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. Author manuscript; available in PMC: 2021 Dec 1.
Published in final edited form as: Neuromolecular Med. 2020 Aug 20;22(4):503–516. doi: 10.1007/s12017-020-08607-1

Cell-Free Extracellular Vesicles Derived from Human Bone Marrow Endothelial Progenitor Cells as Potential Therapeutics for Microvascular Endothelium Restoration in ALS

Svitlana Garbuzova-Davis 1,2,3,4,*, Alison E Willing 1,2,3, Jared Ehrhart 1, Lianchun Wang 5, Paul R Sanberg 1,2,4,6, Cesario V Borlongan 1,2
PMCID: PMC7677172  NIHMSID: NIHMS1622312  PMID: 32820422

Abstract

Repairing the damaged blood-CNS-barrier in amyotrophic lateral sclerosis (ALS) is necessary to prevent entry of detrimental blood-borne factors contributing to motor neuron dysfunction. Recently, we showed benefits of human bone marrow endothelial progenitor cell (hBM-EPC) transplantation into symptomatic ALS mice on barrier restoration by replacing damaged endothelial cells (ECs). Additionally, transplanted cells may endogenously repair ECs by secreting angiogenic factors as our subsequent in vitro study demonstrated. Based on these study results, hBM-EPCs may secrete extracellular vesicles, which may contain and transfer diverse vesicular biomolecules towards maintenance of EC functionality. The study aimed to characterize extracellular vesicles (EVs) derived from hBM-EPCs as potential cell-free therapeutics for endothelium repair in ALS. EVs were isolated from hBM-EPC media at different culture times and vesicle properties were evaluated. The protective effects of EVs on mouse brain endothelial cell (mBEC) exposed to ALS mouse plasma were investigated. Uptake and blockage of EVs from GFP-transfected hBM-EPCs in ECs were determined in vitro. Results showed that EVs isolated from hBM-EPCs as nanosized vesicles significantly reduced mBEC damage from the pathological environment and these EVs were taken up by cells. Blockage of β1 integrin on EVs prevented internalization of vesicles in mBECs. Together, these results provide evidence for potential of hBM-EPC-derived EVs as novel cell-free therapeutics for repair of endothelium in ALS. Although determining translational potential of hBM-EPC-derived EVs will require evaluation in vivo, this in vitro study represents a step towards an extracellular vesicle-based approach for repair of the damaged microvascular endothelium in ALS.

Keywords: extracellular vesicles, human bone marrow endothelial progenitor cells, mouse brain endothelial cell line, ALS mouse plasma, in vitro

Introduction

During the last decade, extracellular vesicles (EVs) have been systematically investigated as important mediators of intercellular communication regulating a diverse range of biological processes under physiological and pathological conditions (reviewed in (Bakhshandeh et al. 2017; EL Andaloussi et al. 2013; Gho and Lee 2017; Iraci et al. 2016; Mulcahy et al. 2014; Tetta et al. 2013; Yoon et al. 2014)). Although EVs is a generic term for all secreted vesicles in the extracellular space, these vesicles are composed of microvesicles (MVs), exosomes (Exo), and apoptotic bodies (Abs). Despite debates on vesicle nomenclature (Gould and Raposo 2013), EV content can be distinguished by size (MVs: 50–1,000 nm; Exo: 40–120 nm; Abs: 500–2,000 nm), content, and different biogenesis pathways (reviewed in (Colombo et al. 2014; EL Andaloussi et al. 2013)). The EVs have been shown to play important roles in cell-to-cell communications by transferring various proteins, peptides, lipids, mRNA, and microRNA to recipient cells thereby regulating stem cell plasticity (Bakhshandeh et al. 2017; Lee et al. 2012), angiogenesis (Cantaluppi et al. 2012; Deregibus et al. 2007), and immune responses (Bobrie et al. 2011; Chaput and Théry 2011; Robbins and Morelli 2014; Théry et al. 2009).

There has been an increased research focus on nanoparticles as potential therapeutic agents for treatment of various diseases, including neurodegenerative disorders, as discussed (EL Andaloussi et al. 2013; Iraci et al. 2016), mainly due to nanovesicles’ capability of crossing the blood-brain barrier (BBB) and limited immunogenicity (Das et al. 2019). For example, studies showed that EVs derived from human bone marrow mesenchymal stromal cells (MSC) reduced inflammation in a mouse model of LPS-induced lung injury (Morrison et al. 2017) or lung ischemia-reperfusion injury (Stone et al. 2017). Also, MSC-EVs have demonstrated therapeutic potential for kidney repair (Aghajani Nargesi et al. 2017) and tissue regeneration in sepsis (Zheng et al. 2018). Additionally, MSC-derived Exo reduced infarct size in a mouse model of myocardial ischemia/reperfusion injury (Lai et al. 2010) and promoted the healing process in a femur fracture mouse model (Furuta et al. 2016). Notably, MVs derived from human endothelial progenitor cells (EPCs) enhanced endothelial cell survival, proliferation, and formation of capillary-like structures in vitro by horizontal transfer of mRNAs and stimulated angiogenesis in vivo in immunodeficient SCID mice (Deregibus et al. 2007). Moreover, xenotransplantation of human pancreatic islets with EPC-derived MVs into SCID mice led to marked improvement towards neoangiogenesis within engrafted tissue and increased insulin secretion (Cantaluppi et al. 2012). These study results demonstrated promising extracellular vesicle approaches for protective and/or regenerative therapeutics strategy. Although functional mechanism(s) are still unclear, EVs may contain diverse vesicular cargo proteins from parent cells, recapitulating many of the beneficial effects of whole-cell therapy.

Our current research primarily aims to develop cell therapy for repair of the damaged blood-CNS-barrier (B-CNS-B) in amyotrophic lateral sclerosis (ALS), a fatal neurodegenerative disease (Eve et al. 2018; Garbuzova-Davis et al. 2017; Garbuzova-Davis, Haller, et al. 2018). Recently, we (Garbuzova-Davis, Kurien, et al. 2019) demonstrated the beneficial effects of intravenously transplanted human bone marrow endothelial progenitor cells (hBM-EPCs) into symptomatic G93A SOD1 mutant mice on restoration of the B-CNS-B via potential replacement of damaged endothelial cells (ECs) with administered cells. Widespread engraftment of transplanted hBM-EPCs into the capillary lumen of the gray/white matter spinal cord and brain motor cortex/brainstem, provided significant restoration of capillary ultrastructure, substantial decrease of capillary permeability, and maintenance of perivascular astrocyte end-feet. The superior barrier repair led to significantly increased spinal cord motor neuron survival and improvement of disease outcomes in cell-treated ALS mice.

Despite robust therapeutic efficacy, the exact mechanisms of transplanted hBM-EPC actions are still uncertain. Our next study was designed to elucidate the constituents of these angiocompetent cells, factors which likely contributed to the repair mechanism. The examination of hBM-EPCs in vitro at different time points under normogenic conditions predominantly revealed gradual significant increases of VEGF-A and angiogenin-1 in conditioned media, re-arrangement of cytoskeletal F-actin filaments, and cellular zonula occludens 1 and occludin immunoexpressions (Garbuzova-Davis, Ehrhart, et al. 2019). These study results provide evidence for potential endogenous endothelium repair in ALS, in addition to replacement of deteriorated ECs, through hBM-EPC transplantation via angiogenic factor secretions and tight junction protein expressions. Our findings support those of a previous study (Urbich et al. 2005) showing that EPCs derived from human peripheral blood highly expressed angiogenic growth factors and other active molecules enhancing formation of new vessels and tissue regeneration. Since hBM-EPCs may have a dual effect (i.e. EC replacement and endogenous EC enhancement) on B-CNS-B restoration in ALS, additive actions of transplanted cells should be considered. We hypothesized that hBM-EPCs secrete extracellular vesicles, which may contain and transfer diverse vesicular molecules towards maintenance of EC functionality.

The aim of this study was to characterize extracellular vesicles (EVs) derived from human bone marrow endothelial progenitor cells (hBM-EPCs) as potential cell-free therapeutics for endothelium repair in ALS. First, EVs isolated from media of cultured hBM-EPCs at different time points were evaluated regarding extracellular vesicle properties. Second, the effects of hBM-EPC-derived EVs on endothelial cells under a pathological environmental condition reminiscent of ALS were examined in vitro. A specific focus was determining the uptake and inhibition of EV incorporation in damaged endothelial cells as potential therapeutic EV mechanisms.

Methods

Ethics

Human bone-marrow-derived endothelial stem cells were purchased from CELPROGEN (Torrance, CA, USA) and a document is available online regarding informed consent of donors: https://www.celprogen.com/uploads/product/14200480134.pdf. No ethical approval was required by the University of South Florida for the use of these cells.

Blood was collected from G93A SOD1 mutant male mice and control non-carrier mutant SOD1 gene male mice used in our previous published studies (Garbuzova-Davis et al. 2017; Garbuzova-Davis, Kurien, et al. 2019). All described procedures for animals used in these studies were approved by the Institutional Animal Care and Use Committee at the University of South Florida and conducted in compliance with the Guide for the Care and Use of Laboratory Animals. All applicable international, national, and/or institutional guidelines for the care and use of animals were followed.

Cell Preparation and Culture Procedure

Human Bone Marrow Endothelial Progenitor Cells

Cryopreserved human bone marrow-derived endothelial progenitor cells (hBM-EPCs, CELPROGEN, Torrance, CA, USA) were thawed rapidly using a water bath at 37°C and then transferred slowly with a pipette into a 15-mL centrifuge tube containing 10 mL of phosphate buffered saline 1X (PBS), pH 7.2 (Cat. No. SH30256.01, HyClone Laboratories, Logan, UT, USA). The cells were centrifuged (100 g/10 min) at room temperature (RT), the supernatant was discarded, and the process was repeated. After the final wash, cell viability was assessed before culture using the Vi-Cell™ cell viability analyzer (Beckman Coulter, Miami, FL, USA). The hBM-EPCs were cultured in separate 24-well plates (2 X 104 cells/500 μL commercial basal media/well) for different experiments.

In experiment 1, hBM-EPCs were cultured for 24 hrs, 72 hrs, and 5 days in vitro (DIV) and conditioned media was collected at each time point for extracellular vesicle (EV) isolation and analyses as described below. Media was renewed after collection at 24 and 72 hrs. Cells were examined under an inverted Olympus IX71 microscope and randomly selected phase-contrast images (n=5/time point) were taken for morphological analysis.

In experiment 2, hBM-EPCs were cultured in basal media for 24 hrs after cell seeding. The next day, media was changed with supplementation of either 1%, 3%, or 5% ALS plasma derived from early symptomatic G93A SOD1 mutant mice in duplicate for each condition. After 48 hrs of incubation, cell viability was determined using the LIVE/DEAD assay as described below.

In experiment 3, hBM-EPCs were initially cultured for 24 hrs in basal media and then transfected with pMAX GFP using lipofectamine for 48 hrs as described below. The media was then renewed and collected after 48 hrs of cell incubation for isolation of GFP labeled EVs.

Mouse Brain Endothelial Cell Line

Cryopreserved mouse brain endothelial cells from line bEnd.3 [BEND3] (ATCC® CRL-2299™) (mBECs, ATCC, Manassas, VA, USA) were thawed rapidly using a water bath at 37°C and then transferred slowly with a pipette into a centrifuge tube containing 10 mL of PBS 1X, pH 7.2. The cells were centrifuged (130 g/7 min) at RT, the supernatant discarded, and the process repeated. After the final wash, cell viability was assessed before culturing as described above. The mBECs were cultured in ATCC-formulated Dulbecco’s Modified Eagle’s Medium (Cat. No. 30–2002) containing 10% fetal bovine serum (Corning®, Cat# 35–010-CV, Lot# 35010161, Corning, NY, USA).

For the initial study, mBECs were cultured in 24-well plate (2 X 104 cells/500 μL media/well) for 48 hrs, reaching 80–85% cell confluence. Media was then changed by adding 3% ALS plasma obtained from early symptomatic mice for an additional 48 hrs incubation. The mBECs were then incubated in media supplemented with 3% ALS mouse plasma and EVs isolated from 72 hrs cultured hBMEPCs at a concentration of 1 μg/mL or 5 μg/mL in duplicate. The next day, the LIVE/DEAD viability/cytotoxicity assay for cell viability was performed as described below.

In a follow-up study, mBECs were cultured in 8-well chamber slides (5 X 103 cells/250 μL media/well, Lab-TekR, Naperville, IL) and the in vitro procedure was conducted as described above. In these experiments, hBM-EPC-derived EVs from 72 hrs cultures were used at 1 μg/mL in duplicate. Cultured mBECs with only 3% ALS mouse plasma, 3% plasma from control mice, or basal media were used as controls. The study was performed in duplicate.

Additionally, mBECs were cultured in 8-well chamber slides as described above. After 48 hrs of exposing cells to 3% ALS mouse plasma, cells were incubated for 24 hrs in media supplemented with 3% ALS mouse plasma and 1 μg/mL of GFP EVs isolated from transfected hBM-EPCs. Cultured mBECs incubated in only basal media with same concentration of GFP EVs were used as a control. The next day, cells were fixed by 4% paraformaldehyde (PFA) in PBS solution and Vectashield® containing DAPI (Vector Laboratories, USA) was applied for imaging analysis of GFP EV distribution within cells. The study was performed in duplicate.

Transfection of hBM-EPCs

The hBM-EPCs were initially seeded for 24 hrs (n=4 well) and then transfected with pMAX GFP (0.5 μg/well, Cat. No. V4YP-2A24, Lonza, Basel, Switzerland) using lipofectamine™ 2000 (1.25 μL/well, Cat. No. 11668019, Thermo Fisher, USA) for 48 hrs accordingly to the manufacturer’s protocol. Media was then replaced, and cells were grown for an additional 48 hrs before media was collected for isolation of GFP labeled EVs. Cells were then fixed in 4% PFA solution and DAPI added into the well. The transfected hBM-EPCs were examined under an epifluorescent inverted Olympus IX71 microscope and images (n=8) were randomly taken for further analysis. The efficiency of GFP labeled hBM-EPCs was analyzed using NIH ImageJ software (version 1.46) and presented as the percentage of GFP positive cells from the total number of cells counted in the entire image.

Extracellular Vesicle Isolation

Conditioned media collected from cultured hBM-EPCs at 24 hrs, 72 hrs, and 5 DIV or transfected and non-transfected cells was processed for EV isolation from two separate experiments using the Total Exosome Isolation kit (Cat. No. 4478359, Invitrogen, USA) according to the manufacturer’s instructions. Briefly, media (1 mL) was centrifuged at 2000 g for 30 min at RT to remove cells and debris. The supernatant was transferred to a new tube without disturbing the pellet. Then reagent, at half the volume of the supernatant, was added and stored at 4° C overnight. The next day, samples were centrifuged at 10,000 g for 60 min at 4° C. After removal of the supernatant, the pellet was re-suspended in 25 μL of PBS and stored at −20° C for further analysis.

Extracellular Vesicle Protein Concentration and Size

Total protein concentrations in collected EVs were determined using the NanoDrop™ OneC (Thermo Scientific, USA) at absorbance of A260/A280. Briefly, EV samples were thawed at RT and 1 μL of each was used in duplicate. Protein concentration is presented as mg/mL.

EV sizes were analyzed in collected samples from cultured hBM-EPCs at 24 hrs, 72 hrs, and 5 DIV using the NanoSight LM10 (Malvern Instruments, UK) with Nanosight NTA 3.1 software. Briefly, the samples were diluted with filtered (0.02 μm filter) distilled water to 25 ng/mL. The sample (350 μL) was then loaded into the instrument analysis chamber with a syringe. Using the Nanosight Tracking Analysis software (version 3.1, Malvern Instruments, UK), the camera was turned on and the screen gain and camera level were set to 4.5 and 11, respectively, to sharply focus the image of the particles. The measurement parameters were set to capture 3 separate measurements of 60 sec duration. After the videos were captured, the program automatically processed the data, measuring particle size and concentration of particles per milliliter. For examination of GFP-labeled EVs, two μL of obtained EVs were smeared on slides and cover-slipped with distilled water. The images were taken under epifluorescence using an Olympus BX60 microscope at high magnification and analyzed for EV size and GFP expression.

Blocking Incorporation of EVs in mBECs

Preventing EV internalization in mBECs via blockage of adhesion molecule was performed as described (Deregibus et al. 2007). Briefly, mBECs were cultured in 8-well chamber slides as described above. After 48 hrs of exposing cells to 3% ALS mouse plasma, cells were incubated for 24 hrs in media supplemented with 3% ALS mouse plasma, 1 μg/mL of hBM-EPC-derived EVs or GFP labeled EVs from transfected hBM-EPCs, and 1 μg/mL of purified mouse anti-CD29 (integrin β1) antibody (Cat. No. 610467, Clone 18/CD29 [RUO], Becton Dickinson Bioscience, Sparks, MD, USA). The hBM-EPC-derived EVs or GFP EVs were pre-incubated with mouse anti-CD29 antibody for 30 min at 37° C. Cultured mBECs incubated in only basal media with same concentration of GFP EVs and CD29 were used as a control. The next day, the LIVE/DEAD viability/cytotoxicity assay for cell viability was performed in duplicate in cultures containing 3% ALS mouse plasma + hBM-EPC-derived EVs + CD29 as described below. In cultures with 3% ALS mouse plasma + hBM-EPC-derived GFP EVs + CD29, cells were fixed by 4% PFA in PBS solution and Vectashield containing DAPI (Vector Laboratories, USA) was applied for imaging analysis of GFP-derived EV incorporation in cells. The study was performed in duplicate.

Cell Viability Assay

Viability of cultured hBM-EPCs and mBECs under exposure of ALS mouse plasma at different concentrations was determined using the LIVE/DEAD viability/cytotoxicity kit (Cat. No. L3224, Fisher Scientific, Pittsburg, PA, USA) as we previously described (Ehrhart et al. 2018). Briefly, the culture media was removed and cells were washed with PBS twice in each well. The combined LIVE/DEAD assay reagents were added to each well (24-well plate - 375 μL; 8-well plate - 250 μL) and incubated at RT for 30 minutes. After incubation, randomly selected images (n=5–8/well) were obtained at 20X magnification for cell quantification using the epifluorescent inverted Olympus IX71 microscope. Live cells were labeled with green fluorescence through the conversion of non-fluorescent cell-permanent calcein acetoxymethyl to intensely fluorescent calcein by ubiquitous intracellular esterase enzyme activity. Dead cells were identified using ethidium homo dimer-1 (EthD-1), which enters cells through damaged membranes and produces a red fluorescence upon binding to nucleic acids. Cell counts of live (green) and dead (red) cells were determined using NIH ImageJ software (version 1.46).

Obtaining Plasma from Mouse Blood

Blood was obtained via submandibular bleeding of G93A SOD1 mutant male mice at 12–13 weeks of age at early symptomatic stage (n=7) and control non-carrier mutant SOD1 gene male mice (n=3) of the same age. These animals were used in our previous published studies (Garbuzova-Davis et al. 2017; Garbuzova-Davis, Kurien, et al. 2019). Briefly, mouse blood was collected in BD Microtainer blood collection tubes containing 1.0 mg K2EDTA anticoagulant (Becton, Dickinson and Company, Franklin Lakes, NJ, USA) and tubes remained at RT for 10 min. Samples were then centrifuged at 3000 rpm for 15 min at RT to separate blood cells and plasma. The plasma was then transferred to a new tube, aliquoted, and stored at −80° C.

Statistical Analysis

Data are presented as means ± S.E.M. One-way ANOVA with post-hoc Tukey HSD (Honesty Significant Difference) multiple comparison test using online statistical software (astatsa.com, 2016 Navendu Vasavada) was performed for statistical analysis. Significance was defined as p < 0.05.

Results

Characteristics of hBM-EPC-derived EVs

The EVs were isolated from conditioned media of cultured hBM-EPCs at different time points: 24 hrs, 72 hrs, and 5 DIV. Morphology of cultured hBM-EPCs changed over time from rounded to elongated cells. Predominantly, rounded cells were detected at 24 hrs after initial seeding (Fig. 1A). Elongated cells with long processes were more apparent at 72 hrs (Fig. 1B) and 5 DIV (Fig. 1C). Only a few rounded cells were observed in these cultures. Notably, some swollen cells with enlarged nuclei were identified in prolonged 5 DIV cell cultures (Fig. 1C).

Fig. 1.

Fig. 1

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Morphology of hBM-EPCs in vitro and total protein concentration in isolated EVs. Morphologies of cultured hBM-EPCs changed from rounded at 24 hrs (A) to elongated at 72 hrs (B) and 5 DIV (C). Decreased appearance of rounded cells was detected in prolonged cell cultures. Of note, swollen cells with enlarged nuclei were observed at 5 DIV (C, center of image). * -rounded cells, < - elongated cells. Scale bar in A-C is 50 μm. (D) Total protein concentration was similar between each time points

Measurements of total protein concentrations in hBM-EPC-derived EVs showed similar amounts at all examined time points: 24 hrs – 20.11 ± 1.85 mg/mL, 72 hrs - 20.62 ± 0.28 mg/mL, and 5 DIV - 19.94 ± 0.88 mg/mL (Fig. 1D). However, there was a diverse size range of EVs collected at different culture time points: 24 hrs – 93–267 nm, 72 hrs – 47–371 nm, and 5 DIV – 28–553 nm (Fig. 2A, 2B, 2C). Although EV sizes tended to increase over culture time (24 hrs – 186.4 ± 21.3 nm, 72 hrs - 242.8 ± 11.4 nm, and 5 DIV - 276.6 ± 63.9 nm), no significant differences were determined between time points (Fig. 2D).

Fig. 2.

Fig. 2

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Sizes of hBM-EPC-derived EVs isolated at different culture time points. A diverse range of EV sizes was determined at 24 hrs (A), 72 hrs (B), and 5 DIV (C). Peaks with numbers refer to concentrations of nanoparticles/mL. (D) EV sizes increased over time in cultures without significant differences between time points

Together, results showed adequate isolation of hBM-EPC-derived EVs from conditioned media at different culture time points based on total protein concentrations and EV sizes. According to range of EV sizes, EVs are referred to as nanosized vesicles. Although EVs isolated from 24 hrs demonstrated reasonable sizes, cultured cells at 72 hrs displayed active morphogenesis vs. 24 hrs and EV sizes were within a tighter range vs. 5 DIV.

Analysis of ALS Mouse Plasma Effect on hBM-EPCs Viability In Vitro

To mimic in vivo environments of intravenous transplantation of hBM-EPCs into ALS mice, cells were exposed in vitro to different concentrations of ALS plasma obtained from early symptomatic G93A SOD1 mutant mice. The hBM-EPC viability was determined by using the LIVE/DEAD Viability/Cytotoxicity assay. Numerous viable cells were observed in cultures with basal media and 1% ALS mouse plasma (Fig. 3A, 3B). There were no significant differences (p > 0.05) of dead hBM-EPCs between their incubation in basal media (0.39 ± 0.28%) and 1% ALS mouse plasma (0.69 ± 0.35%) (Fig. 3E). A significant increase (p < 0.01) of dead cells was found after exposure to 3% (6.06 ± 0.11%) or 5% (8.57 ± 0.63%) vs. 1% ALS mouse plasma or basal media (Fig. 3C, 3D, 3E). Moreover, a significantly (p < 0.05) higher percentage of hBM-EPC death was noted after adding 5% ALS mouse plasma compared to 3% (Fig. 3E). Additionally, culture media supplementation with either 3% or 5% ALS mouse plasma changed live cell morphology, as demonstrated by substantial numbers of swollen cells (Fig. 3C, 3D). Under these supplements, some dead cells showed evidence of ruptured membranes.

Fig. 3.

Fig. 3

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Viability of hBM-EPCs in vitro after exposure to different concentration of ALS mouse plasma. The cells were stained using the LIVE/DEAD Viability/Cytotoxicity assay to identify the viable (green fluorescence) and non-viable cytotoxic (red fluorescence) cell populations. Numerous viable (green) hBM-EPCs cultured in basal media (A) and 1% ALS mouse plasma (B) were observed. Increased numbers of dead (red) cells were detected in cultures of media supplemented with 3% (C) and 5% (D) ALS mouse plasma. Of note, a substantial number of swollen live hBM-EPCs was evident in cultures supplemented with these concentrations of ALS mouse plasma. Scale bar A-D is 50 μm. (E) A significant increase of dead hBM-EPCs was determined in cultures where media was supplemented with 3% or 5% ALS mouse plasma vs. 1% or basal media. Also, a significantly greater percentage of cell death was noted after adding 5% ALS mouse plasma compared to 3%. * - p < 0.05, ** - p < 0.01

Thus, results demonstrated that exposure of hBM-EPCs to plasma derived from symptomatic ALS mice affected cell viability in vitro. As the percentage of supplemental ALS plasma increased in the culture media, moderate cell death also increased; 3% ALS mouse plasma supplement was selected as a reasonable “inducer” for cell damage.

Analysis of hBM-EPC-derived EVs on mBEC Viability under Pathological Condition In Vitro

To evaluate the effect of hBM-EPC-derived EVs upon EC damage induced by ALS mouse plasma, cells from mouse brain endothelial cell line bEnd.3 [BEND3] (mBEC) were used to mimic the in vivo situation. Initially, dose-response effects of EVs were characterized on mBEC viability (LIVE/DEAD Viability/Cytotoxicity assay) under 3% ALS mouse plasma supplementation. Results showed a significant increase (p < 0.01) in dead cells by ALS mouse plasma exposure (12.78 ± 0.77%) vs. cells grown in basal media (6.41 ± 1.75%). A substantial decrease (p < 0.01) of dead mBECs (4.66 ± 0.23%) was found after adding 1 μg/mL of EVs. When culture media was supplemented with 5 μg/mL of EVs, numerous dead cells (14.21 ± 1.71%) were detected similarly to cells exposed to ALS mouse plasma alone. These initial results indicated that 1 μg/mL of EVs was sufficient to alleviate mBEC damage. Concentration of EVs at 5 μg/mL was potentially toxic for cells as indicated by increased cell death.

Based on the EV dose-response analysis, the next experiment was performed to confirm efficacy of the best EV concentration. The mBECs were cultured in basal media, 3% control mouse plasma, or 3% ALS mouse plasma for 48 hrs. Then 3% ALS mouse plasma was supplemented with 1 μg/mL of EVs for an additional 24 hrs. Similar to initial study results, ALS mouse plasma induced a significant (p < 0.01) increase in dead mBECs (26.19 ± 4.11%) in comparison to control mouse plasma (11.92 ± 2.19%) or basal media (5.69 ± 0.81%) (Fig. 4A, 4B, 4C, 4F). Significant (p < 0.01) reduction in cell death (12.73 ± 2.83%) to the level of control mouse plasma was determined after adding 1 μg/mL of EVs (Fig. 4D, 4F). Numerous viable mBECs with improved cell morphology were observed in these cultures. Only a few dead cells were noted. When EVs were pre-treated with anti-CD29 blocking antibody, significant (p < 0.01) increase in percentage of dead mBECs (22.06 ± 3.22%) vs. EVs treatment alone at the same concentration was noted (Fig. 4E, 4F). Cells cultured with CD29 antibody alone demonstrated results similar to those of basal media cultures (data not shown).

Fig. 4.

Fig. 4

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Effect of hBM-EPC-derived EVs on mouse brain endothelial cell viability after ALS mouse plasma exposure in vitro. The cells were stained using the LIVE/DEAD Viability/Cytotoxicity assay to identify the viable (green fluorescence) and non-viable (red fluorescence) cell populations. Numerous viable (green) mBECs cultured in basal media (A) and control mouse plasma (B) were observed. (C) Dead (red) cells were significantly increased by exposure to 3% ALS mouse plasma for 48 hrs. (D) Adding 1 μg/mL of EVs to culture media supplemented with ALS plasma for 24 hrs, promoted cell survival as shown by a substantial decrease in the number of dead cells. (E) Pre-incubation of EVs with anti-CD29 blocking antibody significantly increased dead cells. Scale bar in A-E is 50 μm. (F) A significant increase of dead mBECs was determined when culture media was supplemented with 3% ALS mouse plasma vs. basal media or control mouse plasma. A significant reduction in cell death to level of control mouse plasma was found after adding 1 μg/mL of EVs. Pre-treatment of EVs with CD29 antibody significantly increase cell death in comparison to EVs treatment effect at the same concentration. ** p < 0.01

Together, results showed the beneficial effect of hBM-EPC-derived EVs at 1 μg/mL concentration on mBECs damaged by ALS mouse plasma. Cell morphology was also enhanced after only a short time of EV treatment. Importantly, pre-incubation of EVs with blocking antibody against β1 integrin (CD29) showed no treatment EV effects, presumably by inhibition of EV incorporation in damaged mBECs.

hBM-EPCs Transfection and the Uptake of GFP-labeled EVs by mBECs In Vitro

To elucidate the potential mechanism underlying hBM-EPC-derived EV actions leading to protection of endothelial cells from damage via pathological environment, hBM-EPCs were transfected with pMAX GFP using lipofectamine. Non-transfected cells were cultured in basal media and served as controls. Media was collected from transfected and non-transfected hBM-EPCs for isolation of GFP-labeled and non-GFP labeled EVs. GFP-labeled hBM-EPCs were well apparent in cultures with appropriate cell morphology (Fig. 5A) analogous to non-transfected cells (Fig. 5B). Although, the efficiency of GFP-labeled hBM-EPCs upon culture fixation was 37.64 ± 1.59%, GFP-labeling efficiency on live cells was much higher (~70%, data not shown). GFP-labeled EVs were visible at appropriate vesicle sizes (Fig. 5C). Total protein concentrations in GFP-labeled EVs (20.28 ± 1.27 mg/mL) and non-GFP-labeled EVs (20.05 ± 1.34 mg/mL) were similar (Fig. 5D).

Fig. 5.

Fig. 5

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Transfection of hBM-EPCs with GFP. (A) hBM-EPCs transfected with GFP using lipofectamine showed GFP-labeling (green). Cell nuclei were counterstained with DAPI. Although, the efficiency of GFP-labeled hBM-EPCs upon culture fixation was 37.64 ± 1.59%, GFP-labeling efficiency on live cells was much higher (~70%, data not shown). (B) Non-transfected cultured hBM-EPCs, grown in basal media, were used as controls. (C) Microscopic detection of GFP-labeled EVs demonstrated appropriate vesicle sizes (green). Scale bar in A and B is 50 μm; in C is 20 μm. (D) Total protein concentrations were similar between GFP-labeled EVs and non-GFP-labeled EVs

In the next study, the capability of mBECs to uptake and the potential incorporation inhibition of 1 μg/mL of GFP-labeled EVs under pathological (3% ALS mouse plasma) or normogenic (basal media) condition in vitro was investigated. Numerous GFP-labeled EVs were observed within cytosol of many cultured cells with ALS mouse plasma supplements after 24 hrs incubation (Fig. 6A, 6A’). Moreover, EVs were detected in cell processes. Although GFP-labeled EVs were also seen within some cultured mBECs in basal media, numerous EVs were observed in outer or in inner cell membranes (Fig. 6B, 6B’). Pre-incubation of GFP EVs with anti-CD29 blocking antibody demonstrated prevention of EV incorporation in mBECs. The GFP-labeled EVs were primarily noticed on cell surface after exposure to 3% ALS mouse plasma (Fig. 6C, 6C’). Similar localization of a few GFP EVs was identified in cell cultures with basal media (Fig. 6D, 6D’).

Fig. 6.

Fig. 6

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The uptake of GFP-labeled EVs by mouse brain endothelial cells in vitro. (A, A’) Cytosolic GFP-labeled EV uptake (green, arrowheads) was detected in many mBECs exposed to 3% ALS mouse plasma. The EVs were also detected in cell processes. (B, B’) mBECs cultured in basal media showed GFP-labeled EVs in extracellular space or in inner cell membranes (green, arrowheads). (C, C’) Blocking GFP-labeled EVs with anti-CD29 antibody showed inhibition of EV incorporation in mBECs. These EVs (arrowheads) were mainly detected on cell surface after exposure to 3% ALS mouse plasma. (D, D’) Similarly, GFP EVs were localized in cell cultures with basal media. Only a few EVs (arrowheads) were observed on near cells under normogenic culture condition. Cells in A-D were counterstained with DAPI and cell nuclei displayed in A’-D’, respectfully. Scale bar in A-D’ is 20 μm

Thus, results demonstrated uptake and wide distribution of GFP-labeled EVs within mBECs, largely in cell cultures exposed to ALS mouse plasma. The inhibition of GFP EV internalization in damaged cells was determined by blockage of β1 integrin with specific CD29 antibody.

Discussion

In the present study, human bone marrow endothelial progenitor cell-derived extracellular vesicles (hBM-EPC-derived EVs) as potential cell-free therapeutics for endothelium repair in ALS were characterized under normogenic and pathological environmental conditions in vitro. The major study findings were that hBM-EPC-derived EVs were: (1) classified as nanosized vesicles; (2) shown to significantly reduce mouse brain endothelial cell (mBEC) damage from pathological environment at a concentration of 1 μg/mL; (3) largely taken up by mBECs upon exposure to a pathological environment; and (4) inhibited incorporation in damaged mBECs via blockage of adhesion molecule β1 integrin. These findings provide evidence for potential novel cell-free therapeutics of hBM-EPC-derived EVs for repair of endothelium in ALS by reducing cell damage induced by pathological environment via extracellular vesicle uptake and probable release of various bioactive molecules into cells. Furthermore, results from this in vitro study show promise for an extracellular vesicle-based approach to restore the B-CNS-B in ALS.

Based on beneficial cell transplant outcomes, numerous studies have used MSCs as the cellular source for deriving nanosized vesicles towards therapeutic intervention (Aghajani Nargesi et al. 2017; Dabrowska, Andrzejewska, Lukomska, et al. 2019; Dabrowska, Andrzejewska, Strzemecki, et al. 2019; Furuta et al. 2016; Lai et al. 2010; Qiu et al. 2018; Shojaati et al. 2019; Stone et al. 2017; Zheng et al. 2018). Also, vesicles obtained from human EPCs have shown positive effects in vivo by promoting angiogenesis (Cantaluppi et al. 2012; Deregibus et al. 2007). The authors noted that EPC-derived microvesicles are capable of being internalized into target ECs and thus transfer cargo biomolecules leading to stimulation of angiogenesis and/or enhancement of neoangiogenesis. The vesicles potentially “mimic the effect of the cells from which they are released” (Cantaluppi et al. 2012), acting as mediators of intercellular communication networks (Camussi et al. 2010; Tetta et al. 2013), and may be referred to as “nanosized extracellular organelles” (Gho and Lee 2017).

In our present study, EVs derived from cultured hBM-EPCs at different time points were characterized by total protein concentration and size. Although specific protein content in EVs was not analyzed, total protein concentrations in EVs were consistent across measured time points. According to the range of EV sizes, isolated EVs, comprising a heterogenous group of vesicles, mainly contained both exosomes and microvesicles as discussed in detail (EL Andaloussi et al. 2013). Some apoptotic bodies, likely part of the EVs isolated from 5 DIV cultures, were larger than those obtained at 24 and 72 hrs (more than 500 nm; see Figure 2C). These apoptotic bodies were possibly released from cultured hBM-EPCs, perhaps demonstrating an apoptotic process by changes in cellular nuclei and cytosol shapes. However, this assumption requires confirmation by specific detection of apoptotic cells in vitro, an investigation which is currently underway. Similar to our recent report (Garbuzova-Davis, Ehrhart, et al. 2019), morphology of cultured hBM-EPCs changed over time from rounded (24 hrs) to elongated cells (72 hrs and 5 DIV) suggesting active cell morphogenesis, especially, at 72 hrs with a tighter range of EV sizes vs. 5 DIV.

To mimic a pathologic in vivo environment, in which hBM-EPCs are introduced via intravenous transplantation into early symptomatic G93A SOD1 mutant mice, hBM-EPCs were exposed in vitro to different concentrations of plasma obtained from these symptomatic ALS mice. Results showed a moderate increase in cell death according to the concentration of supplemental ALS plasma (1% - 0.69 ± 0.35%, 3% - 6.06 ± 0.11%, and 5% - 8.57 ± 0.63%) and changes in live cell morphology, suggesting the influence of the pathological environment upon cell survival. These results were expected due to unfavorable humoral factor content in ALS peripheral blood. Meta-analysis of various cytokines in G93A SOD1 mutant mice demonstrated increased levels of pro-inflammatory type I cytokines (IL-1β, IL-1α, IL-12, and TNF-α) vs. anti-inflammatory type II cytokines (IL-4, IL-6, IL-10) across all disease stages with more pronounced elevations at end stage disease (Jeyachandran et al. 2015). Moreover, when Moreno-Martínez et al. (Moreno-Martínez, de la Torre, et al. 2019) recently analyzed 97 cytokines in plasma from G93A SOD1 mutant mice at different disease stages, they reported increased levels of several cytokines that were associated with a shorter mouse lifespan. However, the authors noted that these cytokines may be inadequate prognostic disease biomarkers due to their expression variability. A review from the same authors (Moreno-Martínez, Calvo, et al. 2019) detailed concerns of inflammatory cytokines’ reliability as a diagnostic tool for ALS. ALS patients have also shown significantly increased levels of peripheral blood inflammatory cytokines such as TNF-α, TNF receptor 1, IL-6, and IL-1β (Ehrhart et al. 2015; Hu et al. 2017; Lam et al. 2016). Additionally, the potential role of humoral IL-6 cytokine in mediating EC inflammation via the trans-signaling pathway has been discussed (Garbuzova-Davis, Ehrhart, et al. 2018). Thus, numerous up-regulated circulating inflammatory cytokines found in both ALS patients and an animal model may not only contribute to neuroinflammation, but also impede viability of cells transplanted into the blood circulation. Also, the pathological humoral environment in ALS can affect the integrity of endogenous endothelial cells since these cells, lining the capillary lumen and exposed to circulating blood, may be vulnerable to various harmful blood substances. However, confirmatory studies to elucidate pro- and anti-inflammatory cytokines in ECs isolated from ALS mice during disease progression as well as to correlate EC cytokine levels with those in mouse peripheral blood are planned for a future investigation. Also, since hBM-EPCs showed increased cell death upon exposure to ALS mouse plasma, additional studies are needed to elucidate cell response effect to control mouse plasma in vitro. This study may be imperative to determine efficacy of hBM-EPCs by administration into control non-ALS mice.

To address this issue, mouse brain endothelial cells (mBECs) were used in our follow-up study to determine whether ALS mouse plasma worsens cell viability and to evaluate the effect of hBM-EPC-derived EVs upon induced EC damage in vitro. Results showed substantial cell death after exposure to 3% ALS mouse plasma and hBM-EPC-derived EVs at 1 μg/mL concentration significantly alleviated mBEC damage and improved cell morphology, suggesting an efficacious EV effect. Interestingly, a dose-response study established that EVs at 5 μg/mL increased cell death and may be considered toxic to EC survival at this concentration. Potentially, the observed toxicity is due to this dosage being at the far end of a dose-response bell curve as commonly observed in testing of pharmaceutical drugs. However, mechanism(s) of this high-dose EV toxicity is unknown and warrants future investigation.

Finally, hBM-EPC-derived EV actions leading to alleviation of EC damage from the pathological environment were elucidated. Adding 1 μg/mL of GFP-labeled EVs into culture media supplemented with 3% ALS mouse plasma showed vesicle uptake by numerous mBECs. The GFP-EVs were widely distributed in the cytosol and cell processes. These results may provide support for the beneficial effect of hBM-EPC-derived EVs on damaged ECs by potential transfer of various bioactive molecules sustaining cell functionality. However, a multiomics approach to identify transcripts for intracellular signaling pathways, metabolites, cytokines, biological markers, and/or proteins in EVs may be imperative for elucidating reparative cell mechanisms. Since EVs comprise a heterogenous group of vesicles representing diverse sets of biological processes (reviewed in (Akers et al. 2013; EL Andaloussi et al. 2013; György et al. 2011)), specific surface biomarker expressions were identified for exosomes such as CD9, CD63, CD81 tetraspanins and flotillin (Kalluri and LeBleu 2020; Kowal et al. 2016; Rana and Zöller 2011; Raposo and Stoorvogel 2013); microvesicles – integrins, selectins, and CD40 ligand (Cocucci et al. 2009; Deregibus et al. 2007); and apoptotic bodies – annexin V, thrombospondin, and C3b (Igami et al. 2020; van Engeland et al. 1998; van Genderen et al. 2008). Defining the ratio of these marker expressions in hBM-EPC-derived EVs may afford better characteristics of vesicle subpopulations. Additionally, determining specific cargo proteins in EVs via proteomics, protein(s) responsible for restoration of EC integrity under pathological condition, may provide vital information. Proteomics is a powerful method allowing detection of numerous vesicular proteins with “essential clues to the molecular mechanisms involved in vesicle cargo sorting and biogenesis” (Choi et al. 2015). The authors also noted the importance of this EV analysis for elucidating diagnostic and therapeutic target proteins. The EVs may also contain RNA that contribute to beneficial vesicle functions, a possibility discussed in comprehensive reviews (Hill et al. 2013; Kim et al. 2017; Mateescu et al. 2017). Furthermore, EVs are lipid-bilayered vesicles and composed of numerous membrane lipids, which play an essential role for the stability, rigidity, and function of EVs (reviewed in (Yoon et al. 2014)). Since the specific lipid components in EVs are dependent upon the originating cells (Subra et al. 2007), investigating lipid composition in hBM-EPC-derived EVs may identify their functional roles not only in maintaining vesicle structure, but also in modulating intracellular fusion and fission of EVs (Chernomordik and Kozlov 2003). In this context, characteristic of the protein-to-lipid ratio “may prove useful for quality control of extracellular vesicle related basic and clinical studies” (Osteikoetxea et al. 2015). Thus, these important studies mentioned above are planned for our future investigations. Though, EV constituents via biomarker expressions and proteomics are currently under our investigation.

Mechanisms of EV uptake, however, need further discussion. A comprehensive review by Mulcahy et al. (Mulcahy et al. 2014) summarized pathways by which EVs taken up into target cells, such as fusion with the plasma membrane or intracellular membrane, various types of mediated endocytosis, macropinocytosis, and phagocytosis. The authors emphasized that EVs internalization into cells may occur via multiple routes and may “depend on proteins and glycoproteins found on surface of both the vesicle and the target cell” (Mulcahy et al. 2014). Despite different origins of pathways (endolysosomal or budding from the plasma membrane), in which EVs are generated, numerous studies support an endocytic uptake mechanism as the primary entry route of EVs into recipient cells (reviewed in (Colombo et al. 2014; EL Andaloussi et al. 2013; Mulcahy et al. 2014)). However, fusion of EV membrane with the cell plasma membrane of receiving cells should be considered as well. Also, inhibition of EV uptake by ECs via blocking ligand and/or receptor-mediated internalization (reviewed in (Chaput and Théry 2011; EL Andaloussi et al. 2013; Mulcahy et al. 2014)) may verify therapeutically relevant roles of EVs demonstrated in our current in vitro study. A study by Deregibus et al. (Deregibus et al. 2007) investigated the role of adhesion molecules in the incorporation of MVs into human EPCs and showed that blocking only α4 integrin and β1 integrin (CD29) inhibited MV internalization in cells in contrast to ICAM-1 or α6 integrin. In our study, pre-incubation of EVs with anti-CD29 blocking antibody demonstrated prevention of GFP-labeled EV incorporation in mBECs after exposure to 3% ALS mouse plasma and as result the percentage of dead mBECs was significantly increased vs. EVs treatment alone at the same concentration. Thus, our results support a previous finding (Deregibus et al. 2007) regarding importance of specific adhesion molecules in mediating EV uptake by target cells.

In summary, the present study results demonstrated that hBM-EPC-derived EVs effectively isolated from cell culture media were categorized as nanosized vesicles. These EVs significantly reduced mouse brain endothelial cell damage from the pathological environment in vitro by internalization and probable transfer of cargo bioactive molecules into damaged cells. Although identification of specific protein(s) responsible for mediating EC integrity is still underway, our study evidence suggests that EVs released from hBM-EPCs promote endothelial cell repair and can be promising novel cell-free therapeutics for restoration of the B-CNS-B in ALS. However, whether the protective effect of EVs upon damaged cells is durable and cell injury is reversible are currently unknown. Also, one limitation in our study is that the in vitro results need to be evaluated in vivo by EV administrations into ALS mice to confirm therapeutic efficacy of vesicles for restoration of the damaged microvascular endothelium. Since we just initiated an investigation on development of an extracellular vesicle-based approach for ALS therapy towards restoration of CNS barrier integrity, further optimization of this cell-free approach toward translation into clinical application will be addressed in our future animal studies.

Acknowledgments

This study was supported by the NIH, NINDS (1R01NS090962) grant. We thank Dr. Bickford (Center of Excellence for Aging & Brain Repair, Department of Neurosurgery and Brain Repair at the University of South Florida, Morsani College of Medicine, and James A Haley VA Hospital, Tampa) for technical assistance.

Footnotes

Conflicts of Interest

The authors declare no conflict of interest.

Publisher's Disclaimer: This Author Accepted Manuscript is a PDF file of an unedited peer-reviewed manuscript that has been accepted for publication but has not been copyedited or corrected. The official version of record that is published in the journal is kept up to date and so may therefore differ from this version.

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