Characterisation of exosomes derived from human

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ARTICLE IN PRESS Colloids and Surfaces B: Biointerfaces xxx (2011) xxx–xxx

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Characterisation of exosomes derived from human cells by nanoparticle tracking analysis and scanning electron microscopy Viktoriya Sokolova a,1 , Anna-Kristin Ludwig b,1 , Sandra Hornung b , Olga Rotan a , Peter A. Horn b , Matthias Epple a,∗ , Bernd Giebel b,∗∗ a Institute of Inorganic Chemistry, Faculty of Chemistry, University of Duisburg-Essen, and Center for Nanointegration Duisburg-Essen (CeNIDE), Universitaetsstr. 5–7, 45117 Essen, Germany b Institute for Transfusion Medicine, University Hospital Essen, Hufelandstr. 55, 45122 Essen, Germany

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Article history: Received 15 February 2011 Received in revised form 2 May 2011 Accepted 5 May 2011 Available online xxx Keywords: Exosomes Vesicles Dynamic light scattering Nanoparticle tracking analysis

a b s t r a c t Exosomes from three different cell types (HEK 293T, ECFC, MSC) were characterised by scanning electron microscopy (SEM), dynamic light scattering (DLS) and nanoparticle tracking analysis (NTA). The diameter was around 110 nm for the three cell types. The stability of exosomes was examined during storage at −20 ◦ C, 4 ◦ C, and 37 ◦ C. The size of the exosomes decreased at 4 ◦ C and 37 ◦ C, indicating a structural change or degradation. Multiple freezing to −20 ◦ C and thawing did not affect the exosome size. Multiple ultracentrifugation also did not change the exosome size. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Initially exosomes were described as vesicles that were released by rat reticulocytes as a consequence of the fusion of multivesicular endosomes with the plasma membrane [1,2]. More than a decade later, exosomes were isolated from B lymphocytes, and it was demonstrated that as antigen-presenting vesicles, these exosomes can induce T-cell responses [3]. Mainly within the last 5 years, the research on exosomes started to burst and it became evident that exosomes are released from an increasing variety of different cells types. In addition to a number of proteins being involved in intercellular cell signalling, they have been found to transport micro as well as messenger RNAs that can be incorporated and translated in target cells [4]. Acting as multi-signalling batteries they can modulate the immune response, and tumour cells were shown to use such mechanisms to escape from the host immune system [5,6]. As they contain cell-specific signatures analyses of exosomes may

∗ Corresponding author at: Institute of Inorganic Chemistry, University of Duisburg-Essen, Universitaetsstr. 5–7, 45117 Essen, Germany. Tel.: +49 201 1832413; fax: +49 201 1832621. ∗∗ Corresponding author at: Institute for Transfusion Medicine, University Hospital Essen, Virchowstr. 179, 45147 Essen, Germany. Tel.: +49 201 7234204; fax: +49 201 7235906. E-mail addresses: [email protected] (M. Epple), [email protected] (B. Giebel). 1 Both authors share first authorship.

be used for diagnostic purposes, e.g. in melanoma [7] or in ovarian cancer diagnostics [8]. Additionally, positive impacts of exosomes on tissue regeneration have been observed [9]. Thus, in addition to being very interesting intercellular signal mediators whose function will be elaborated within the next few years, exosomes appear as promising new tools for the clinical diagnostics and maybe for novel treatment strategies [10,11]. Up to now, exosomes have mainly been studied with biological techniques, which is often very challenging due to their comparatively small size of 50–100 nm [12]. Scanning electron microscopy (SEM) and atomic force microscopy was reported by Sharma et al. [13]. Here, we applied two well-established colloid-chemical techniques [14], i.e. SEM and dynamic light scattering (DLS), and one comparatively new technique, i.e. nanoparticle tracking analysis, to characterise the size of exosomes derived from different human cell types. The exosome size was followed during storage at −20 ◦ C (including multiple thawing), 4 ◦ C, and 37 ◦ C, and also during ultracentrifugation. These are all established techniques to handle exosomes in biology, and it is of prime importance to assess the integrity of exosomes under these conditions. 2. Materials and methods 2.1. Cells and culture conditions Cells of the human embryonic kidney cell line HEK 293T were cultured in Dulbecco’s Modified Eagle Medium (DMEM; PAA

0927-7765/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2011.05.013

Please cite this article in press as: V. Sokolova, et al., Characterisation of exosomes derived from human cells by nanoparticle tracking analysis and scanning electron microscopy, Colloids Surf. B: Biointerfaces (2011), doi:10.1016/j.colsurfb.2011.05.013


ARTICLE IN PRESS V. Sokolova et al. / Colloids and Surfaces B: Biointerfaces xxx (2011) xxx–xxx

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Fig. 1. Light-microscopic images of the exosome-producing cells: HEK 293T (left), human ECFC (center) and MSC (right) (scale bar: 100 ␮m).

Laboratories, Paschingen, Austria) supplemented with 10% fetal calf serum (FCS; Biochrom AG, Berlin, Germany) and 1% penicillin, streptomycin, glutamine (Gibco, Invitrogen GmbH, Frankfurt, Germany). Primary human mesenchymal stem/stroma cells (MSCs) which had initially been raised from umbilical cord tissue were cultivated in alpha-Minimum Essential Medium (alpha-MEM, PromoCell GmbH, Heidelberg, Germany) supplemented with 10% FCS of a selected, pretested batch (PAN-Biotech GmbH, Passau, Germany). Primary endothelial colony forming cells (ECFC) which had initially been raised from umbilical cord blood were cultured in EGM2 Medium (Lonza Cologne GmbH, Cologne, Germany). One day after cell culture, the supernatants were harvested for the isolation of exosomes, the former media were removed, the cells were washed once with phosphate-buffered saline (PBS) and the supernatant of fresh media was added that had been depleted from serum exosomes by ultracentrifugation at 110,000 × g for 2 h. 2.2. Exosome isolation and purification Exosomes were isolated from cell culture supernatants of HEK 293T cells, ECFCs and MSCs. To remove cells, conditioned media were centrifuged for 5 min at 900 × g, and to remove remaining debris, another centrifugation step was performed for 1 h at 10,000 × g. To remove all particles bigger than 200 nm, the supernatants were filtered through 0.2 ␮m pore filters. To concentrate the exosomes, the filtrate was passed through a Vivacell 100 Filter (Sartorius AG, Goettingen, Germany) during 30 min centrifugation at 4 ◦ C and 400 × g. 10 mL of the concentrated supernatant were passed through a Sepharose CL-2B column (1.5 cm × 45 cm; GE Healthcare, Munich, Germany) and 1 mL fractions were eluted. Following the estimation of the protein contents by a Bradford Ultra protein assay (Expedeon Ltd., Harston, UK), the proteincontaining fractions belonging to the main peak were pooled and centrifuged for 2 h at 110,000 × g at 4 ◦ C. The obtained precipitates were resuspended in 150 ␮L PBS and used for further colloidchemical analyses. 2.3. Analytical methods For scanning electron microscopy (ESEM Quanta 400 instrument; FEI), the exosomes were fixed with 3.7% glutaraldehyde (Sigma–Aldrich GmbH, Taufkirchen, Germany) in PBS for 15 min. After washing twice with PBS, the fixed exosomes were dehydrated with an ascending sequence of ethanol (40%, 60%, 80%, 96–98%). After evaporation of ethanol, the samples were left to dry at room temperature for 24 h on a glass substrate, and then analysed by SEM after gold–palladium sputtering. Dynamic light scattering and zeta potential determinations were performed with a Zetasizer nanoseries instrument (Malvern Nano-Zetasizer, = 532 nm laser wavelength). The exosome size data refers to the scattering intensity distribution (z-average). For particle size determination, nanoparticle tracking analysis (NTA) was performed with a NanoSight LM10 instrument equipped with the NTA 2.0 analytical software. All experiments were carried

out at 1:1000 dilution, leading to particle concentrations around 6 × 107 mL−1 . Each experiment was carried out in triplicate. The 50% median value (D50) is given in all cases, and the standard deviation is given for all data. The particle size distribution in a typical experiment was D10 183 nm, D50 258 nm, D70 277 nm, and D90 381 nm. For dynamic light scattering and nanoparticle tracking analysis, the exosomes which were initially dispersed in PBS after the isolation were diluted 1:1000 with pure water. The salt concentrations after dilution were 137 ␮M NaCl, 2.7 ␮M KCl, 8.1 ␮M Na2 HPO4 , and 1.76 ␮M KH2 PO4 at a pH of 7.4. The total ion strength was therefore about 166 ␮M. Multiple centrifugation steps to assess the stability of the exosomes were performed three times for 2 h each and once for 16 h at 110,000 × g at 4 ◦ C. Following each centrifugation step, the precipitates were resuspended in 200 ␮L PBS. For analytical purposes 50 ␮L of each obtained sample were retained. Multiple deep-freezing of the exosomes was accomplished by freezing of the exosome dispersion (10 mL) at −20 ◦ C for 24 h, followed by thawing under gentle shaking at room temperature for about 5 min. For the NTA experiment, 1 mL of the samples was measured for about 15 min, returned into the storage vessel and deep-frozen again. 3. Results and discussion Three different exosome-producing human cell types were used (Fig. 1). We used exosomes isolated from a human embryonic kidney cell line (HEK 293T), from human umbilical cord blood-derived endothelial cells, so-called ECFC [15], and from human umbilical cord derived MSCs. Fig. 2 shows the results of SEM. All exosomes had a spherical shape with a diameter of about 30–50 nm. Agglomeration occurred due to the drying process before the SEM analysis. The size in dynamic light scattering was 212 nm (PDI = 0.626) for HEK exosomes, 226 nm (PDI = 0.481) for ECFC exosomes and 208 nm (PDI = 0.605) for MSC exosomes. Note that the high polydispersity index (PDI) indicates a multimodal particle size distribution. In dynamic light scattering, the exosomes showed a strongly negative zeta potential of −54 ± 6 mV for HEK exosomes, −49 ± 13 mV for ECFC exosomes and −52 ± 4 mV for MSC exosomes due to the negatively charged phospholipid membrane. The size of the exosomes from nanoparticle tracking analysis (Fig. 3) was around 120 nm for HEK exosomes, 110 nm for ECFC exosomes, and 110 nm for MSC exosomes. The difference to the results from SEM (30–50 nm) compared to nanoparticle tracking analysis (around 110 nm) is due to the fact that the latter one monitors the hydrodynamic diameter of the exosomes in solution, and also that larger particles contribute more strongly to the light scattering than the smaller particles which leads to a shift compared to the D50 value from NTA. Even though the human cells which we used in this study are morphologically very heterogeneous (Fig. 1), their exosomes had comparable sizes and shape (Fig. 2). Recently, it has been shown that components of the evolutionary conserved ESCRT complexes deform the membrane of late endosomes in a defined manner to




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Fig. 2. Scanning electron micrographs of fixed and dehydrated exosomes from three different cell types.

form buds of comparable sizes that are cleaved as intraluminal vesicles into the interior of the late endosome [16]. Since upon fusion with the plasma membrane late endosomes release these intraluminal vesicles as exosomes into the environment, our observation that different cell types secrete exosomes of comparable sizes corresponds very well with the current understanding of the exosome biogenesis. Mainly, two different methods have emerged in biology to enrich exosomes. In the first method, exosomes are purified with a differential centrifugation method which by sequentially increasing the centrifugal forces removes cells and larger particles from conditioned cell culture supernatants or from body fluids. Exosomes are commonly precipitated at a final, at least 2 h lasting centrifugation step at ≥100,000 × g [3,17]. In the second method, exosomes can be purified by size exclusion chromatography, similar to the bigger microvesicles. After removing cells and larger particles by a low-g centrifugation step, exosomes can be

pre-concentrated by two filtration steps with 0.2 ␮m pore filters and 100 kDa MWCO. From these concentrates, exosomes can be purified by size exclusion chromatography. To concentrate the eluted exosomes, the obtained fractions are typically centrifuged at ≥100,000 × g like during the final step of centrifugation based methods [18]. First, we investigated whether the high centrifugal forces that are applied to concentrate the exosomes had any impact on the integrity or the size of the purified exosomes. To this end we size-fractioned supernatants of the human HEK cell line 293T on a Sepharose CL-2B column as described above. The eluted exosome fraction was split into two different aliquots and repeatedly centrifuged at 110,000 × g up to four times. Exosome size and integrity were measured after each centrifugation step by nanoparticle tracking analysis (Table 1). We did not find any differences between the size and integrity of the exosomes that had not been ultra-centrifuged and those that were centrifuged at 110,000 × g for up to 22 h. Therefore, we conclude that the high centrifugal forces have no impact on the size and integrity of the exosomes. Accordingly, to concentrate the exosomes we implemented one ultracentrifugation step within our subsequent experiments. Coupled with our increasing knowledge about exosomes as mediators of intercellular communication and their potential use within the clinics, the question arose how stable they are during storage. Thus, we have measured size and integrity of exosomes

Table 1 Colloid-chemical data of HEK 293T exosomes from nanoparticle tracking analysis after centrifugation at 110,000 × g. The size of exosomes is given as average ± standard deviation (n = 3). Centrifugation steps

Fig. 3. Representative image from nanoparticle tracking analysis (NTA) of HEK exosomes.

After preparation 1 time (2 h) 2 times (2 + 2 h) 3 times (2 + 2 + 2 h) 4 times (2 + 2 + 2 + 16 h)

Size (nm) 119 ± 21 110 ± 18 118 ± 17 108 ± 16 109 ± 15




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Table 2 Colloid-chemical data of exosomes from nanoparticle tracking analysis by storage at 4 ◦ C. The size of exosomes is given as average ± standard deviation (n = 3). The results marked with an asterisk (*) are statistically significant in comparison to in comparison to the results at t = 0 h (P < 0.001). Time (h) 0 24 48 72 96 192 360 480 600

Size (nm) (HEK) 116 ± 27 97 ± 22 88 ± 26 76 ± 22 74 ± 15* 63 ± 12* 61 ± 14* 56 ± 12* 55 ± 10*

Size (nm) (ECFC) 113 ± 15 110 ± 28 114 ± 39 81 ± 20* 75 ± 34* 78 ± 26* 71 ± 18* 52 ± 12* 34 ± 13*

Size (nm) (MSC) 107 ± 19 91 ± 33 93 ± 25 81 ± 14* 72 ± 24* 69 ± 18* 64 ± 12* 50 ± 14* 48 ± 11*

Table 3 Colloid-chemical data of exosomes by nanoparticle tracking analysis during storage at 37 ◦ C. The size of exosomes is given as average ± standard deviation (n = 3). The results marked with an asterisk (*) are statistically significant in comparison to the results at t = 0 h (P < 0.001). Time (h)

Size (nm) (HEK)

Size (nm) (ECFC)

Size (nm) (MSC)

0 1 3 5 24 48 72 96 192

136 ± 37 139 ± 33 158 ± 22 119 ± 22 116 ± 32 64 ± 16* 83 ± 21* 67 ± 18* 53 ± 14*

120 ± 18 138 ± 27 103 ± 34 108 ± 37 86 ± 47 60 ± 16* 63 ± 11* 70 ± 19* 47 ± 16*

109 ± 19 101 ± 31 91 ± 19 93 ± 18 58 ± 15* 56 ± 21* 49 ± 13* 50 ± 17* 39 ± 16*

that were stored in liquid solutions for different time periods, either at 37 ◦ C, at 4 ◦ C or frozen at −20 ◦ C (Tables 2 and 3, and Figs. 4 and 5). Nanoparticle tracking analysis showed that the size of the exosomes decreased after an initial period of about 2 days during storage at 37 ◦ C by about 60%. The size of the exosomes did not change during the first 2 days during storage at 4 ◦ C, but we observed a decrease in the size of exosomes from 120 nm down to 80 nm after 3–4 days. After 25 days of storage at 4 ◦ C, the exosome size had decreased to 30–50 nm (Table 2 and Fig. 3). The rate of decrease was significantly slower than at 37 ◦ C, indicating a higher stability at 4 ◦ C. Notably, multiple deep-freezing and thawing did not affect the exosome size, demonstrating that this method is well suited to store exosomes (Table 4).

Fig. 5. Particle size by nanoparticle tracking analysis for HEK exosomes during storage at 37 ◦ C.

Table 4 Colloid-chemical data of exosomes from nanoparticle tracking analysis after freezing (up to 10 times). The size of exosomes is given as average ± standard deviation (n = 3). Number of ultracentrifugation steps 1 2 3 4 5 6 7 8 9 10

Size (nm) (HEK)

104 ± 31 110 ± 25 97 ± 22 102 ± 24 116 ± 29 113 ± 14 100 ± 32 109 ± 32 106 ± 23 94 ± 18

Size (nm) (ECFC)

112 ± 23 100 ± 17 115 ± 30 94 ± 21 103 ± 17 120 ± 44 112 ± 14 92 ± 28 108 ± 44 109 ± 24

Size (nm) (MSC)

109 ± 29 117 ± 45 102 ± 26 112 ± 21 105 ± 27 98 ± 31 115 ± 26 94 ± 43 101 ± 25 107 ± 23

4. Conclusions We have analyzed exosomes in different experimental settings by established colloid-chemical methods. Altogether our analyses qualify nanoparticle tracking analysis and SEM as valuable techniques to analyze in complementation to appropriate biological read-out systems the size and integrity of exosomes in dispersion. In contrast, dynamic light scattering can deliver the exosome charge as represented by the zeta potential, but the exosome size cannot be safely determined due to the polydispersity and probably a weak scattering contrast. We found that the size and integrity of the exosomes was strongly dependent on the storage conditions: the exosome diameter significantly decreased within 2 days at 37 ◦ C and 4 days at 4 ◦ C, but storage at −20 ◦ C did not affect their size. For the typically applied purification procedures, our result indicates that ultracentrifugation does not affect the exosome size it of high importance.

Acknowledgements

Fig. 4. Particle size by nanoparticle tracking analysis for HEK exosomes during storage at 4 ◦ C.

We thank Liska Horsch and Stefan Radtke for providing human ECFCs and MSCs. We are especially thankful to Prof. Theresa Whiteside and her team (University of Pittsburgh) for the training of A.-K.L. in exosome purification techniques. The project was supported by a travel grant to A.-K.L. and an M.D. fellowship to S.H. within the University’s internal IFORES programme.




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