Characterization of Microvesicles Released from Human Red Blood Cells

Under physiological as well as pathological conditions, various cell types release small spherical fragments called membrane vesicles or extracellular vesicles (EVs). So far many different terms for these EVs such as ectosomes, microvesicles (MVs), shedding vesicles, apoptosomes or microparticles have been used in a number of reports [1,2,3,4,5]. Recently, based on their size and origin of formation, EVs have been classified as exosomes, MVs and apoptotic bodies [6,7,8,9,10].

Exosomes are small enclosed membrane vesicles of nearly uniform size from 30 to 100 nm already described by Johnstone during the in vitro culture of sheep reticulocytes [11]. They were also observed in a variety of cultured cells such as lymphocytes, dendritic cells, cytotoxic T cells, mast cells, neurons, oligodendrocytes, Schwann cells, and intestinal epithelial cells [12,13]. In these cells, exosomes originated from the endosomal network that locates within large sacs in cytoplasm. The release of exosomes to extracellular environment is carried out by the fusion of these sacs to the plasma membrane [7,12,14].

Distinct from exosomes, the biogenesis of MVs arises through direct outward budding and fission of the plasma membrane following different kinds of cell activation or during early state of apoptosis [15]. Normally, MVs are larger compared to exosomes with the size ranging from 50 to 1000 nm [15,16,17]. However, there is an overlapping of the size between exosomes and MVs. So far the mechanism of biogenesis is primarily used to distinguish MVs and exosomes [4,6,16]. The formation and release of MVs is the result of dynamic processes of phospholipid redistribution and cytoskeletal protein breakdown [6].

A vast amount of literature dealing with the release mechanism of MVs in RBCs and their properties in response to A23187, ATP depletion, oxidative stress and storage has been reported (e.g. [18,19,20,21,22,23,24]). In addition, an increase of the levels of RBC-derived MVs in the circulating blood of patients with sickle cell anemia (SCA), thalassemia and glucose-6-phosphate-deficiency (G6PD) has been observed [25,26]. Morphological transitions of RBCs stimulated by exogenous compounds, according to the bilayer-couple hypothesis, also result in the formation of MVs [19,22,27]. Furthermore, the reorganization of the cell membrane such as loss of asymmetrical membrane phospholipid distribution leads to membrane blebbing and formation of MVs [28,29,30,31]. Recently, numbers of studies have shown that an increase of the intracellular Ca²⁺ concentration by opening Ca²⁺ channels or activation of protein kinase C (PKC) leads to PS exposure and formation of MVs in many different cell types including human RBCs [29,30,32,33].

In contrast, apoptotic bodies have been characterized as largest EVs with the size varying from 1 to 5 µm. Nucleated cells undergoing apoptosis pass through several stages, beginning with condensation of the nuclear chromatin, followed by membrane blebbing, and finally releasing EVs and apoptotic bodies [6]. However apoptotic cells are also known to release smaller vesicles (MV) in response to apoptosis induction and these MVs are known to stimulate innate immune responses [34]. Although mature human RBCs have no nucleus and organelles, however, they are able to undergo an apoptosis-like process (also called eryptosis [28,29]) with similar characteristics, e.g. membrane blebbing and formation of MVs [28,29].

Although there is a number of reports about the formation of MVs in RBCs, investigations of the kinetics of this process as well the characterization of released MVs are still limited. By using different methods and techniques we investigated the kinetics of formation of MVs. In addition, the MVs released from human RBCs under different conditions were also isolated and characterized. The study may contribute for understanding the role of MVs in normal biological process and diseases.

Fresh human venous blood from healthy donors was obtained from the Institute of Sports and Preventive Medicine (Saarland University, Saarbruecken) and from the Institute of Clinical Haematology and Transfusion Medicine (Saarland University Hospital, Homburg). Blood was withdrawn by venipuncture into citrate-coated tubes or with heparin as anticoagulant, stored at 4°C and used within one day. Freshly whole blood was centrifuged at 2,000 g for 5 min at room temperature and the plasma and buffy coat was removed by aspiration. Subsequently, RBCs were washed 3 times in HEPES buffered physiological solution (HPS) containing (mM): NaCl 145, KCl 7.5, glucose 10, HEPES 10, pH 7.4. Finally, RBCs were re-suspended in HPS and kept at 4°C for experiments.

For experiments, RBCs were used at 0.05% haematocrit in HPS containing additionally 2 mMCaCl₂ and incubated with Ca²⁺ inonophore 4-bromo-A23187 (positive control, further abbreviated as A23187) or LPA or PMA. After 2 h incubation at 37°C cells were washed in HPS. The concentrations of A23187, LPA, and PMA chosen in the present study are in accordance with previous investigations showing an optimal effect [30,35].

The formation and shedding of MVs was investigated by using an inverted fluorescence microscope (Eclipse TE2000-E, Nikon, Tokyo, Japan) under both transmitted bright field illumination and fluorescence modes. The exposure of phosphatidylserine (PS) in the outer layer of the plasma membrane and in MVs can be displayed by annexin V-FITC with excitation and emission wavelength at 488 nm and 512 nm, respectively. RBCs were prepared in HPS in the presence of 2 mM CaCl₂and 5 µl of annexin V-FITC (0.05% haematocrit). The suspension was transferred to a cover slip (previously coated with poly-L-lysine 0.01%). When the cells were settled on the surface of the cover slip, an imaging procedure for the kinetics of MVs formation was recorded immediately after adding A23187 at final concentration of 2 µM or PMA at 6 µM. For experiment with LPA, the substance was added before the addition of Ca²⁺ (to avoid the influence of Ca²⁺ on the solubility of this moiety [36]). Experiments in the presence of A23187, LPA, or PMA were also carried out in the absence of extracellular Ca²⁺ by adding 2 mM EGTA. For controls, ethanol or DMSO was added.

To take into consideration a possible effect of the intracellular glutathione (GSH) level, bright field imaging experiments were performed with RBCs pre-incubated with N-Acetyl-L-Cysteine (NAC) for 1 h and then stimulated with PMA under 4 different conditions: without NAC, treated with NAC (1 mM), PMA, and NAC plus PMA. All these experiments were carried out in HPS with and without Ca²⁺.

Our experimental approach was designed to provide a real-time assessment of the formation of MVs, while maintaining identical imaging parameters under all conditions. This was accomplished by taking images with an electron multiplication CCD camera (CCD97, Photometrics, Tucson, USA) using a 100×1.4 (NA) oil immersion lens with infinity corrected optics. An image was taken every 20 s for 120 min using the imaging software VisiView (Visitron Systems GmbH, Puchheim, Germany). Exposure time was 4000 ms for fluorescence and 100 ms for bright field transmitted light. Experiments were performed at room temperature in a dark room.

RBCs were treated with LPA, PMA or A23187 as stated before for stimulating the formation of EVs. RBCs were suspended in HPS solution at 0.05% haematocrit in the presence of 2 mM CaCl₂. Subsequently, A23187 or LPA or PMA was added at concentrations of 2 µM, 2.5 µM or 6 µM, respectively (LPA was added before the addition of Ca²⁺). After incubation for 2 h at 37°C with occasionally shaking, cell suspensions were subjected to differential centrifugation. First, a centrifugation step at 1,500 g for 10 min was applied followed by centrifugation at 3,000 g for 15 min twice to remove intact cells, cell debris, and apoptotic blebs. The supernatants were collected and further centrifuged at 25,000 g for 1 h at 4°C to harvest large size MVs. Subsequently, the collected supernatants were transferred to new tubes and ultra-centrifuged at 35,000 rpm (corresponding to about 200,000 g) for 2 h at 4°C for the isolation of small size MVs (Beckman Coulter, Swinging Bucket rotor SW40 Ti, k-factor 137).

To obtain all MVs, after removing intact cells, cell debris, and apoptotic blebs, the supernatants were directly subjected to ultra-centrifuge at 200,000 g for 2 h at 4ºC. Pellets obtained after each centrifugation step were re-suspended in Milli-Q water (ultra-pure) for morphological, size and zeta potential measurements.

Samples were analyzed using a FACScalibur Flow Cytometer (Becton Dickinson Biosciences, Franklin Lakes, USA) and BD CellQuest™ Pro Software 5.2.1. All acquisition and analysis were done using log mode. The parameters were set up using standard calibration kit (BD Calibrite™, BD Calibrite 3 Beads, BD Biosciences). Parameters of both forward and side scatter were adjusted to remove instrument noise (dust). The gating process was done by using a combination of sheath fluid (blank), samples of stimulated RBCs, isolated MVs stained with fluorescence dyes annexin V-FITC for PS and Dil as lipophilic tracer.

Stimulated RBCs under conditions mentioned above were fixed with 2% glutaraldehyde at room temperature for 10 min and washed in phosphate buffered saline, pH 7.4 (PBS-TWEEN® Tablets, 1 tablet per litre, Calbiochem - Merck, Darmstadt, Germany) by centrifugation at 5,000 g for 5 min at room temperature to remove glutaraldehyde. The pellets were re-suspended in Milli-Q water and immediately applied on glass slides and air-dried for 1 h. The slides were dipped quickly and gently washed stepwise with ethanol from 50, 70, 90 to 100% for dehydration. For SEM analysis, the prepared slides were sputtered with a gold layer of 15 nm thickness prior to SEM imaging (Sputter coater: Quorum Q150R ES, Quorum Technologies Ltd, East Grinstead, UK) and kept in a closed box at room temperature.

To prepare samples of MVs for SEM analysis, these particles were fixed with 2% glutaraldehyde for 10 min at room temperature. Subsequently the samples were washed in phosphate buffered saline by an ultra-centrifuge step at 200,000 g for 30 min at 4°C. Further steps were performed similar to the preparation of stimulated RBCs described above.

For SEM imaging (EVO HD15, Carl Zeiss Microscopy GmbH, Jena, Germany), several randomly selected frames from each sample were captured for morphological observation and statistical purpose. SEM imaging was carried out using 5 kV acceleration voltage and secondary electron (SE) detector.

AFM equipment (Bioscope IV, Veeco Instruments, Santa Barbara, USA) was used for topographical imaging. Glutaraldehyde-fixed samples of stimulated RBCs, MVs were scanned in air by tapping-mode. The cantilever OMCL-AC160TS series (Olympus) was used with nominal spring constant of 26 N/m, tip radius less than 10 nm and reflective side coated with Al. Data of height, amplitude and phase modes were recorded simultaneously. The images were scanned at the resolution of 256 × 256 pixels, scanning rate in the range from 0.3 to 0.75 Hz. Topographical images were analyzed and displayed using NanoScope Analysis 1.5 software (Brucker Corporation).

A Zetasizer Nano ZS (Malvern, Worcestershire, UK) was used for MVs size and zeta potential measurement. Uniform polystyrene particles of 100, 200 and 400 nm diameter (Bangs Laboratories, Fishers, USA) at 001% in phosphate buffered saline (PBS) were used to verify instrument operation. For size measurement, the MVs samples were diluted in Milli-Q water (attenuator index position in the range from 7 to 9) and analyzed using the standard operation procedure (SOP) as follows: sample refractive index 1.43 (phospholipid liposomes), dispersant refractive index 1,33 (water), system temperature at 25°C and sample equilibration time for 60 s. One ml of each sample was measured in disposable polystyrene (DTS0012, Malvern Instruments) with a path length of 10 mm. Observed populations of particles were characterized by associated Z-average size (nm) and polydispersity index (PdI).

For zeta potential measurement, the disposable capillary cell DTS1070 (Malvern Instruments) was used. Samples were measured in different solutions and buffers: Milli-Q water, NaCl 5 mM or 10 mM, and Tris-HCl 5 mM or 10 mM, pH 7.0. The SOP was set up similar as described above. Each sample was measured 3 times with maximum 100 runs in automatic mode. Observed populations of particles were characterized by associated phase (rad) and zeta potential (mV).

The Malvern Zetasizer software v7.03 was used to collect and analyze the data. The error bars displayed on the DLS graphs were presented by the standard deviation (SD) of three measurements of the same sample.

A23187, LPA, PMA and NAC were purchased from Sigma-Aldrich (St. Louis, USA). Annexin V-FITC and Dil stain (DiIC18(3)) were obtained from Life Technologies (Carlsbad, USA). In this study, A23187 was dissolved in absolute ethanol at 1 mM. LPA and PMA were dissolved in DMSO at 1 mM and stored at -20°C. NAC was prepared as 0.5 M stock solution in water placed in a heating block for 1 h. Glutaraldehyde (for electron microscopy), and all other chemicals were at analytical grade and purchased from Sigma-Aldrich.

Data are presented as the mean values ± S.D. of at least 3 experiments of different blood samples. Statistical analysis was performed using Student's t-test when Gaussian distributed. The values were taken as significant difference when p ≤ 0.05 or p ≤ 0.01. Otherwise, a Mann-Whitney test was performed.

It has been shown that treatment of human RBCs with A23187, LPA, or PMA in the presence of extracellular Ca²⁺ leads to an increase of intracellular Ca²⁺ and exposure of PS in the outer cell membrane leaflet [30,32,37]. In our recent publication we mentioned that after treatment of human RBCs with these substances MVs have been observed [30]. However, this observation was not in the focus of that paper. Therefore, in this paper we investigated the process of membrane blebbing and formation of MVs and characterized these particles in more detail. Figure 1 shows the kinetics of the formation and the outward budding process of MVs in human RBCs after stimulation with A23187, LPA or PMA. The lag time before the formation of MVs started was different among different substances used for stimulation.

Treatment with A23187 resulted in a reduction of cell volume already after 5 min (not shown but can be seen in [30,35]). The formation of MVs can be clearly seen after 15 min (small spots on the surface of RBCs and in the solution). The MVs were formed and subsequently released into the medium. The exposure of PS can be observed also after 15 min. After 2 h almost all cells show PS exposure and MVs on the cell membrane (Fig. 1, upper row).

In case of LPA treatment, after 10 min, there was an alteration in the morphology of RBCs from the echinocyte or discocyte to a spherical shape (not shown). Subsequently, MVs begin to appear on the cell surface. Although the process of PS exposure on the outer leaflet of cell membrane was faster in comparison to the case of A23187 treatment, the number of cells showing a positive annexin V- FITC signal was about 60% of the cells only (Fig. 1, middle row).

In the presence of PMA, RBCs with echinocyte or discocyte shape showed a tendency to change their morphology to stomatocytes after 10 min of treatment. The stomatocyte shape was unchanged over time and coupled with the formation and shedding of MVs (Figs. 2 and 3.1) after 15 min of treatment. The exposure of PS in the outer leaflet of the cell membrane could be observed in about 70% of the cells after 2 h of treatment (Fig. 1, lower row).

Under all experimental conditions, the exposure of PS in the outer cell membrane leaflet and the formation of MVs were observed. However, the fluorescence signal of annexin V-FITC in case of PMA is significant lower in comparison to LPA or A23187 (Fig. 1).

Fluorescence imaging using annexin V-FITC for PS staining (Fig. 1) showed that under all stimulating conditions (A23187, LPA, PMA) only a small but more or less identical amount of MVs could be observed. However under bright-field imaging much more MVs can be seen in case of PMA treatment compared to fluorescence imaging (cp. Figs. 1, 2B) and compared to bright field imaging of cells and MVs after A23187 and LPA treatments (not shown). Furthermore, with bright field imaging, the formation of MVs was also clearly observed when RBCs were treated with PMA in the absence of Ca²⁺ (Fig. 2A).

Treatment of RBCs with NAC (1 mM) did not lead to changes of cell morphology in the presence or absence of extracellular Ca²⁺ (Fig. 3.1 A, B and Fig. 3.2 A, B). In addition, no significant difference in cell morphology between PMA-treated and NAC- plus PMA-treated cells in the presence or absence of extracellular Ca²⁺ could be observed (Fig. 3.1 C, D and Fig. 3.2 C, D). In all cases no significant differences in the formation of MVs were seen. There was also no PS exposure when RBCs were treated with NAC (data not shown).

The formation of MVs was also analyzed by flow cytometry. In the controls (freshly washed RBCs) only a very small amount of MVs was observed (Fig. 4). For stimulated RBCs, the distribution of MVs was gated in R2 regions (Fig. 4). There was an overlapping of MVs with instrument noise but this was not significant. By staining with annexin V-FITC, the fractions of MVs showing PS on the outer surface were clearly separated from the noise. In addition, the fluorescent dye Dil was applied to stain MVs, RBCs and all particles derived from cell membrane. By using Dil, based on its lipophilic properties, we were able to separate RBCs, MVs, and instrument noise due to the high fluorescence signal of Dil bound to phospholipid components (Fig. 5).

All released MVs from stimulated RBCs were analyzed by flow cytometer (Fig. 6). The FSC vs. SSC dot plots displayed the distribution of MVs in the region R2. By staining with annexin V-FITC almost all MVs showed PS on their surfaces (gate R5). In case of LPA stimulation, more than 85% MVs showed positive signal of annexin V-FITC and about 15% of MVs displayed no signal (gate R4). In all experiments, instrument noise was accounted for less than 1% of all detected events.

After removing intact cells, cell debris and apoptotic bodies, all MVs released from stimulated RBCs were collected by centrifugation at 200,000 g for 2 h at 4°C. DLS analysis showed that the size of released particles was very heterologous (PDI = 1.0). A symmetric multimodal distribution of particles in the range from 70 to 1000 nm with the mean value of about 200 nm was observed (Fig. 7A).

The population of larger particles (MVs) obtained by centrifugation at 25,000 g for 1 h (see Materials and methods) has sizes from 150 nm to 300 nm (Fig. 7B). The mean value of the size of MVs from 3 different measurements was 205.8 ± 51.4 nm. After collecting MVs, the population of MVs with smaller size was obtained by a further step of ultracentrifugation at 200,000 g for 2 h. DLS data showed that the size of these small MVs was 125.6 ± 31.4 nm (Fig. 7C). There was an overlapping in the size of two populations in the region from 150 to 200 nm (Fig. 7B and C) Under the experimental conditions, no significant difference was observed in the size distribution of MVs released from RBCs stimulated by LPA, PMA or A23187.

Under experimental conditions, MVs are negatively charged (Fig. 8). There was no significant difference in the charge between the populations of MVs. Data analysis showed that the charge of MVs depend on the concentration of ions and the pH of solutions or buffers (p < 0.01). No significant difference in the charge of MVs released from RBCs stimulated by A23187 and LPA was observed (Fig. 8). However, there was a slight difference in the charge of MVs released from RBCs stimulated by PMA in comparison to those released after LPA or A23187 treatment (p < 0.05) when they were measured in Milli-Q water. Under all stimulating conditions, the lowest negative charge of MVs was determined of approximately -40 mV in Milli-Q water (Fig. 8). In case of phosphate saline buffer, the zeta potential values of MVs were recorded of about -10 mV, however, no distribution curve was observed (data not shown).

The morphology and size of stimulated RBCs, MVs were analyzed by SEM (Fig. 9). In comparison to control (RBCs in PBS), the morphology of stimulated RBCs was altered. The formation of MVs has been observed on the surface of stimulated RBCs. In addition, an adhesion of stimulated RBCs was clearly observed (Fig. 9 A1, A2, B1, B2 and C). Under all stimulating conditions, released MVs showed spherical shape and high polydispersity in their size. Statistical analysis showed that the size of MVs is in the range from 100 to 300 nm with the average value of about 200 nm (Fig. 9 E and F).

The size and morphology of MVs were also analyzed by AFM technique using tapping-mode. Data analysis from height, amplitude and phage modes showed that the size of these particles varied from about 70 to 200 nm (Fig. 10). There was no significant difference in size and morphology of MVs released from stimulated RBCs using PMA, LPA or A23187. In addition, the adhesion of MVs was observed in all samples.

Already 5 - 10 min after stimulation of RBCs with LPA, PMA or A23187 changes of cell volume and cell morphology were observed. In addition, exposure of PS on the outer cell membrane leaflet together with the formation and shedding of MVs were clearly seen on the surface of the cells by staining with annexin V-FITC. There was a relation between the exposure of PS and the formation of MVs. In the population of MVs released from RBCs stimulated by A23187 and LPA, the number of MVs showing positive signal for annexin V-FITC was accounted for about 85%. In case of PMA, the number of MVs with positive signal was only about 60% (see gates R4 and R5 of Fig. 4). By comparison the obtained data from MVs stained with annexin V-FITC and Dil, a lipophilic tracer, it revealed that not all MVs showed PS in their outer surface (Fig. 5).

So far, the mechanism for the exposure of PS and shedding of MVs in human RBCs have been described by sustained elevation of intracellular calcium [30,38,39,40] and other events associated with Ca²⁺ such as the opening of the Gardos channel leading to K⁺ efflux, accompanied by an Cl^- efflux, and subsequently to a water efflux and reduction of cell volume [41]; activation of the scramblase [42,43]; activation of calcium-dependent proteases, including calpain [17,44,45]; cytoskeletal alterations [46,47] and change in erythrocyte morphology from the normal discoid shape to a spherical shape [48]. However, in case of PMA, the formation and shedding of MVs occurs in both Ca²⁺-dependent and -independent ways (Figs. 2 and 3.1). The results suggest that this process is not only affected by Ca²⁺-dependent parameters. Other pathways, e.g., the activation of protein kinase C, could also play a substantial role [30,33]. However, a direct physical effect of PMA due to partitioning in the cell membrane on annexin V-FITC binding and releasing of MVs cannot be ruled out completely.

NAC has been reported as a thiol-oxidizing agent which is able to decrease the level of intracellular GSH in RBCs [49] and finally leading to the formation of MVs in G6PD deficient RBCs [50]. However, in the present study we show that treatment of RBCs from healthy people with NAC in the absence of PMA does not result in any changes of cell morphology and formation of MVs (Figs. 3.1 and 3.2). In the presence of PMA, the morphology of RBCs changed from echinocytes or discocytes to stomatocytes coupled with the release of MVs. However, there was no significant difference between RBCs treated with only PMA and both NAC and PMA (Figs. 3.1 and 3.2). It suggests that the effect of oxidative stress caused by the reduction of GSH level is not sufficient to stimulate the formation of MVs within the time-course of the experiment.

At the moment, it is unclear whether the MVs are formed and shed randomly or only in certain regions of the plasma membrane [3,6]. If this process takes place randomly, in principle, the phospholipid components and membrane proteins in MVs will be comparable with the plasma membrane. Therefore, the regions of plasma membrane of RBCs showing PS on the outer leaflet may be a part of all MVs. However, as mentioned above, there was a subpopulation of MVs showing no PS on their outer layer. This suggests that in the whole population of MVs there exist different types of MVs carrying different components of phospholipids and probably membrane proteins on their surfaces. By differential centrifugation, the population of MVs at different size can be partly separated. However, these particles should be clearly separated based on other criteria rather than merely based on their size. However, so far efficient and specific markers to distinguish populations of MVs released from human RBCs have not been described [6,51].

The negative charge of MVs can be explained based on the negative charged PS on the outer layer of MVs [17,52]. Therefore, the negative charge of MVs depends on the number of PS on their outer layer. In addition, the adhesion of MVs themselves and RBCs carrying MVs should be clarified (Fig. 9). Since MVs have a negative surface charge they will repel each other and spread apart. However, Ca²⁺ could play an intermediate role forming molecular cross-bridges among MVs or stimulated RBCs showing PS on their surfaces. In this case it is not easy to explain the adhesion of stimulated RBC with PMA in the absence of Ca²⁺[35]. In order to explain this adhesion, the involvement of receptors and adhesion proteins should be taken into account [4,8,17,35]. Based on these findings, one could suggest that the exposure of PS in the outer cell membrane leaflet and the release of MVs may be directly involved in the aggregation of RBCs during thrombus formation as well as the RBC clearance by macrophages under normal physiological conditions or at certain diseases [37,53,54].

Treatment of RBCs with LPA, PMA or A23187 in the presence of Ca²⁺ led to the exposure of PS on the outer cell membrane leaflet and the release of MVs. A similar effect was seen in RBCs when protein kinase C was activated. The exposure of PS was observed in almost all MVs. However, there was a subpopulation of these particles, which did not show PS on their outer layers. The size of released MVs varied from 70 to 1000 nm. By differential centrifugation, two populations of MVs were separated with size of 205.8 ± 51.4 nm and 125.6 ± 31.4 nm, respectively. All population of MVs revealed a negative charge of about - 40 mV in Milli-Q water. Adhesion among MVs and simulated RBCs was also observed. These findings suggest that the MVs may be involved in blood clot formation and the RBC clearance by macrophages.

This research is funded by Vietnam National Foundation for Science and Technology Development (NAFOSTED) under grant number 106-YS.06-2013.16 and Researcher Links Travel Grant from British Council. MCW is supported by a grant from CNPq (Brasil).

The authors declare no conflict of interest.