsequently, the RBCs were washed 3 times in physiological solution (NaCl 140 mM, KCl 7.5 mM, HEPES 10 mM, glucose 10 mM, pH 7.4) to remove the buffy coat. The diluted blood was gently layered onto lymphoprepTM (purchased from AXIS-SHIELD) and centrifuged at 400g for 30 min at room temperature. RBCs have a higher density so they are in the lower layer while peripheral blood mononuclear cells are in the upper layer. RBCs then collected and washed 3 times in physiological solution and stored at 4°C for experiments. Intracellular Ca2+ measurement The washed RBCs were suspended in physiological solution at 1% haematocrit with Fluo-4, AM, cell permeant (F-14201, Life technology) at final concentration of 2.5 µM. Fluo-4, AM was prepared as stock solution in DMSO at 0.5 mM. The cell suspension was mixed by vortexing and incubated for 45 min at 37°C with occasionally shaking. Subsequently, the cells were washed 3 times with the physiological solution by quick centrifugation at 12,000 g and re-suspended in physiological solution (haematocrit 0.5%). The intracellular Ca2+ in RBCs wasmeasured by fluorescence microscope (Eclipse TE 2000 E, Nikon) at room temperature. In all experiments, the extracellular Ca2+ concentration was added in physiological solution at 2 mM. Influence of LPA, PMA and A23187 on the level of intracellular Ca2+ was evaluated by measurement of fluo-4 fluorescence signal. The control experiments were performed under the same conditions in the absence of LPA, PMA or A23187. For intracellular Ca2+ measurement, the fluo-4 loaded RBCs were suspended in the physiological solution containing 2 mM Ca2+ and applied on a cover slip. When RBCs were settled down and kept in focus, A23187, LPA or PMA was gently added to final concentrations of 2.0, 2.5 or 6 µM, respectively. The measurements were carried out immediately after adding these substances. The time was set for the starting point. The excitation and emission wavelength were used at 488 nm and 530/15 nm, respectively. The fluorescence signals were normalized with background correction. Experiments were carried out with at least three different bloods from healthy donors. The data were analysed using MetaVue software. In parallel with the fluorescence microscope, intracellular Ca2+ content was also measured using a flow cytometer/fluorescence activated cell sorter (FACS) (BD FACSCalibur™). Parameters for flow cytometer analysis were adjusted using a calibration bead kit (BD Calibrite™ ). Determination of the shedding microvesicles The formation of MVs was observed under transmission light microscope at 600 times of magnification. The formation and shedding of MVs on the outer leaflet of the RBC membrane was also visualized by using fluorescence microscope (Eclipse TE 2000 E, Nikon). RBCs were stained with Annexin V, FITC conjugate (A13199, Life technology) in Annexin binding buffer (NaCl 145 mM, HEPES 10 mM CaCl2 2.5 mM, pH 7.4). A volume of 500 µl of annexin binding buffer and 5 µl of annexin V-FITC were added into each sample. The excitation and emission wavelength of annexin V-FITC are 488 and 530 nm, respectively. For quantitative analysis, number of RBCs showing positive signal with Annexin V- FITC also was measured by flow cytometer (Becton dickinson/BD) using a 530/30 nm band pass filter. For each measurement, 30,000 RBCs were counted. RBCs showing positive signal ofannexin V-FITC can be calculated in percentage by comparing positive and negative events with the control. BD CellQuest™ Pro Software was used for data acquisition and analysis. RESULTS AND DISCUSSION Measurement of of intracellular Ca2+ Figure 1. Kinetic measurement of intracellular Ca2+ in RBCs RBCs were suspended in physiological solutions containing 2 mM Ca2+. The experiments were started immediately after adding A23187, LPA or PMA. a: The Ca2+ flux in physiological solution (control); b: in the presence of 2 µM A23187; c: in the presence of 6 µM PMA; d: in the presence of 2.5 µM LPA. Each single curve represents one single cell. The level of intracellular Ca2+ concentration in RBCs was measured based on the signal of fluo-4. In control experiment, the values of fluorescence intensity in single cells were stable during 30 min (fig. 1a). Three substances A23187, LPA and PMA were used for these experiments. Calcium ionophore (A23187) acts as a divalent cation ionophore, allowing these ions to cross cell membranes. Lysophosphatidic acid (LPA) is a phospholipid derivative that can act as a signaling molecule. LPA a water-soluble lipid second messenger is released from activated platelets fibroblasts, adipocytes, and cancer cells [7]. The most common phorbol ester is 12-O-tetradecanoylphorbol-13-acetate also called phorbol-12-myristate-13-acetate (PMA), which has been used as an activator for protein kinase C family [4]. In the presence of 2 µM A23187, the increase of intracellular Ca2+ took place after 5 min of incubation (fig. 1b). At lower concentrations of A23187, the delay time was from 15 to 20 min. At concentrations from 100 µM to 5 mM extracellular Ca2+, there is no significant difference in both delay time and fluorescence intensity (data not shown). In average, the highest fluorescence intensity reached after 15 min. In the presence of 6 µM PMA and 2.5 µM LPA, a very fast influx of Ca2+ was observed just after adding these substances (fig. 1b, 1c, 1d). Data analysis showed that an increase of Ca2+ influx that can be observed just after 45 to 60 seconds after a treatment of RBCs with 2.5 µM LPA (fig. 2). The lag time was from 45 to 60 seconds depending on the bloods. An increase of the LPA concentration from 2.5 µM to 5.0 µM does not change the delay time but the number of haemolysed RBCs is significantly increased. To compare the level of intracellular Ca2+ in RBCs treated with A23187, LPA and PMA, an overlay histogram of fluorescence intensity was created. The data showed that the highest level of intracellular Ca2+ was observed in RBCs treated with A23187 and LPA (fig. 3). This can be explained based on the difference in acting mechanism of these substances. In case of LPA, the Ca2+ influx should be due to a channel mediated transport rather than a leak transport. LPA is believed to bind to a G protein-coupled receptor that activates a C-type phospholipase that in turn generates diacylglycerol and 1,4,5-inositol trisphosphate [11, 24]. It has been also demonstrated that LPA opens the non-selective voltage dependent cation channel in human RBCs [4, 11]. For PMA, this substance activates protein kinase C and promotes the uptake of Ca2+ into the cell. The ω-agatoxin-TK-sensitive, Cav2.1-like (P/Q-type) Ca2+ channel is present in the RBC membrane and it may function under the control of kinases and phosphatases. This Ca2+ channel is responsible for the uptake of Ca2+ into RBCs in the presence of PMA [2]. Figure 2. Kinetics of the Ca2+ uptake in RBCs in the presence of LPA RBCs were suspended in physiological solutions containing 2 mM Ca2+. The kinetic experiments were started immediately after adding LPA at final concentration of 2.5 µM LPA. Figure 3. Overlay histogram of the intracellular Ca2+ in RBCs Original FACS data showing the overlay histogram of fluo-4 signal for intracellular Ca2+ in RBCs treated with A23187, LPA and PMA. 1: control in physiological solution containing 2 mM Ca2+, 2: 6 µM PMA, 3: 2.5 µM LPA, 4: 2 µM A23187. M1 and M2 represent the number of the negative and positive events. For each measurement, 30,000 events were analysed. Figure 4. Reduction of the cell volume of RBCs Original FACS data showing the side scatter vs. forward scatter. Left: RBCs were suspended in physiological solution containing 2 mM Ca2+ (control). Right: RBCs were treated with 2 µM A23187 for 15 min incubation at room temperature. Formation of microvesicles Treatment of RBCs with A23187, PMA or LPA led to increase of intracellular Ca2+. An increase of intracellular Ca2+ led to the reduction of cell volume (fig. 4). In this figure, a decrease of forward scatter value reflected the reduction of the size of RBCs. In the presence of A23187, PMA or LPA, the formation of MVs in RBCs was observed. In this paper, only the formation of MVs induced by 6 µM PMA for 120 min was demonstrated in figure 5. Recently, elevation of the intracellular Ca2+ activates different cellular processes such as calcium dependent scramblase, calpain, ceramide formation [4, 12, 21, 26]. The consequences are cell shrinkage, membrane blebbing, PS exposure, and the formation of microvesicles [4, 13]. In this study, the formation of MVs can be explained by the activation of phospholipid scramblases leading to disrupt the asymmetric distribution of phospholipids in the membrane and the externalization of PS. In addition the increase of cytosolic calcium ions also activate the calcium-dependent protease calpain leading to the breakdown of cellular skeleton and the shedding of membrane structure [4, 9, 11]. The consequence of these processes is the formation and release of MVs. Figure 5. The formation and release of microvesicles of RBCs Left: RBCs were suspended in physiological solutions containing 2 mM Ca2+ (control). Right: RBCs were suspended in physiological solutions containing 2 mM Ca2+ and 6 µM of PMA. The cells were observed under light transmission microscope at 600 times of magnification. The MVs were formed and shed from cell membrane of RBCs. Figure 6. Kinetics of the formation and release of MVs RBCs were suspended in physiological solution containing 2 mM Ca2+, annexin V-FITC in the presence of 2 µM A23187. Experiment was measured continuously for 120 min under fluorescence microscope (Eclipse TE 2000 E, Nikon). A detail of the kinetic process of formation of MVs in human RBCs by the action of A23187 was presented in figure 6. In this figure, cell shrinkage, membrane blebbing and microvesicles formation can be clearly observed. Similar results with PMA or LPA were also observed. However, in the absence of Ca2+ and 1 mM EGTA (no influx of Ca2+) treatment of RBCs with 6 µM of PMA also led to the formation of MVs in small number of RBCs (data now shown). It means that the formation of MVs in RBCs may also depend on other pathway(s). Kinetics of microvesicle formation In this study, the kinetics of microvesicles formation was measured by using fluorescence microscope. The exposed PS on the outer leaflet of membrane can be detected by using annexin V-FITC. Annexin V-FITC is a protein conjugated with fluorescein isothiocyanate which specifically binds to PS with very high affinity. The formation of MVs has been reported in couple with the externalization of MVs [12, 26]. Phosphatidyl serine (PS), a phospholipid component located in the inner leaflet of membrane is transported to outerleaflet by the activity of scramblase [5, 6]. The formation of MVs when RBCs treated with A23187 was presented in figure 6. After 30 min of measurement using A23187, some RBCs showing PS exposure and a small amount of microvesicles were observed. In this figure, the formation and release of MVs was clearly observed after 60 min and 120 min of experiment. It is necessary to mention that in artificial liposomes, lipids form symmetrical and stable bilayers with a random spontaneous transbilayer lipid diffusion between both leaflets [6]. However, lipids in biological membranes are asymmetrically distributed across the bilayer. The choline-containing lipids, phosphatidyl choline and sphingo myelin are enriched on the external leaflet of the plasma membrane. In contrast, the amine-containing glycerophospholipids, phosphatidyl ethanolamine and PS, are located preferentially on the cytoplasmic leaflet [5]. Therefore, the exposure of PS on the outer leaflet of RBCs and on MVs detected by annexin V-FITC (fig. 6) was an evidence for the formation and release of MVs from RBCs. By comparison with A23187, the process of MVs formation in the presence of LPA and PMA took place faster within 60 to 90 minutes of incubation (data not shown). In case of LPA, the number of RBCs haemolysed increased significantly after 60 min. The results confirmed that an elevation of intracellular Ca2+stimulated the formation and release of MVs from RBCs. CONCLUSION Treatment of human RBCs with A23187, LPA and PMA leds to an increase of intracellular Ca2+, formation and release of MVs. By using fluorescence labelling technique, fluorescence microscope and flow cytometer, the intracellular Ca2+ and kinetics of MVs formation in RBCs were investigated. The obtained results were useful for underlying the biological conditions and the mechanisms leading to the formation of MVs in RBCs in health and diseases. Based on obtained results, MVs released from RBCs can be collected for further characterization and functional study. Acknowledgements: This research is funded by Vietnam National Foundation for Science and Technology Development (NAFOSTED) under grant number 106-YS.06-2013.16 and The German Academic Exchange Service (DAAD). REFERENCES Aatonen M. T., Ohman T., Nyman T. A., Laitinen S., Gronholm M., Siljander P. R., 2014. Isolation and characterization of platelet-derived extracellular vesicles. Journal of extracellular vesicles. DOI: 10.3402/jev.v3.24692. Andrews D. A., Yang L., Low P. S., 2002. Phorbol ester stimulates a protein kinase C-mediated agatoxin-TK-sensitive calcium permeability pathway in human red blood cells. Blood., 100(9): 3392-3399. Antwi-Baffour S., Kholia S., Aryee Y. K., Ansa-Addo E. A., Stratton D., Lange S., 2010. Human plasma membrane-derived vesicles inhibit the phagocytosis of apoptotic cells-possible role in SLE. Biochem Biophys Res Commun., 398(2): 278-283. Bach N. D., Wagner-Britz L., Maia S., Steffen P., Wagner C., Kaestner L., 2011. Regulation of phosphatidylserine exposure in red blood cells. Cell Physiol Biochem., 28(5): 847-856. Daleke D. L., 2003. Regulation of transbilayer plasma membrane phospholipid asymmetry. J. Lipid Res., 44(2): 233-242. Daleke D. L., 2008. Regulation of phospholipid asymmetry in the erythrocyte membrane., Curr. Opin. Hematol., 15(3): 191-195. Eichholtz T., Jalink K., Fahrenfort I., Moolenaar W. H., 1993. The bioactive phospholipid lysophosphatidic acid is released from activated platelets. The Biochemical Journal, 291(3): 677-680. Elmore S., 2007. Apoptosis: a review of programmed cell death. Toxicol. Pathol., 35(4): 495-516. Gyorgy B., Szabo T. G., Pasztoi M., Pal Z., Misjak P., Aradi B., 2011. Membrane vesicles, current state-of-the-art: emerging role of extracellular vesicles. Cell Mol Life Sci., 68(16): 2667-2688. Huebner A. R., Somparn P., Benjachat T., Leelahavanichkul A., Avihingsanon Y., Fenton R. A., 2015. Exosomes in urine biomarker discovery. Adv. Exp. Med. Biol., 845: 43-58. Kaestner L., Steffen P., Nguyen D. B., Wang J., Wagner-Britz L., Jung A., 2012. Lysophosphatidic acid induced red blood cell aggregation in vitro. Bioelectrochemistry, 87: 89-95. Lee S. H., Meng X. W., Flatten K. S., Loegering D. A., Kaufmann S. H., 2013. Phosphatidylserine exposure during apoptosis reflects bidirectional trafficking between plasma membrane and cytoplasm. Cell Death Differ., 20(1): 64-76. Lee T. H., D'Asti E., Magnus N, Al-Nedawi K., Meehan B., Rak J., 2011. Microvesicles as mediators of intercellular communication in cancer-the emerging science of cellular 'debris'. Semin Immunopathol., 33(5): 455-467. Lee Y., El Andaloussi S., Wood M. J., 2012. Exosomes and microvesicles: extracellular vesicles for genetic information transfer and gene therapy. Hum Mol Genet, 21(R1): R125-R134. Ludwig A. K., Giebel B., 2012. Exosomes: small vesicles participating in intercellular communication. Int J. Biochem. Cell Biol., 44(1): 11-15. Ouyang L., Shi Z., Zhao S., Wang F. T., Zhou T. T., Liu B., 2012. Programmed cell death pathways in cancer: a review of apoptosis, autophagy and programmed necrosis. Cell Prolif., 45(6): 487-498. Pasquet J. M., Dachary-Prigent J., NurdenA. T., 1998. Microvesicle release is associated with extensive protein tyrosine dephosphorylation in platelets stimulated by A23187 or a mixture of thrombin and collagen. The Biochemical journal., 333(3): 591-599. Raposo G., Stoorvogel W., 2013. Extracellular vesicles: exosomes, microvesicles, and friends. J. Cell Biol., 200(4): 373-383. Reinhardt T. A., Lippolis J. D., Nonnecke B. J., Sacco R. E., 2012. Bovine milk exosome proteome. J. Proteomics., 75(5): 1486-1492. Tetta C., Ghigo E., Silengo L., Deregibus M. C., Camussi G., 2013. Extracellular vesicles as an emerging mechanism of cell-to-cell communication. Endocrine., 44(1): 11-19. Trajkovic K., Hsu C., Chiantia S., Rajendran L., Wenzel D., Wieland F., 2008. Ceramide triggers budding of exosome vesicles into multivesicular endosomes. Science., 319(5867): 1244-1247. Turturici G., Tinnirello R., Sconzo G., Geraci F., 2014. Extracellular membrane vesicles as a mechanism of cell-to-cell communication: advantages and disadvantages. Am J Physiol Cell Physiol., 306(7): C621-C633. Vidal M., 2010. Exosomes in erythropoiesis. Transfus Clin. Biol., 17(3): 131-137. Yang L., Andrews D. A., Low P. S., 2000. Lysophosphatidic acid opens a Ca++ channel in human erythrocytes. Blood, 95(7): 2420-2425. Zhou Q., Li M., Wang X., Li Q., Wang T., Zhu Q., 2012. Immune-related microRNAs are abundant in breast milk exosomes. Int. J. Biol. Sci., 8(1): 118-123. Zwaal R. F., Comfurius P., Bevers E. M., 2005. Surface exposure of phosphatidylserine in pathological cells. Cell Mol. Life Sci., 62(9): 971-988. VAI TRÒ CỦA Ca2+ NỘI BÀO TRONG VIỆC HÌNH THÀNH VÀ GIẢI PHÓNG VI THỂ CỦA TẾ BÀO HỒNG CẦU Nguyễn Đức Bách1*, Nguyễn Hữu Đức1, Phạm Kim Đăng1, Lưu Thảo Linh, Lý Thị Bích Thủy2, Ingolf Bernhardt 3 1Học viện Nông nghiệp Việt Nam 2Viện Công nghệ sinh học, Viện Hàn lâm KH & CN Việt Nam 3Đại học Tổng hợp Saarland TÓM TẮT Sự hình thành và giải phóng các cấu trúc màng có kích thước nhỏ (vesicles) đã được phát hiện ở nhiều loại tế bào khác nhau. Dựa vào kích thước người ta chia các cấu trúc này thành 3 dạng khác nhau, dạng có kích thước rất nhỏ từ 40 đến 100 nm có nguồn gốc từ bên trong tế bào được gọi là exosome, dạng có kích thước từ 0,1 đến 1,0 µm tách ra từ màng tế bào được gọi là vi thể hay microvesicle. Cấu trúc có kích thước lớn từ 1,0-4,0 µm được tách ra từ màng của tế bào đang trong giai đoạn chết (apoptosis). Mặc dù cơ chế hình thành và giải phóng các vi thể còn chưa rõ nhưng chúng tham gia quá trình vận chuyển lipid, các yếu tố sinh trưởng, protein, các phân tử tín hiệu, các mRNA và các RNA không mã hóa giữa các tế bào. Ở người, tế bào hồng cầu chiếm số lượng rất lớn được vận chuyển tới tất cả các mô của cơ thể. Một số nghiên cứu gần đây cho thấy có sự hình thành và giải phóng các vi thể từ tế bào hồng cầu tuy nhiên cơ chế của quá trình này chưa sáng tỏ. Trong nghiên cứu này, các điều kiện cho sự hình thành và giải phóng vi thể từ tế bào hồng cầu người đã được khảo sát. Bằng các kỹ thuật đánh dấu quỳnh quang kết hợp với sử dụng kính hiển vi huỳnh quang và máy đếm tế bào, kết quả cho thấy phorbol-12-myristat-13-acetate (PMA), lysophosphatidic acid (LPA) và calcium inonophore (A23187) đã làm tăng hàm lượng Ca2+ nội bào. Việc tăng nồng độ Ca2+ nội bào dẫn đến hình thành và giải phóng các cấu trúc vi thể. Quá trình hình thành vi thể đi kèm với sự vận chuyển của phosphatidylserine ra lớp màng ngoài của tế bào. Động học của quá trình hình thành vi thể được khảo sát bằng cách đánh dấu tế bào với annexin V-FITC. Kết quả nghiên cứu cho thấy khi nồng độ Ca2+ nội bào tăng đã dẫn đến kích thước tế bào bị thu nhỏ, hình thành và giải phóng các cấu trúc vi thể khỏi tế bào hồng cầu. Từ khóa: Canxi nội bào, phosphatidyl serine, tế bào hồng cầu, vi thể. Ngày nhận bài: 22-10-2014