A standardized method to functionalize plasma-extracellular vesicles via copper-free click chemistry for targeted drug delivery strategies

doi:10.21203/rs.3.rs-3148779/v1

Download PDF

Method Article

A standardized method to functionalize plasma-extracellular vesicles via copper-free click chemistry for targeted drug delivery strategies

https://doi.org/10.21203/rs.3.rs-3148779/v1

This work is licensed under a CC BY 4.0 License

Version 1

posted

You are reading this latest preprint version

Background: Extracellular vesicles (EVs) have emerged as potential vehicles for targeted drug delivery and diagnostic applications. However, achieving consistent and reliable functionalization of EV membranes remains a challenge. Copper-catalyzed click chemistry, commonly used for EV surface modification, poses limitations due to cytotoxicity and interference with biological systems. To overcome these limitations, we developed a standardized method for functionalizing EV membrane via copper-free click chemistry. EVs derived from plasma hold immense potential as diagnostic and therapeutic agents. However, the isolation and functionalization of EVs from such a complex biofluid represents a considerable challenge.

Results: We compared three different EV isolation methods to obtain an EV suspension with an optimal purity/yield ratio, and we identified sucrose cushion ultracentrifugation (sUC) as the ideal protocol. We then optimized the reaction conditions to successfully functionalize plasma-EV surface through a copper-free click chemistry strategy with a fluorescently labeled azide, used as a proof-of-principle molecule. Click-EVs maintained their identity, size, and, more importantly, the capacity to be efficiently uptaken by responder tumor cells. Moreover, once internalized, click-EVs partially followed the endosomal recycling route.

Conclusions: The optimized reaction conditions and characterization techniques presented in this study offer a foundation for future investigations and applications of functionalized EVs in drug delivery, diagnostics, and therapeutics.

extracellular vesicles

surface functionalization

click chemistry

tumor cells

Nanoscience, the study and manipulation of matter at the nanoscale, has emerged as a burgeoning field with vast implications for numerous scientific disciplines. At the heart of this fascinating field lies the visionary contributions of Richard Feynman, a renowned physicist and Nobel laureate, whose seminal lecture in 1959 sparked the modern-day interest in the potential of nanoscience [1]. In the last decade, the possibility to apply nanotechnologies in biomedicine for both diagnosis and therapy of human diseases has increased considerably [2][3][4]. In this context, targeted drug delivery has emerged as a promising strategy to enhance the efficacy and specificity of therapeutic interventions while minimizing off-target effects. While various platforms, including artificial nanoparticles, have been investigated for targeted drug delivery [5], the remarkable potential of extracellular vesicles (EVs) has captured the attention of the scientific community due to their unique properties and inherent biological advantages [6].

EVs are small membranous vesicles released by cells into the extracellular space [7]. They play crucial roles in intercellular communication by transferring bioactive molecules, such as proteins, lipids, and nucleic acids, between cells. EVs can be broadly classified into exosomes, microvesicles, and apoptotic bodies, based on their biogenesis, size, and molecular composition [8] [9].

Functionalizing the membrane of EVs offers a myriad of possibilities, including the incorporation of specific ligands, therapeutic agents, or imaging probes [10]. These modifications can enhance the targeting efficiency, stability, and cargo delivery capabilities of EVs, expanding their therapeutic and diagnostic applications [11]. However, to achieve optimal functionalization, it is essential to start with a reliable and representative source of EVs. In addition to circumventing the challenges associated with the use of cells as a source for extracellular vesicles, isolating EVs from biofluids such as plasma provides the possibility of using a holistic and comprehensive vesicle population, reflecting the complex and dynamic nature of EVs in physiological conditions.

In this work, we propose a fast and simple chemical-conjugation-based method to functionalize the EV membrane. In particular, our strategy provides, for the first time, evidence of the engineering of plasma-derived EVs by a copper-free click chemistry alkyne-azide cycloaddition. This bioorthogonal reaction provides a powerful tool for site-specific functionalization of EV membranes with various molecules of interest [12] [13] [14].

With the purpose of obtaining an EV suspension with a tolerable purity and sufficient enrichment to allow downstream applications, we compared three different isolation protocols, namely size-exclusion chromatography (SEC), sucrose cushion ultracentrifugation (sUC) and a combination of iodixanol density ultracentrifugation and SEC (IDC+SEC). Size, concentration and purity analyzed by nanoparticle tracking analysis (NTA), transmission electron microscopy (TEM), western blot (WB), and enzyme-linked immunosorbent assay (ELISA) confirmed that the sUC method represented the optimal compromise between purity and yield.

We focused on the EV membrane functionalization using a reporter molecule, a fluorescent azide, to be used in a two-step copper-free click chemistry reaction. The use of a fluorescent azide as a reporter molecule allows for straightforward and reliable detection of functionalized EVs, facilitating the optimization and quality control of the membrane engineering procedure. Functionalized EVs were able to be efficiently internalized by responder tumor cells in a time-dependent manner, being in part recycled by the endosomal compartment of the target cells. The standardization of the procedures here described provides a robust foundation for future studies, where more specific and tailored molecules could be employed to impart desired functionalities to EV membranes.

Sucrose cushion ultracentrifugation represents the optimal compromise between purity and yield for plasma-EV isolation
Plasma is one of the most challenging biofluids for the isolation of EVs, as it contains a massive pattern of proteins, protein aggregates, as well as a large set of lipidic nanoparticles with size or density comparable to EVs [15]. To evaluate the appropriate EV isolation procedure that could guarantee the optimal compromise between yield and purity, we compared three different protocols: size-exclusion chromatography (SEC), a combination of iodixanol density gradient and SEC (IDC+SEC) [15], and a sedimentation rate-based method (sucrose cushion ultracentrifugation, sUC) [16]. The latter has been performed using either 1 or 2.3 ml as starting volume of plasma.
The concentration of contaminating HDLs and LDLs/VLDLs in plasma-EVs separated by the three different methods has been measured by ELISAs specific for Apolipoprotein A1 (ApoA1) and Apolipoprotein B100 (ApoB100), respectively (Fig. 1a and 1b).

While ApoA1 expression increased by 13-fold in SEC-EVs compared to sUC-EVs-1ml, sUC-EVs-2.3ml, and IDC+SEC-EVs, no significant differences were observed among sUC-EVs from 1ml, sUC-EVs from 2.3ml and IDC+SEC-EVs [sUC-1mL: 9.0x10⁴ + 0.6 ng/mL; sUC-2.3mL: 8.7x10⁴ + 0.1 ng/mL; SEC: 120.2x10⁴ + 4.9 ng/mL; IDC+SEC: 8.9x10⁴ + 0.4 ng/mL; p<0.0001; N=3] (Fig. 1a) Similarly, the concentration of ApoB100 was significantly higher in samples separated by SEC compared to the other considered protocols (Fig. 1b). The concentration of ApoB100 detectable in SEC-EVs was respectively about 110, 88 and 330-fold more compared to sUC-EVs-1ml, sUC-EVs-2.3ml, and IDC+SEC-EVs [sUC-1mL: 3.1 + 0.4 x10² ng/mL; sUC-2.3mL: 3.9x10² + 0.6 ng/mL; SEC: 34.30x10² + 13.4 ng/mL; 1.0x10² + 0.1 ng/mL, p<0.0001; N=3] (Fig. 1b). To eliminate the ApoB100-expressing lipoproteins, the IDC+SEC method turned out to be the optimal method, while we didn’t observe significant differences in ApoB100 concentration applying the sUC method to 1 or 2.3 ml of plasma (Fig. 1b). Accordingly, we decided to adopt the modified method (sUC-2.3mL) and we compared by NTA the size distribution and concentration of EVs isolated by SEC, IDC+SEC and sUC-2.3ml (Fig. 1c and 1d). NTA revealed that no significant differences in terms of size distribution were observed among the three different isolation protocols. Indeed, the X50 parameter, indicating the diameter of particles where 50% of the distribution has a smaller and 50% has a larger particle size, was 130.0±4.5 nm,146.1±12.7 nm and 134.9±10.3 nm for sUC-, SEC and IDC+SEC-EVs, respectively (Fig. 1c). Nevertheless, significant differences in terms of EV concentration were observed comparing the separation protocols (Fig. 1d). IDC+SEC was the one resulting in the lowest amount of EVs, being able to isolate 11-fold less EVs than the sUC method and 16-fold less than the SEC method [sUC: 2.5 + 0.8 x10⁹; SEC: 3.5 + 0.7 x10⁹; IDC+SEC: 2.2 + 0.3 x10⁸; sUC vs IDC+SEC: p=0.0014; SEC vs IDC+SEC: p=0.0001; N=4] (Fig. 1d). Taken together, these data indicate that sUC is the ideal method for enrichment of EVs from human plasma, representing the optimal compromise between EV purity and yield compared to the other two considered protocols.
Therefore, we focused on sUC-EV characterization (Fig. 2).
Tetraspanin (CD9, CD81 and CD63) expression was evaluated by non-conventional flow cytometry [17] (Fig. 2a, 2b, and Supplementary Fig.1a) and by super resolution microscopy (Fig. 2c, 2d).

Both techniques indicated that CD9 is the most abundant tetraspanin expressed by plasma-EVs (Fig. 2b, 2d). Interestingly, the characterization at single EV level revealed a heterogeneous EV population expressing different tetraspanin combinations (Fig. 2d), even if CD9 is confirmed as the most expressed one. Moreover, sUC-EVs expressed the typical EV markers Alix, Flotillin-1, TSG-101 and Syntenin, as revealed by western blot analysis (Supplementary Fig.1b).
NTA showed the presence of a homogeneous size distribution with an average X50 of 130.0±4.5 nm (Fig. 2e). In particular, 10% of the analyzed particles were smaller than 69.1±2.5 nm and 90% of them were smaller than 233.3±8.8 nm (Fig. 2e).

Copper-free click chemistry has no impact on plasma-EV identity and size, but severely affects their concentration
A copper-free click chemistry approach has been used to functionalize the plasma-EV membrane. The method has been set-up using, as reagents, the alkyne Dibenzocyclooctyne-N-hydroxysuccinimidyl ester (DBCO-NHS) and a fluorescently labelled azide (AZ-647). The latter has been selected as a proof-of-principle molecule to standardize the whole procedure. Firstly, the aminic groups of the EV surface proteins reacted with the N-hydroxysuccinimidyl (NHS) group of DBCO. The alkyne group of DBCO was then coupled to AZ-647 by copper-free click chemistry. As already reported [12], the highest labelling yield was obtained when EVs were reacted with DBCO-NHS for 1h at RT and then immediately coupled with AZ-647 for 4h at RT. To obtain an efficient plasma-EV labeling, the particle number to be used in the click chemistry reaction has been titrated (Fig. 3a-e).

AZ-647 mixed with the EV suspension, without DBCO-NHS, served as negative control. Titration experiments, conducted using 5 different EV amounts (from 10⁶ to 5x10⁹) with fixed concentrations of both DBCO-NHS and AZ-647, revealed that the incubation with only AZ-647 did not induce an unspecific EV labelling (Fig. 3a-e, grey histograms) and, more importantly, the reaction was successful when working with at least 10⁹ EVs (Fig 3d, 3e, blue histograms). Indeed, experimental replicates conducted with 10⁹ EVs indicated that the percentage of labeled EVs, ranging from 40 to 60% of total EVs used in the reaction, was significantly higher than the corresponding negative control (Fig. 3f; p<0.0001; N=11) and these results were confirmed by fluorescent NTA analysis (Fig. 3g; p<0.0001; N=4). Subsequent analyses were performed to evaluate whether click chemistry reaction affected the EV identity. The expression of the vesicle markers Flotillin-1, CD63 and Syntenin did not vary in Click-EVs compared to both naïve EVs and EVs incubated with only AZ-647 (Fig. 3h). These data were confirmed by TEM and NTA analysis which did not show any significant variation in EV size among naïve and corresponding Click-EVs (Fig. 3i, 2e and 3l). On the contrary, click chemistry severely affected EV concentration, with a loss of 80-90% in both negative control and Click-EVs, compared to naïve sample (Fig. 3m).
These results suggest that click chemistry doesn’t affect EV size and identity, but it profoundly affects the final EV number, and this is probably due to the washing step necessary after EV labelling to remove the unlabeled reagents and to possible EV aggregation. This observation explains why it was so important to identify an EV isolation protocol that allows obtaining a sufficient number of vesicles to be efficiently functionalized.
Click-EVs are internalized by responder cells in a time dependent manner and follow the endosomal recycling route
The capacity of Click-EVs to be internalized by responder cells (MDA-MB-231 tumor cell line) was evaluated. Responder cells were analyzed by flow cytometry after 18h of co-incubation with either negative control (1x10⁹ EVs+AZ-647) or 1x10⁹ Click-EVs. Untreated cells were used as an additional negative control. No fluorescence was detected with neither untreated cells nor cells incubated with EVs+AZ-647 (Fig. 5a), confirming the previous results related to the absence of an unspecific EV labeling given by the incubation with the fluorescent azide (Fig. 4f, 4g). Interestingly, 54.6 + 22.6% of cells treated with Click-EVs were AF-647 positive, clearly indicating an efficient uptake of Click-EVs (Fig. 5a).

In addition, we evaluated by confocal microscopy the timing and the recycling route of Click-EVs within responder cells at three different time points: 3h, 10h and 18h (Fig. 4b). A few Click-EVs were internalized by responder cells after 3h incubation (Fig 4b, upper-left panel) and their numbers increased during time, as determined by the increased fluorescence/cell (Fig 4b, bottom-right panel). At 10 h, the mean fluorescence intensity associated with internalized EVs was almost 20 times higher than the one measured at 3h and augmented even more at 18 h (50 times more than 3h), indicating that Click-EVs internalization progressively increased with time (Fig 4b, bottom-right panel).
Moreover, analyzing the distribution of Rab5, an early endosome marker, we observed, at longer times, the presence of Rab5 surrounded Click-EVs (Fig. 5).
In particular, both the number of cells displaying Click-EVs surrounded by Rab5 and the number of Rab5-surrounded EVs/cell increased progressively from 10h to 18h (Fig 5, bottom histogram). Taken together, these data indicate that Click-EVs are efficiently internalized after 10h co-incubation with responder cells, and partially follow the endosomal recycling route.

EVs, including exosomes and microvesicles, are naturally released by various cell types and are involved in intercellular communication by transferring bioactive molecules, including proteins, nucleic acids, and lipids, between cells [18]. This intrinsic function makes EVs an attractive option for targeted drug delivery, as they can be engineered to carry therapeutic agents to specific target cells or tissues [19]. By modifying the surface of EVs with targeting ligands, such as antibodies or peptides, it is possible to enhance their specificity, and improve their delivery efficiency while potentially reducing adverse and side effects [20].

Nowadays, all commercially available nanoparticle-related therapeutics are based on artificial ones, especially liposomes, with about 14 liposomal products authorized by FDA and EMA [21]. Similarities and differences between liposomes and natural nanoparticles, particularly in protein and lipid composition, are under discussion [22] [23] [24]. Low immunological response [25], targeting potential [26], and ability to cross various biological barriers [27] [28] [29] [30] [31] emerge as the most considerable advantages of using EVs as drug delivery vehicles. In addition, based on the different cell sources, the employed isolation procedure, or the specific protein/lipid profile, EVs could present longer clearance kinetics than artificial nanovesicles [24].

The asymmetrical lipid distribution and specific protein composition of EV membrane can also justify their peculiar organotropism and homing ability [31] [32]. It has been demonstrated that cancer-derived EVs show a tissue tropism also at the early stages of neoplastic transformation [33], and that the transplantation of autologous extracellular vesicles can be used for cancer-specific targeting [34]. Especially based on this evidence, biofluids, and in particular blood, would represent an ideal source of EVs to be functionalized for targeted drug delivery or diagnostic purpose. The addition of the “acquired” to the natural homing characteristic could result in a more efficient targeting effect. In this regard, our study builds upon existing literature that has primarily focused on the functionalization of EVs derived from cell supernatants [13] [35]. Although these studies have provided valuable insights into EV biology and targeting strategies, the isolation of EVs from plasma represents a significant advancement in the field. Several recent reports have highlighted the unique characteristics and potential advantages of plasma-derived EVs. For instance, plasma-EVs are more likely to interact with target cells in vivo due to their presence in the systemic circulation [36]. Therefore, developing a standardized methodology to functionalize plasma-derived EVs could have broader implications for targeted drug delivery and personalized medicine. Furthermore, it has to be taken into account that plasma is one of the most difficult body fluids to isolate EVs from, due to the presence of a massive pattern of particles [37]. Plasma proteins and protein aggregates, as well as the large set of lipidic nanoparticles, represent the most abundant contaminants [38]. The commonly used methods to separate plasma-EVs are based on size or density. Recently, it has been observed that combining these two isolation strategies it’s possible to remove a large part of lipoproteins [15]. However, this combined strategy profoundly affects the yield, an important parameter to be considered when EVs have to be applied in downstream procedures which require a high number of vesicles. To obtain enough and highly pure EVs we selected a sedimentation rate- (sucrose cushion ultracentrifugation-sUC) method which has been in the past efficiently used for EV-isolation from conditioned media [39] and more recently also successfully applied to isolate EVs from plasma [16].

The standardization of plasma-EV membrane functionalization is of paramount importance for several reasons. Firstly, a standardized approach would enable the comparison of results across different laboratories and facilitate the exchange of EV-based therapeutic strategies, ultimately accelerating the progress in the field. Secondly, a standardized methodology would enhance the reproducibility of EV-based drug delivery studies, allowing researchers to validate and build upon previous findings. Additionally, a consistent methodology would aid in the evaluation and regulatory approval of EV-based therapeutics, ensuring their safe and effective use in clinical settings.

We adopted a copper-free click chemistry approach to functionalize the EV membrane. Copper-free click chemistry is an alternative approach to click chemistry that proceeds at a lower activation barrier and is free of cytotoxic transition metal catalysts [40] [41]. Indeed, copper-free click chemistry approaches offer distinct advantages over copper-catalyzed click reactions, which can be cytotoxic and potentially affect the integrity and functionality of EVs [42]. By eliminating the need for copper ions, this bioorthogonal reaction reduces potential cytotoxicity concerns, and ensure the preservation of EV structure and biological activity [43]. This enables the precise modification of EV membranes without compromising their native properties, thereby enhancing their therapeutic potential and targeting capabilities. Moreover, the alkyne-azide cycloaddition reaction exhibits excellent selectivity, efficiency, and biocompatibility [44]. The reaction proceeds rapidly under physiological conditions, allowing for efficient modification of EV membranes in a controlled manner. We here adopted a reporter molecule, a fluorescent azide, to functionalize the EV membrane. The use of a fluorescent azide served two purposes in our study. Firstly, it allowed us to track and visualize the successful incorporation of the molecule onto the EV membrane. This provided a convenient and reliable method to assess the efficiency and homogeneity of the functionalization process. Secondly, the fluorescent azide served as a model molecule, enabling us to validate the applicability and efficacy of our EV membrane functionalization protocol. By replacing the fluorescent azide with azo-modified sequence-specific molecules, our protocol could be easily adapted to incorporate diverse targeting moieties in the EV membrane. We observed that the whole functionalization procedure did not affect EV integrity and identity, but profoundly impaired their number. This further emphasizes the importance of selecting an isolation protocol that allows the recovery of a sufficient EV number. There is no consensus regarding the mode of EV uptake, which could be receptor-dependent or not. Moreover, the fate of EV within the recipient cell is not clearly characterized. Only recently it has been proposed that the EV cargo could be released in the cytosol, degraded within lysosomes or re-secreted through recycling within newly formed EVs, exploiting the endosomal compartment of the cells [45]. By incubating click-EVs with responder cells we focused on the analysis of two aspects: the timing of the uptake and their recycling route into cells. We could confirm not only that clicked-EVs were efficiently internalized and able to transfer their fluorescent signal but also that they partially followed the same endosomal recycling route followed by unmodified EVs.

Through the application of copper-free click chemistry, the engineering of plasma extracellular vesicle membranes provides a promising approach for tailoring EVs to specific therapeutic or diagnostic applications. By precisely functionalizing the EV membrane, it’s possible to enhance their stability, targeting efficiency, and cargo delivery capabilities, thereby unlocking the full potential of EVs as versatile and effective drug delivery vehicles or diagnostic tools.

Our study highlights the importance of standardizing the methodology for EV membrane functionalization to advance the field of targeted drug delivery. By focusing on plasma-derived EVs, we have demonstrated the novelty and clinical relevance of our approach, which complements existing literature that primarily utilizes EVs derived from cell supernatants. Moving forward, efforts should be directed towards establishing standardized protocols that can be readily implemented by researchers in the field, thereby facilitating the translation of EV-based drug delivery systems into clinical practice.

Blood collection and processing

Blood samples were provided by the Institutional Transfusion Center of Ospedale Policlinico San Martino, Genova. Blood was collected from healthy donors in K2E EDTA tubes. Plasma was isolated from blood by serial centrifugation steps, performed at 4°C: 120xg 20 min, 120xg 5 min, 360xg 20 min and 3000xg 10 min. The resulting plasma was stored at –80 °C until EV separation.

EV isolation

Three different EV isolation protocols were tested.

Size exclusion chromatography (SEC) was performed using IZON smart columns qEV/70. According to manufacturer’s instructions, 500 µL of plasma was loaded on the column and eluted with PBS containing 2mM EDTA (PBS-EDTA) to overcome EV aggregation. Twenty eluted fractions (F) of 500 μL were collected in protein low binding tubes, and EVs were collected in fractions 7-9. The elution profile of both EVs and contaminating proteins has been assessed measuring the absorbance at 280nm. Samples were further diluted depending on subsequent analysis. The EV-enriched fractions have been rinsed in dPBS and centrifuged at 100 000×g for 2h with a SN985 swinging rotor (Optima TL, Beckman).

Regarding the combination of iodixanol density gradient and SEC (IDC+SEC) method, we followed a previously described protocol [15]. Briefly, 6 mL of plasma was layered on top of a 2 mL 50%, 2 mL 30%, and 2 mL 10% OptiPrep layer. Cushion and sample were centrifuged at 178.000×g for 2 h at 4 °C in a SW41It swinging rotor. The visible band between the 10% and 30% layers was collected (high-density band, 1.06-1.16g/cm³; EV enriched band) and loaded onto a SEC column. The low-density band (< 1,025g/cm³) on the top of the tube, enriched in lipoproteins, was discarded. The EV enriched fractions have been further centrifuged at 100.000×g for 2h with a SN985 swinging rotor (Optima TL, Beckman).

The sucrose cushion ultracentrifugation (sUC) was performed as previously reported [16]. Either 1 mL or 2.3 mL (the latter amount has been modified from the original protocol) of plasma were diluted in 0.22 mm filtered dPBS to a final volume of 9.5mL, carefully loaded on the top of 2mL 20% sucrose and centrifuged at 100 000×g for 135 minutes in a SW41Ti swinging rotor. Then, the supernatant was discarded and the EV pellet was resuspended in 1ml of PBS-EDTA, centrifuged at 3000×g for 10 minutes, and the supernatant has been further centrifuged at 100 000×g for 2h with a SN985 swinging rotor (Optima TL, Beckman).

EV quantification by Nanoparticle Tracking Analysis (NTA)

EV size distribution and concentration were analyzed by ZetaView® TWIN-NTA PMX-220 (Particle Metrix GmbH, Inning am Ammersee, Germany), equipped with a sample cell and two lasers (488 nm and 640 nm) and Zetaview 8.05.14_SP7 software. Briefly, after calibration with 100 nm polystyrene beads, samples were diluted in filtered dPBS and injected into the sample cell using a 1 mL syringe. For both size and concentration measurements, the 488 nm laser in scatter mode was used; for the fluorescent NTA, 640 nm laser with a 660 nm filter wavelength was used. Size distribution analyses of 11 different positions were performed for each sample and the reading parameters were set as 80% sensitivity, and shutter speed of 100, minimum brightness of 20. For fluorescent NTA, sensitivity was increased to 90%, minimum brightness of 30.

ELISA

EV-enriched samples were diluted in dPBS (100,000-times for measurements of ApoA1 in all samples, 3000-times for measurements of ApoB100 in samples after SEC and 80-times for measurements of ApoB100 in samples after sUC) and the concentration (ng/mL) of

ApoA1 and ApoB100 measured in duplicates by specific ELISAs (ELISA-ApoA1: #3710-1HP-2, ELISA-ApoB100: #3715-1HP-2, Mabtech, Sweden), according to the manufacturer’s instructions. Measured concentrations were normalized to 1 mL of starting plasma and any potential dilution of the samples was accounted for.

EV characterization by flow cytometry

EVs were characterized by non-conventional flow cytometry, as previously described[17]. Briefly, 1 ×10⁸ EVs in 100µL of PBS-EDTA were stained with 1mM CFDA-SE (Vybrant™ CFDA-SE Cell Tracer Kit, Thermo Fisher Scientific, Waltham, MA, USA) at either 4°C or room temperature (RT). The expression of CD9 (APC Mouse Anti-Human CD9, Clone HI9a, 312108; BioLegend, San Diego, CA, USA), CD63 (PE-Cy7 Mouse Anti-Human CD63, Clone H5C6, 561982; BD Biosciences, San Jose, CA, USA ), and CD81 (BV421 Mouse Anti-Human CD81, Clone JS-81, 740079; BD Biosciences ) was evaluated within the CFDA-SE-positive events and compared to the corresponding isotype controls using CytoFLEX S (Beckman Coulter). FlowJo software was used to analyze the data.

Western blot

EVs were resuspended in RIPA buffer (1% NONIDET p-40, 0.1% SDS, 0.1% Sodium deoxycholate, protease inhibitor cocktail 1x, in PBS pH7.5) and protein content was quantified by Bicinchoninic acid (BCA) assay (Thermo Fisher Scientific). Five μg of proteins for each sample were loaded on 4%–12% NuPAGE Bis–Tris gel (Thermo Fisher Scientific). Electrophoresis was performed at 160 V and proteins were blotted on a polyvinylidene fluoride membrane. After blocking nonspecific sites with 5% non-fat dry milk (EuroClone, Italy) in Tris Buffered Saline with Tween 20 (TTBS, 20 mM Tris pH 7.5, 500 mM NaCl, 0.05% Tween 20), the membrane was incubated overnight at 4 °C with specific primary antibodies for: Flotillin-1 (Recombinant Anti-Flotillin 1 antibody [EPR6041] (ab133497), 1:10 000, Abcam), Syntenin (Recombinant Anti-Syntenin antibody [EPR8102] (ab133267), 1:1000, Abcam), Alix (Recombinant Anti-ALIX antibody [EPR15314] (ab186429), 1:1000, Abcam), TSG-101 (Anti-TSG101 antibody produced in rabbit, 1:1000 dilution, T5701, SIgma) diluted in 2.5% non-fat dry milk/TTBS. After washing three times with TTBS, membranes were incubated with specific HRP-conjugated secondary antibodies anti-mouse (Mouse IgG HRP Linked Whole Ab, GENA931) or anti-rabbit (Rabbit IgG HRP Linked Whole Ab, GENA934). Positivity was highlighted by providing the substrates for the chemiluminescence reaction of HRP (Amersham ECL Prime Western Blotting Detection Reagent, GE Healthcare, Chicago, Illinois, USA) and impressing a photographic sheet by autoradiography (GE Healthcare).

Super resolution microscopy

Super-resolution microscopy analysis of plasma-EVs was performed using Nanoimager S Mark II microscope from ONI (Oxford Nanoimaging, Oxford, UK) equipped with a 100x, 1.4NA oil immersion objective, an XYZ closed-loop piezo 736 stage, and triple emission channels split at 640, 488 and 555 nm. The EV profiler Kit (ONI) was utilized for the experiments following manufacturer’s protocol. The Kit is composed by fluorescent antibodies anti CD9-488, CD63-568 and CD81-647 and all the buffer and reagent necessary for the experiment. dSTORM mode acquired sequentially in total reflection fluorescence (TIRF) mode was used for the acquisition of images. Single-molecule data was filtered using NimOS software (v.1.18.3, ONI). Data has been processed with the Collaborative Discovery (CODI) online analysis platform www.alto.codi.bio from ONI and the drift correction pipeline version 0.2.3 was used [46].

Transmission Electron Microscopy (TEM)

EVs were resuspended in 20 μl dPBS and fixed by adding an equal volume of 2% paraformaldehyde in 0.1 mol/l phosphate buffer (pH 7.4), as previously described [47]. EVs were then adsorbed for 10 min to formvar-carbon coated copper grids by floating the grids on 5 μl drops on parafilm. Subsequently, grids with adhered vesicles were rinsed in PBS and negatively stained with 2% uranyl acetate for 5 min at room temperature. Stained grids were embedded in 2.5% methylcellulose for improved preservation and air dried before examination. Electron micrographs were taken at Hitachi TEM microscope (HT7800 series, Tokyo, Japan) equipped with Megaview 3 digital camera and Radius software (EMSIS, Germany). To visualize EV size distribution, the results were plotted as colorblind safe scatter dot plot in which each size measured is represented as a point along with lines for the median value and the range.

EV surface functionalization by copper-free click chemistry

EVs (from 10⁶ to 5x10⁹ for the titration experiments, and in a range between 10⁹ to 10¹⁰ for all the other experiments) were resuspended in 200µL of PBS-EDTA and mixed for 1h at RT with 1μg of DBCO-NHS ester (Dibenzocyclooctyne-N-hydroxysuccinimidyl ester for copper free click chemistry, 761524, Sigma). After the first incubation, 1.9μl of a 5.9mM AlexaFluor647- azide (AZ-647) solution (Alexa Fluor™ 647 Azide, Triethylammonium Salt, A10277, ThermoFisher) was added to the reaction and incubated for 4h at RT (Click-EVs). The fluorescent azide mixed with the EV suspension (without DBCO-NHS ester) was considered as negative control (EV + AZ-647). After the last incubation, samples were washed by SEC to remove unlabeled chemicals, stained with CFDA-SE, and analyzed by flow cytometry to evaluate, within the CFDA-SE positive events, the AF647 fluorescence signal associated with the azide. For uptake experiments, Click-EVs and the negative control were concentrated by ultracentrifugation (3h, 100 000×g, 4°C) and resuspended in 100µL of phenol red-free Dulbecco's modified eagle medium (DMEM).

EV internalization

The triple negative human breast cancer cell line MDA-MB 231 has been selected as responder cell line to evaluate the internalization capacity of functionalized EVs. MDA MB 231 were seeded in a 24-multiwell plate (25.000 cells/well) and cultured in DMEM supplemented with 10% Fetal Bovine Serum (FBS) (Gibco, Milan, Italy), 2 mM L-glutamine, and 50 mg/ml penicillin/streptomycin (complete medium) for 1 day, to let them adhere to the plastic support. Cells were then stimulated as follows: i) EVs+AZ647 (negative control, 100µL of sample + 100µL of phenol red-free 2X complete medium); ii) EVs+DBCO+AZ647 (Click-EVs, test group, 100µL of sample + 100µL of red-phenol red-free 2X complete medium). Untreated cells were used as additional negative control.

After 18h of incubation with either EVs+AZ647 or Click-EVs, cells were detached by Trypsin-EDTA (Gibco), rinsed in sterile PBS, centrifuged, resuspended in 300µL of PBS and analyzed by flow cytometry to evaluate the percentage of AF647 positive cells.

EV internalization was evaluated also by confocal microscopy. Cells were seeded on a glass slide fitting in a 24-multiwell plate (40.000 cells/well) and cultured in a complete medium for 1 day, to let them adhere to the glass support. Cells were then stimulated as above described for different time points (3h, 6h, 10h, 18h), rinsed 3 times with PBS, fixed with PFA 4% for 5 minutes at RT and permeabilized with 0,1% (v/v) Triton X-100 for 5 minutes at RT. Anti-HLA-I (supernatant of the clone W6.32, IgG2a) and anti-Rab5 (Abcam, Recombinant Anti-Rab5 antibody [EPR21801] - Early Endosome Marker (ab218624), 1:500) antibodies were incubated for 1h at RT to stain cell plasma membrane and early endosomes, respectively. Cells were then rinsed 3 times with PBS and incubated with the respective secondary antibodies (Goat anti-Mouse Alexa Fluor 546 and Goat anti-Rabbit Alexa Fluor 488) for 1h at RT and counterstained with DAPI for 20 minutes at RT. Samples were mounted on a glass slide with a water-based mounting medium (Permafluor, Thermo Fisher Scientific) and fluorescence imaged with the TCS SP2-AOBS laser scanning confocal microscope (Leica, Mannheim, Germany) through a plan apochromatic oil immersion objective 40x (1.4 NA). The 488 nm argon laser line was used for Rab5 fluorescence excitation, while the 546 nm and the 633 nm He-Ne lasers were utilized for excitation of HLA class I and Click-EVs fluorescence, respectively. DAPI was excited by a 405 nm diode laser. Leica software was used for image acquisition and analysis.

Statistical Analysis

The data were analyzed using unpaired t-test (to compare the percentage of AlexaFluor-647 positive EVs in negative control and Click-EVs), Ordinary one-way ANOVA (to evaluate the lipoprotein contaminants, to evaluate EV characterization by flow cytometry and super resolution microscopy, to compare the EV concentration measured by NTA, and to evaluate the EV internalization by flow cytometry) and Mann-Whitney Test (to evaluate the differences in terms of EV size by TEM and EV internalization at different time points by confocal microscopy). Data are presented as mean ± SD considering at least three independent replicates for each assay and analyzed by GraphPad Prism (Graph Pad Software, Inc.). For all analyses p < 0.05 was considered statistically significant. In all cases: ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05.

Competing interests

The authors declare that they have no competing interests.

Funding

“Programmi Operativi Nazionali” (PON), European Community, Grant number: ARS01_00693.

Author’s contributions

M.C.C. conceptualized, analyzed and interpreted all the data and was a major contributor in writing the manuscript and preparing the figures. S.B. performed the confocal microscopy analysis, reviewed the manuscript and prepared figures. N.R. performed the ELISA experiments. C.G. performed and analyzed all the NTA data and reviewed the manuscript. D.R. performed the flow cytometry analysis. M.C.G. performed the TEM analysis. K.C. analyzed the TEM data. C.Gr. performed and analyzed the super resolution microscopy analysis. B.B. analyzed the super resolution microscopy results. R.Q. conceptualized and reviewed the manuscript. R.T. conceptualized, analyzed and interpreted all the data and was a major contributor in writing the manuscript. All authors read and approved the final manuscript.

Acknowledgements

Not applicable.

Bayda S, Adeel M, Tuccinardi T, Cordani M, Rizzolio F. The History of Nanoscience and Nanotechnology: From Chemical–Physical Applications to Nanomedicine. Molecules. 2020; 25:112 .
Patra JK, Das G, Fernandes Fraceto L, Vangelie E, Campos R, Del Pilar Rodriguez-Torres M, et al. Nano based drug delivery systems: recent developments and future prospects. J Nanobiotechnology. 2018;16:71.
Li J, Yao M, Shao Y, Yao D. The application of bio-nanotechnology in tumor diagnosis and treatment: A view. Nanotechnol Rev. 2018;7:257-66.
Kinnear C, Moore TL, Rodriguez-Lorenzo L, Rothen-Rutishauser B, Petri-Fink A. Form Follows Function: Nanoparticle Shape and Its Implications for Nanomedicine. Chem Rev. 2017;117:11476-521.
Joseph TM, Kar Mahapatra D, Esmaeili A, Piszczyk Ł, Hasanin MS, Kattali M, et al. Nanoparticles: Taking a Unique Position in Medicine. Nanomaterials. 2023;13:574.
Li M, Fang F, Sun M, Zhang Y, Hu M, Zhang J. Extracellular vesicles as bioactive nanotherapeutics: An emerging paradigm for regenerative medicine. Theranostics. 2022;2022:4879-903.
Couch Y, Buzàs EI, Vizio D Di, Gho YS, Harrison P, Hill AF, et al. A brief history of nearly EV-erything – The rise and rise of extracellular vesicles. J Extracell Vesicles. 2021;10:e12144.
Yáñez-Mó M, Siljander PRM, Andreu Z, Zavec AB, Borràs FE, Buzas EI, et al. Biological properties of extracellular vesicles and their physiological functions. J Extracell Vesicles. 2015;4:1-60.
Raposo G, Stoorvogel W. Extracellular vesicles: Exosomes, microvesicles, and friends. Journal of Cell Biology. 2013;200:373-83.
Villata S, Canta M, Cauda V. Molecular Sciences EVs and Bioengineering: From Cellular Products to Engineered Nanomachines. Int J Mol Sci. 2020;21:6048.
Kwon S, Shin S, Do M, Oh BH, Song Y, Bui VD, et al. Engineering approaches for effective therapeutic applications based on extracellular vesicles. Journal of Controlled Release. 2021;330:15-30.
Xu L, Faruqu FN, Liam-or R, Abu Abed O, Li D, Venner K, et al. Design of experiment (DoE)-driven in vitro and in vivo uptake studies of exosomes for pancreatic cancer delivery enabled by copper-free click chemistry-based labelling. J Extracell Vesicles. 2020;9:1779458.
Smyth T, Petrova K, Payton NM, Persaud I, Redzic JS, Graner MW, et al. Surface functionalization of exosomes using click chemistry. Bioconjug Chem. 2014;25:1777-84.
Tian T, Zhang HX, He CP, Fan S, Zhu YL, Qi C, et al. Surface functionalized exosomes as targeted drug delivery vehicles for cerebral ischemia therapy. Biomaterials. 2018;150:137-49.
Karimi N, Cvjetkovic A, Jang SC, Crescitelli R, Hosseinpour Feizi MA, Nieuwland R, et al. Detailed analysis of the plasma extracellular vesicle proteome after separation from lipoproteins. Cellular and Molecular Life Sciences. 2018;75:2873-86.
Holcar M, Ferdin J, Sitar S, Tušek-Žnidarič M, Dolžan V, Plemenitaš A, et al. Enrichment of plasma extracellular vesicles for reliable quantification of their size and concentration for biomarker discovery. Scientific Reports. 2020;10:1–13.
Gorgun C, Reverberi D, Rotta G, Villa F, Quarto R, Tasso R. Isolation and Flow Cytometry Characterization of Extracellular-Vesicle Subpopulations Derived from Human Mesenchymal Stromal Cells. Curr Protoc Stem Cell Biol. 2019;48:e76.
Doyle LM, Wang MZ. Overview of Extracellular Vesicles, Their Origin, Composition, Purpose, and Methods for Exosome Isolation and Analysis. Cells. 2019;8:727.
Walker S, Busatto S, Pham A, Tian M, Suh A, Carson K, et al. Extracellular vesicle-based drug delivery systems for cancer treatment. Theranostics. 2019;9:8001-17.
Comprehensive Biotechnology - 2nd Edition, 2011. Elsevier.
Liu P, Chen G, Zhang J. A Review of Liposomes as a Drug Delivery System: Current Status of Approved Products, Regulatory Environments, and Future Perspectives. Molecules. 2022;27:1372.
van der Koog L, Gandek TB, Nagelkerke A. Liposomes and Extracellular Vesicles as Drug Delivery Systems: A Comparison of Composition, Pharmacokinetics, and Functionalization. Adv Healthc Mater. 2022;11:2100639.
Gaurav I, Thakur A, Iyaswamy A, Wang X, Chen X, Yang Z. Factors Affecting Extracellular Vesicles Based Drug Delivery Systems. Molecules. 2021;26:1544.
Elsharkasy OM, Nordin JZ, Hagey DW, de Jong OG, Schiffelers RM, Andaloussi S EL, et al. Extracellular vesicles as drug delivery systems: Why and how? Adv Drug Deliv Rev. 2020;159:332-43.
Zhu X, Badawi M, Pomeroy S, Sutaria DS, Xie Z, Baek A, et al. Comprehensive toxicity and immunogenicity studies reveal minimal effects in mice following sustained dosing of extracellular vesicles derived from HEK293T cells. J Extracell Vesicles. 2017;6:1324730.
Kooijmans SAA, Schiffelers RM, Zarovni N, Vago R. Modulation of tissue tropism and biological activity of exosomes and other extracellular vesicles: New nanotools for cancer treatment. Pharmacol Res. 2016;111:487-500.
Elliott RO, He M. Unlocking the Power of Exosomes for Crossing Biological Barriers in Drug Delivery. Pharmaceutics. 2021;13:1-20.
Alvarez-Erviti L, Seow Y, Yin H, Betts C, Lakhal S, Wood MJA. Delivery of siRNA to the mouse brain by systemic injection of targeted exosomes. Nature Biotechnology. 2011;29:341-5.
van Dongen HM, Masoumi N, Witwer KW, Pegtel DM. Extracellular Vesicles Exploit Viral Entry Routes for Cargo Delivery. Microbiology and Molecular Biology Reviews. 2016;80:369-86.
Liu Y, Li D, Liu Z, Zhou Y, Chu D, Li X, et al. Targeted exosome-mediated delivery of opioid receptor Mu siRNA for the treatment of morphine relapse. Scientific Reports. 2015;5:1-10.
Pedrioli G, Piovesana E, Vacchi E, Balbi C. Extracellular Vesicles as Promising Carriers in Drug Delivery: Considerations from a Cell Biologist’s Perspective. Biology (Basel). 2021;10:376.
Antimisiaris SG, Mourtas S, Marazioti A. Exosomes and Exosome-Inspired Vesicles for Targeted Drug Delivery. Pharmaceutics. 2018;10:218.
Garofalo M, Villa A, Brunialti E, Crescenti D, Dell’omo G, Kuryk L, et al. Cancer-derived EVs show tropism for tissues at early stage of neoplastic transformation. Nanotheranostics. 2021;5:1-7.
Villa A, Garofalo M, Crescenti D, Rizzi N, Brunialti E, Vingiani A, et al. Transplantation of autologous extracellular vesicles for cancer-specific targeting. Theranostics. 2021;11:2034-47.
Ruan H, Li Y, Wang C, Jiang Y, Han Y, Li Y, et al. Click chemistry extracellular vesicle/peptide/chemokine nanocarriers for treating central nervous system injuries. Acta Pharm Sin B. 2023;13:2202-18.
Alberro A, Iparraguirre L, Fernandes A, Otaegui D. Extracellular vesicles in blood: Sources, effects and applications. Int J Mol Sci. 2021; 22:8163.
Tsamchoe M, Petrillo S, Lazaris A, Metrakos P. Isolation of extracellular vesicles from human plasma samples: The importance of controls. Biotechnol J. 2023;18:2200575.
Reymond S, Gruaz L, Sanchez JC. Depletion of abundant plasma proteins for extracellular vesicle proteome characterization: benefits and pitfalls. Anal Bioanal Chem. 2023;415:3177-87.
Gupta S, Rawat S, Arora V, Kottarath SK, Dinda AK, Vaishnav PK, et al. An improvised one-step sucrose cushion ultracentrifugation method for exosome isolation from culture supernatants of mesenchymal stem cells. Stem Cell Res Ther. 2018; 9:180.
Jewett JC, Bertozzi CR. Cu-free click cycloaddition reactions in chemical biology. Chem Soc Rev. 2010;39:1272-9.
Gordon CG, MacKey JL, Jewett JC, Sletten EM, Houk KN, Bertozzi CR. Reactivity of biarylazacyclooctynones in copper-free click chemistry. J Am Chem Soc. 2012;134:9199-208.
Bernardin A, Cazet A, Guyon L, Delannoy P, Vinet F, Bonnaffé D, et al. Copper-Free Click Chemistry for Highly Luminescent Quantum Dot Conjugates: Application to in Vivo Metabolic Imaging. Bioconjug Chem. 2010;21:583-8.
Bird RE, Lemmel SA, Yu X, Zhou QA. Bioorthogonal Chemistry and Its Applications. Bioconjug Chem. 2021;32:2457-79.
Li Y, Fu H. Bioorthogonal Ligations and Cleavages in Chemical Biology. ChemistryOpen. 2020;9:835-53.
Bonsergent E, Grisard E, Buchrieser J, Schwartz O, Théry C, Lavieu G. Quantitative characterization of extracellular vesicle uptake and content delivery within mammalian cells. Nat Commun. 2021;12(1):1864.
Skovronova R, Grange C, Dimuccio V, Deregibus MC, Camussi G, Bussolati B. Surface marker expression in small and medium/large mesenchymal stromal cell-derived extracellular vesicles in naive or apoptotic condition using orthogonal techniques. Cells. 2021;10:2948.
Gorgun C, Ceresa D, Lesage R, Villa F, Reverberi D, Balbi C, et al. Dissecting the effects of preconditioning with inflammatory cytokines and hypoxia on the angiogenic potential of mesenchymal stromal cell (MSC)-derived soluble proteins and extracellular vesicles (EVs). Biomaterials. 2021; 269:120633.

No competing interests reported.

SupplementaryFigure1.pdf
Supplementary Figure 1. (a) Flow cytometry strategy used to characterize sUC-EVs. Green dots identify CFDA-SE positive events. EVs were stained with CFDA-SE at 4°C as "blank tube" (left panel) to define the appropriate dimensional gate when considering EVs stained with CFDA-SE at room temperature (right panel). (b) Western blot analysis of Alix, Flotillin-1, TSG-101 and Syntenin on sUC-EVs.

Download PDF

Version 1

posted

You are reading this latest preprint version

A standardized method to functionalize plasma-extracellular vesicles via copper-free click chemistry for targeted drug delivery strategies

Status:

Version 1

Abstract

Figures

Background

Results

Discussion

Conclusion

Methods

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1