Enrichment and Characterization Protocol for Plant‐Derived Nanovesicles from the Aquatic Plant Wolffia globosa

Abstract

This article presents a rigorous and reproducible protocol for the enrichment of plant‐derived nanovesicles (PDNVs) from the aquatic plant Wolffia globosa using differential ultracentrifugation combined with sucrose density gradient enrichment. The protocol is specifically optimized to address the technical challenges associated with PDNV enrichment from extremely small aquatic plants that require whole‐plant homogenization, a process that inherently increases contamination from intracellular organelles, soluble proteins, and chlorophyll. The workflow integrates gentle homogenization, sequential low‐ and medium‐speed centrifugation, sterile filtration, and an initial ultracentrifugation step to obtain crude PDNVs, followed by density‐based separation using a discontinuous sucrose gradient (8%–30% w/v). This combined size‐ and density‐based strategy enables effective enrichment of PDNVs while substantially reducing non‐vesicular contaminants. Quality control and characterization are performed in accordance with MISEV2023 recommendations using nanoparticle tracking analysis (NTA) and transmission electron microscopy (TEM). Gradient‐enriched PDNVs exhibit a predominantly nanoscale, homogeneous particle population with a mean diameter of approximately 105.3 nm and a particle concentration on the order of 10¹⁰ particles/ml. TEM analysis confirms well‐preserved membrane‐bound vesicles with characteristic round‐to‐cup‐shaped morphology and minimal background contamination, demonstrating a clear improvement over crude PDNV preparations. Quantitative normalization of PDNV yield to plant biomass and assessment of particle‐to‐protein ratios provide standardized metrics for evaluating preparation quality and reproducibility across independent preparations. Overall, this protocol provides a robust, scalable approach for obtaining enriched PDNVs prepared from W.globosa using commonly available laboratory equipment. The resulting PDNV preparations are suitable for downstream physicochemical characterization, molecular profiling, and functional studies, supporting their application in plant PDNV biology and emerging biomedical research. © 2026 Wiley Periodicals LLC.

Basic Protocol 1: Enrichment of plant‐derived nanovesicles (PDNVs) from the aquatic plant W. globosa by differential ultracentrifugation and sucrose density gradient

INTRODUCTION

Plant‐derived nanovesicles (PDNVs) are membrane‐bound nanoparticles secreted by cells into the extracellular environment, serving as fundamental mediators of intercellular communication across all domains of life. In plants, PDNVs transfer diverse biological cargo—including proteins, lipids, and nucleic acids (mRNA, microRNA, and small interfering RNA)—between cells and organisms, playing critical roles in development, stress responses, and pathogen defense (He et al., 2021). Plant‐derived PDNVs have also emerged as promising candidates for biomedical applications due to their biocompatibility, low immunogenicity, and ability to cross biological barriers, with potential uses in drug delivery, functional foods, and immunomodulation (Lian et al., 2022; Yang et al., 2023).

Wolffia globosa (family Lemnaceae, order Alismatales), commonly known as duckweed or Asian watermeal, is the smallest flowering plant, with a body diameter of approximately 0.5–1.5 mm (Bog & Sree, 2019). This aquatic angiosperm offers several unique advantages as a source of PDNVs, including exceptionally rapid growth (doubling time of 24–48 h), a simple morphology lacking true roots and conventional leaves, high nutritional content, and cultivation in aquatic environments that minimize soil‐derived contaminants. However, enrichment of PDNVs from W.globosa presents technical challenges that differ substantially from those encountered in larger terrestrial plants. Standard plant extracellular vesicle (EV) enrichment methods based on vacuum infiltration and recovery of apoplastic wash fluid—well established for species such as Arabidopsis thaliana and Nicotiana benthamiana (Eldahshoury et al., 2024; O'Leary et al., 2014)—are impractical for W. globosa's small, delicate structure, necessitating whole‐plant homogenization. This approach inevitably increases the risk of contamination from intracellular organelles (chloroplasts and mitochondria), soluble proteins (particularly Rubisco), and chlorophyll released by damaged photosynthate‐active cells. Current plant PDNV enrichment methods face additional challenges, including heterogeneous EV populations (TET8+, PEN1+, and EXPO vesicles; He et al., 2021), lack of standardized protocols and universal markers, co‐enrichment of protein aggregates during differential ultracentrifugation, and incomplete contamination removal (Cvjetkovic et al., 2014; Pinedo et al., 2021; Welsh et al., 2024).

This protocol addresses these challenges by combining differential ultracentrifugation with sucrose density gradient enrichment, exploiting both sedimentation rate (size‐dependent) and buoyant density differences between PDNVs (1.08–1.15 g/ml; 8%–30% sucrose) and contaminants (protein aggregates: >1.20 g/ml; >45% sucrose). Key optimizations include the following: (1) gentle homogenization balancing PDNV release against cellular disruption; (2) sequential low‐speed centrifugation (2,000 × g and 10,000 × g) removing intact cells and large organelles; (3) sterile filtration (0.22 µm) removing particulates; (4) optional repeated clarification spins (10,000 × g for 30 min) critically reducing chloroplast contamination; (5) initial ultracentrifugation (120,000 × g, 60 min) pelleting crude PDNVs; (6) sucrose gradient ultracentrifugation (120,000 × g, 90 min, swinging‐bucket SW 41 Ti rotor) achieving density‐based enrichment; and (7) systematic quality control by nanoparticle tracking analysis (NTA) and transmission electron microscopy (TEM). Comparison of crude PDNVs (post‐initial ultracentrifugation) and gradient‐enriched PDNVs by NTA shows substantial improvements: a narrower size distribution (predominantly <200 nm), reduced background contamination, minimal chlorophyll, and higher purity. Enriched PDNVs are suitable for physicochemical characterization [NTA for concentration and size distribution; TEM for morphology; dynamic light scattering (DLS) and zeta potential], comprehensive molecular profiling (proteomics, lipidomics, RNA sequencing, and metabolomics), and functional studies (cellular uptake, cargo delivery, and bioactivity assays). This protocol offers key advantages: It is optimized for small aquatic plants requiring whole‐plant processing; it provides superior purity compared with differential centrifugation alone; it uses standard laboratory equipment; it offers flexibility (crude PDNVs for moderate‐purity applications or full‐gradient enrichment for maximum purity); and it is scalable (50 g to >500 g starting material). Limitations include time requirements (1.5–2 days), unavoidable residual contamination from whole‐plant homogenization despite optimization, enrichment of mixed PDNV populations rather than specific subtypes, moderate yields (1–5 × 10⁹ particles/ml in 50–100 µl final volume from 50–100 g tissue), and potential PDNV aggregation during ultracentrifugation despite gentle handling. Iodixanol (OptiPrep™) density gradient centrifugation is recommended as an iso‐osmotic alternative to sucrose, as it preserves vesicle morphology at all density ranges and avoids the osmotic shrinkage artifact observed at sucrose concentrations above 30% (Greening et al., 2015; van Deun et al., 2017).

This article presents a Basic Protocol for the comprehensive enrichment of PDNVs from W. globosa using differential ultracentrifugation and sucrose density gradient enrichment. The protocol proceeds stepwise from fresh plant biomass as follows: washing and gentle homogenization; sequential centrifugation (2,000 × g, 10,000 × g) removing debris and organelles; sterile filtration (0.22 µm); optional clarification spins (10,000 × g, 30 min, repeated as needed) reducing chloroplast contamination; initial ultracentrifugation (120,000 × g, 60 min) pelleting crude PDNVs; resuspension and loading onto discontinuous sucrose gradients (8%, 15%, 30%, 45%, and 60% w/v in PBS); density gradient ultracentrifugation (120,000 × g, 90 min, SW 41 Ti rotor); collection of PDNV‐enriched fractions (8%–30% interfaces); optional washing removing residual sucrose; aliquoting and storage (4°C short‐term or −80°C long‐term); and confirmation by NTA, TEM, and protein quantification for marker enrichment/contaminant depletion. When properly executed, this protocol yields high‐purity small PDNVs with preserved membrane integrity, suitable for comprehensive characterization and functional studies, and addresses specific challenges in aquatic plant PDNV enrichment while maintaining the rigor and reproducibility standards outlined in the Minimal Information for Studies of Extracellular Vesicles (MISEV2023) guidelines (Pinedo et al., 2021; Welsh et al., 2024).

NOTE: All protocols involving animals must be reviewed and approved by the appropriate Animal Care and Use Committee and must follow regulations for the care and use of laboratory animals. Appropriate informed consent is necessary for obtaining and using human study material.

Basic Protocol 1. ENRICHMENT OF PLANT‐DERIVED NANOVESICLES (PDNVs) FROM THE AQUATIC PLANT W. globosa BY DIFFERENTIAL ULTRACENTRIFUGATION AND SUCROSE DENSITY GRADIENT

This protocol describes the enrichment of PDNVs from W.globosa using differential ultracentrifugation followed by sucrose density gradient enrichment, as shown in Fig. 1. Due to the unique morphology and small size (0.5–1.5 mm) of this aquatic plant, PDNVs are enriched from whole‐plant homogenates rather than from apoplastic wash fluid. The protocol combines gentle homogenization to release extracellular vesicles while minimizing intracellular contamination, followed by sequential differential centrifugation steps to remove cellular debris, organelles, and soluble contaminants (Fig. 1).

Figure 1.

Representative stages of plant‐derived nanovesicle enrichment from Wolffia globosa

Critical features of this protocol include sequential low‐speed centrifugation (2,000 × g and 10,000 × g) to remove intact cells, large debris, and organelles, followed by sterile filtration through 0.22‐µm membranes to remove remaining particulates. Optional clarification centrifugation steps (10,000 × g for 30 min, repeated if necessary) are included to further reduce chloroplast and protein contamination, particularly important when processing whole‐plant material from photosynthetically active tissue. Crude PDNVs are then pelleted by ultracentrifugation at 120,000 × g for 60 min, providing a preparation suitable for preliminary characterization or further enrichment.

The final enrichment by sucrose density gradient ultracentrifugation (120,000 × g for 90 min using a swinging‐bucket rotor, SW 41 Ti) separates PDNVs based on their buoyant density (1.08–1.15 g/ml), effectively removing protein aggregates, ribosomal complexes, and residual organellar fragments. This stepwise approach results in a superior‐quality preparation. Comparison of crude PDNVs (pelleted at 120,000 × g prior to gradient enrichment) with gradient‐enriched PDNVs by NTA demonstrates substantial improvements in particle size distribution, reduction in background contamination, and enhanced preparation purity. The enriched PDNVs exhibit minimal chlorophyll contamination, a narrow particle size distribution (predominantly <200 nm), and preserved vesicular morphology as confirmed by TEM.

When performed correctly, this protocol yields enriched PDNVs suitable for downstream physicochemical characterization (NTA, TEM, DLS, and zeta potential analysis), biochemical analysis (proteomics, lipidomics, and RNA profiling), and functional studies. The complete workflow from fresh plant material to characterized PDNVs is illustrated in Fig. 2, and representative NTA profiles comparing crude and enriched PDNV preparations demonstrate the effectiveness of density gradient enrichment in achieving superior sample quality. This optimized protocol addresses the specific challenges of enriching PDNVs from aquatic plants with minimal structural complexity while maintaining the rigor and reproducibility standards outlined in current guidelines for extracellular vesicle research (MISEV2023; Welsh et al., 2024; Pinedo et al., 2021).

Figure 2.

Workflow for enrichment of plant‐derived nanovesicles from Wolffia globosa

Materials

• Fresh, healthy W.globosa plants, free of visible contamination; healthy, bright green, 50–200 g fresh weight (Commercial fresh, Nature's Wolffia)

  • Use immediately after purchase; long‐term storage is not recommended for EV enrichment.

• Distilled or deionized water (ultrapure, 18.2 MΩ·cm; see recipe)

• Phosphate‐buffered saline, pH 7.4 ± 0.1; 280–300 mOsm/kg; sterile (PBS; Gibco, 10010023)

  • Store at 4°C. If unopened, it can be kept for 12–24 months; once opened, use within 6 months.

• Sucrose density gradient solutions, 8%, 15%, 30%, 45%, and 60% (w/v) in PBS (see recipe)

• Protease inhibitor cocktail, 100 × (Sigma‐Aldrich, P9599)

  • Broad‐spectrum, ready to use; keep stock solution at −20°C and working solution (1×) on ice.

• Analytical balance (accuracy ±0.01 g)

• High‐speed blender or homogenizer

• Cheesecloth or sterile gauze (four layers)

• 50‐ml conical centrifuge tubes (Falcon type, polypropylene)

• Refrigerated centrifuge capable of 2,000–10,000 × g

• Low‐protein‐binding syringe filters, 0.22 µm [polyethersulfone (PES) membrane]

• Sterile syringes, 10–60 ml

• Micropipettes with sterile tips

• Ultracentrifuge (Beckman Coulter Optima XPN‐100)

• Ultracentrifugation tubes, thin‐walled polypropylene, 13.2 ml (Beckman Coulter; cat. no. 331372)

• Ultracentrifugation rotor (swinging‐bucket rotor, type SW 41 Ti; Beckman Coulter)

• Microcentrifuge tubes, 1.5 or 2.0 ml (low protein binding)

• Malvern NS300 nanoparticle tracker (NanoSight‐NTA)

• Transmission electron microscope (TEM; Hitachi HT7700)

• Zetasizer Pro DLS analyzer (Malvern)

• 1Pre‐cool all equipment and reagents.

  Set all centrifuges and the ultracentrifuge to 4°C at least 30 min before use. Pre‐chill sterile‐filtered PBS and sucrose solutions at 4°C. Maintaining low temperature (4°C) throughout the procedure minimizes vesicle degradation, reduces proteolytic activity, and prevents aggregation.

• 2Weigh fresh W . globosa biomass (Fig. 3). Accurately weigh the wet biomass using an analytical balance and record the starting weight (in grams). Accurate recording of starting material weight is essential for calculating PDNV yield, ensuring reproducibility, and enabling comparison across experiments. A minimum of 50 g fresh weight is recommended.

Figure 3.

Fresh Wolffia globosa biomass used to enrich plant‐derived nanovesicles

• 3Wash plant material.

  Transfer the weighed biomass to a clean beaker. Rinse once with tap water to remove debris and then rinse twice with distilled or deionized water. After each rinse, drain water thoroughly by allowing plants to rest briefly on clean paper towels.

  Excessive mechanical agitation during washing should be avoided to minimize cell rupture and subsequent chlorophyll contamination. Gentle handling preserves cell integrity.

• 4Homogenize the plant material.

  Transfer the washed biomass to a pre‐chilled blender or homogenizer. Add a small amount of ice‐cold distilled or deionized water (approximately 1:1 w/v, plant weight to water volume). Homogenize using short pulses (3–5 s per pulse, 3–4 pulses total) at low‐to‐medium speed until a uniform slurry is obtained.

  Over‐homogenization markedly increases contamination by intracellular organelles (chloroplasts and mitochondria) and soluble metabolites. The optimal homogenization time should be determined empirically for each blender model, with other parameters held constant. The resulting slurry should appear uniformly green, with no visible leaf fragments.

• 5Perform coarse filtration.

  Pass the homogenate through four layers of sterile cheesecloth or gauze into pre‐chilled 50‐ml conical tubes. Gently squeeze the cheesecloth to recover filtrate without excessive force.

  This step removes large plant debris (cell walls and tissue fragments) and reduces clogging in subsequent centrifugation steps. Avoid excessive squeezing, which may rupture cells.

• 6Perform the first centrifugation to remove large debris.

  Centrifuge the filtrate at 2000 × g for 30 min at 4°C using a fixed‐angle rotor. Carefully collect the supernatant without disturbing the pellet. Transfer supernatant to fresh 50‐ml tubes.

  This low‐speed centrifugation removes intact cells, large cell debris, and aggregated material.

• 7Perform the second centrifugation to remove residual debris.

  Centrifuge the collected supernatant at 10,000 × g for 60 min at 4°C. Carefully transfer the supernatant to fresh tubes, avoiding disturbance of the pellet.

  This step further reduces large particles, cellular fragments, and organelles (e.g., broken chloroplasts and mitochondria). The supernatant should appear clearer than after step 6.

• 8Perform sterile filtration. Filter the supernatant through a sterile 0.22‐µm low‐protein‐binding syringe filter (PES membrane) into a clean container. Apply gentle, steady pressure to avoid filter rupture and vesicle loss. Filtration removes large particles and aggregates that may interfere with ultracentrifugation. Use low‐protein‐binding filters to minimize PDNV adhesion. Pre‐wet filters with PBS if processing large volumes. This filtration step is employed primarily to reduce bacterial contamination and large cellular debris. Particles slightly larger than 220 nm may still be present in the final preparation, owing to the inherent measurement uncertainty of NTA and the physical heterogeneity of plant‐derived nanovesicle populations (Théry et al., 2018; Liu et al., 2020). This step is therefore not intended as a strict size‐exclusion cut‐off.
• 9Perform the clarification spin (optional but strongly recommended). Centrifuge the filtered supernatant at 10,000 × g for 30 min at 4°C. Collect the supernatant. This prolonged clarification step improves the removal of remaining contaminants (protein aggregates and small organelle fragments) prior to ultracentrifugation, resulting in higher PDNV purity.
• 10Perform the optional chlorophyll reduction step.

  If the supernatant retains visible green coloration, perform an additional centrifugation at 10,000 × g for 30 min at 4°C. Retain only the clarified supernatant. Discard samples with a persistent dark green color.

  Green coloration indicates chloroplast contamination from broken cells. Repeat washing and homogenization with fresh tissue if contamination persists. This step is critical in preparations intended for proteomic or RNA analysis, where chloroplast‐derived material may confound results.

• 11Prepare ultracentrifugation tubes.

  Transfer the clarified supernatant into ultracentrifugation tubes appropriate for the selected rotor (thin‐wall polypropylene tubes for SW 41 Ti rotor). Fill tubes to within 2–3 mm of the top to avoid tube collapse during centrifugation.

  Underfilling or overfilling tubes may cause tube deformation or leakage during high‐speed centrifugation. Avoid introducing air bubbles, which interfere with rotor balance and may cause tube failure.

• 12Balance ultracentrifugation tubes.

  Balance tubes in pairs to within ±0.01 g using sterile‐filtered PBS. Record tube weights if required by institutional safety protocols.

  Proper balancing is critical for ultracentrifuge safety, rotor longevity, and reproducibility. Imbalanced tubes can cause rotor damage, sample loss, and inconsistent results.

• 13Pellet crude PDNVs. Ultracentrifuge at 120,000 × g for 60 min at 4°C using a swinging‐bucket rotor. Set acceleration to “slow” (rate 3–4) and deceleration to “slow” or “no brake” (rate 0–2) to minimize PDNV aggregation. The PDNV pellet is typically invisible or faintly translucent. Mark the expected pellet position on the tube exterior before centrifugation (bottom of the tube for swinging‐bucket rotor). Gentle acceleration/deceleration reduces mechanical stress on PDNVs and minimizes aggregation (Cvjetkovic et al., 2014).
• 14Remove supernatant.

  Using a sterile pipette, carefully aspirate and discard the supernatant, starting from the top of the tube. Leave approximately 200–500 µl of buffer above the pellet to avoid disturbing it. Aspirate slowly from the side opposite the expected pellet location. The final buffer can be removed with a micropipette after visually confirming the pellet's position.

• 15Resuspend the crude PDNV pellet. Add 2 ml of ice‐cold, sterile‐filtered PBS to each tube and gently resuspend the PDNV pellet by pipetting (10–15 cycles) using a 1‐ml pipette tip. Avoid vortexing. If multiple tubes are processed, pool the resuspended material. Gentle resuspension minimizes PDNV aggregation and preserves vesicle integrity. This suspension represents the crude PDNV fraction. If resuspension is difficult, incubate the pellet on ice for 10–15 min before pipetting. NTA of crude PDNVs (Fig. 4A, 4B) shows a heterogeneous particle population with a broad size distribution, consistent with a mixed vesicle population obtained after differential ultracentrifugation.

Figure 4.

Nanoparticle tracking analysis–based characterization of crude plant‐derived nanovesicles

• 16Prepare a sucrose step gradient.

  Using a serological pipette or gradient maker, prepare a discontinuous sucrose gradient in clean ultracentrifugation tubes (Ultra‐Clear™ tubes for SW 41 Ti rotor). Layer 2.0 ml of each sucrose solution sequentially from bottom to top in the following order: 60%, 45%, 30%, 15%, and 8% (w/v in PBS). Add each layer slowly along the tube wall to maintain sharp interfaces.

  Accurate layering is critical for effective density‐based separation. Tilt tubes at a 45° angle and allow each layer to flow slowly down the wall. Visible interfaces should be maintained between layers. Pre‐chilled solutions prevent mixing due to temperature‐induced density change.

• 17Overlay crude PDNV suspension. Carefully overlay 2 ml of the resuspended crude PDNV suspension onto the top of the gradient (above the 8% sucrose layer). Use a pipette and add slowly to avoid disturbing the gradient. The crude PDNV suspension should form a distinct layer on top of the 8% sucrose. If the gradient is disturbed, discard and prepare a fresh gradient.
• 18Perform density gradient ultracentrifugation. Ultracentrifuge the gradient at 120,000 × g for 90 min (or up to 16 h for sharper separation) at 4°C using a swinging‐bucket rotor. Use slow acceleration (rate 3) and minimal deceleration (rate 1–2, or no brake). This step separates PDNVs from protein aggregates, soluble proteins, and other contaminants based on buoyant density. Plant PDNVs typically equilibrate at densities of 1.08–1.15 g/ml, corresponding to the 8%–30% sucrose interface (Welsh et al., 2024). Extended centrifugation (16 h) may improve purity but is not essential.
• 19Collect PDNV‐enriched fractions. After centrifugation, carefully remove and discard approximately the top 2 ml of the gradient. Using a sterile pipette, collect 2.0 ml fractions corresponding to the interfaces between 8%–15%, 15%–30%, and 30%–45% sucrose layers. Transfer each fraction to labeled tubes. PDNV‐enriched fractions are typically found at the 8%–15% and 15%–30% interfaces. Collect fractions slowly to avoid cross‐contamination between layers. The 30%–45% fraction may contain larger vesicles or contaminants and should be analyzed separately.
• 20Pool PDNV‐containing fractions. Combine the collected 8%–15% and 15%–30% fractions (approximately 4–6 ml total) into a single container. Record the total pooled volume. Pooling increases the final PDNV yield. If analyzing individual fractions separately (e.g., for subpopulation characterization), skip this step.
• 21Add protease inhibitors (optional). If samples are intended for protein analysis (western blotting or proteomics), add a protease inhibitor cocktail to the pooled fractions according to the manufacturer's instructions (typically a 1:100 dilution of a 100 × stock). Mix gently by inversion. Protease inhibitors prevent degradation of the PDNV surface and cargo proteins during subsequent handling. This step is not necessary for RNA extraction or lipid analysis.
• 22Wash PDNVs to remove sucrose (optional but recommended).

  Dilute the pooled PDNV fractions 1:5 with sterile PBS (e.g., 6 ml PDNV suspension + 24 ml PBS = 30 ml total). Transfer to ultracentrifugation tubes and centrifuge at 120,000 × g for 60 min at 4°C. Remove the supernatant and resuspend the pellet in 50–100 µl of PBS.

  This washing step removes residual sucrose, which may interfere with downstream applications (e.g., protein quantification or NTA). For functional studies, sucrose removal is strongly recommended.

• 23Aliquot samples. Divide the enriched PDNV suspension into appropriately sized aliquots (e.g., 10–20 µl per tube for single‐use aliquots) in sterile, low‐adhesion microcentrifuge tubes. Label tubes clearly with sample ID, date, and storage conditions. Aliquoting minimizes repeated freeze–thaw cycles, which can cause PDNV aggregation, membrane damage, and loss of biological activity. Use low‐binding tubes to minimize PDNV adhesion.
• 24Store enriched PDNVs. Store aliquots at 4°C for short‐term use (up to 1 week) or at −80°C for long‐term storage (up to 6 months). For ultra‐long‐term storage, consider lyophilization or storage in cryopreservation medium (e.g., 10% DMSO or trehalose). Record storage conditions, date, and freeze–thaw history in laboratory records. Avoid repeated freeze–thaw cycles (maximum 2–3 cycles). For critical experiments, use fresh PDNV preparations. PDNVs stored at −80°C maintain structural integrity and bioactivity for several months, but activity should be verified before use.
• 25Confirm PDNV enrichment and proceed with downstream analysis.

  Verify PDNV size, concentration, and morphology using NTA and TEM. If properly executed, this protocol yields predominantly nanoscale vesicles with a narrow size distribution and particle concentrations of 1–8 × 10¹⁰ particles/ml (Fig. 5A, 5B). TEM analysis confirms well‐preserved cup‐shaped vesicular structures with diameters of 50–200 nm and minimal background contamination (Fig. 4A–4D). These results demonstrate significant improvement over crude PDNVs and confirm successful enrichment of homogeneous PDNV populations suitable for downstream characterization, functional assays, and mechanistic studies.

Figure 5.

Nanoparticle tracking analysis (NTA)–based characterization of enriched plant‐derived nanovesicles. Fractions from the 8%–30% sucrose interface (buoyant density 1.10–1.18 g/ml) were selected for NTA analysis shown here. Fractions above 1.20 g/ml were excluded due to potential osmotic deformation artifacts at high sucrose concentrations (Greening et al., 2015; van Deun et al., 2017).

REAGENTS AND SOLUTIONS

Use ultrapure deionized water for all preparations.

Deionized water

Using a laboratory purification system (e.g., Milli‐Q, PURELAB, or Arium) to produce deionized water with a purity of 18.2 MΩ·cm, <10 ppb TOC. Store at room temperature and prepare fresh every day.

Sucrose solutions (8%, 15%, 30%, 45%, and 60% w/v)

Dissolve sucrose in PBS to obtain a final volume of 100 ml (see amounts below). The densities should be 1.03‐1.29 g/ml. The solutions can be stored at 4°C for a week, but using freshly prepared solutions is recommended. Table 1 presents the solution specifications.

Table 1.

Sucrose Amounts for 100 ml Final Volume

Concentration	Sucrose Amount	Density
8% w/v	8.0 g	∼1.03 g/ml
15% w/v	15.0 g	∼1.06 g/ml
30% w/v	30.0 g	∼1.13 g/ml
45% w/v	45.0 g	∼1.20 g/ml
60% w/v	60.0 g	∼1.29 g/ml

Note: Sucrose solutions were prepared as % w/v (grams of sucrose dissolved and brought to a final volume of 100 ml with PBS). The density values presented in Table 1 are approximate densities estimated from published sucrose density reference tables at 20°C. These values are intended as theoretical references corresponding to the nominal sucrose concentrations and were not determined experimentally for the prepared solutions. Because the reference tables report sucrose concentrations as % w/w, density values were estimated by interpolation of published density data for sucrose solutions at the corresponding % w/w concentrations equivalent to the nominal % w/v values. Consequently, minor discrepancies between the reported and actual densities of the prepared solutions may occur, particularly at higher sucrose concentrations. The values in Table 1 should therefore be regarded as theoretical estimates rather than experimentally determined densities.

COMMENTARY

Background Information

Plant‐derived nanovesicles (PDNVs) are membrane‐bound nanostructures secreted by plant cells that mediate intercellular and interorganismal communication through the transfer of proteins, lipids, metabolites, and nucleic acids, including mRNA and small RNAs. Growing evidence demonstrates that plant PDNVs play critical roles in development, stress responses, and plant–pathogen interactions, and they have recently attracted substantial interest in biomedical and biotechnological applications due to their biocompatibility, low immunogenicity, and capacity to deliver bioactive cargo across biological barriers. Compared with mammalian EVs, plant PDNVs present additional methodological challenges, including the absence of universally accepted markers, intrinsic heterogeneity of vesicle populations, and the frequent co‐enrichment of non‐vesicular contaminants such as protein aggregates and ribonucleoprotein complexes. These issues have led to calls for increased rigor, standardized workflows, and transparent reporting in plant PDNV research, as reflected in recent community guidelines and consensus statements (He et al., 2021; Pinedo et al., 2021; Welsh et al., 2024).

Wolffia globosa, the smallest flowering plant known, represents a unique and promising source of plant‐derived PDNVs due to its exceptionally rapid growth, aquatic cultivation, and simple morphology lacking differentiated roots and leaves. However, these same features impose distinct technical constraints on PDNV enrichment. Established plant PDNV protocols often rely on vacuum infiltration and recovery of apoplastic wash fluid, an approach that is not feasible for W. globosa because of its minute size and fragile structure. Consequently, PDNV enrichment from this species requires whole‐plant homogenization, which inherently increases the risk of intracellular contamination. Mechanical disruption can release chloroplasts, mitochondria, and abundant soluble proteins such as ribulose‐1,5‐bisphosphate carboxylase/oxygenase (Rubisco) and chlorophyll, all of which can confound downstream physicochemical and molecular analyses if not effectively removed (Cvjetkovic et al., 2014; Pinedo et al., 2021).

To address these challenges, rigorous separation strategies that combine size‐ and density‐based principles are required. Differential ultracentrifugation enables enrichment of vesicle‐containing fractions based on sedimentation behavior, while subsequent sucrose density gradient ultracentrifugation exploits differences in buoyant density to separate PDNVs (typically 1.08–1.15 g/ml for small PDNVs) from protein aggregates and organellar remnants, which equilibrate at higher densities. This combined approach has been widely adopted as a gold‐standard enrichment strategy in both mammalian and plant PDNV research and is strongly aligned with the MISEV2023 recommendations, which emphasize contamination control, transparent reporting, and method‐dependent interpretation of EV preparations (Welsh et al., 2024). Within this framework, careful optimization of homogenization intensity, clarification steps, and gradient fraction selection is essential to obtain reproducible, high‐purity PDNV preparations from W. globosa suitable for robust downstream characterization and functional studies.

Quantification of PDNV yield and purity from Wolffia globosa

PDNV yield from W. globosa was quantified by NTA and normalized to plant biomass. Particle yield was expressed as particles per gram of fresh plant material (particles/g), calculated from the average particle concentration multiplied by the final suspension volume and divided by the fresh weight of plant tissue used for enrichment.

The first enrichment yielded 6.51 × 10⁸ particles/g, whereas the second enrichment yielded 3.43 × 10⁹ particles/g, corresponding to an average PDNV yield of 2.04 × 10⁹ particles/g across independent preparations. Particle concentration measurements demonstrated good analytical precision within each run. For the second preparation, NTA reported 9.38 × 10¹⁰ ± 7.85 × 10⁹ particles/ml, corresponding to a coefficient of variation (CV) of 8.4%, which is within the expected technical variability range for NTA measurements. PDNV purity was evaluated using the particle‐to‐protein ratio (particles per µg protein), a commonly applied indicator of protein contamination. Based on a mean protein concentration of 12.33 µg/ml (range 8–17 µg/ml), the calculated purity was 7.61 × 10⁹ particles/µg protein, with a conservative range of 5.1 × 10⁹ to 1.27 × 10¹⁰ particles/µg protein.

Together, these results indicate reproducible particle quantification with acceptable technical precision and demonstrate that the enriched vesicles exhibit moderate‐to‐high purity, suitable for downstream functional and molecular analyses. These reporting practices are consistent with current recommendations for extracellular vesicle characterization.

Sucrose density gradient ultracentrifugation conditions

Sucrose density gradient ultracentrifugation is a critical step for improving the purity of PDNVs enriched from W. globosa, as it enables separation based on buoyant density rather than sedimentation alone. This approach is particularly important for plant‐derived PDNVs obtained through whole‐plant homogenization, which introduces abundant soluble proteins, pigments, and organellar debris. Small PDNVs typically equilibrate at densities of approximately 1.08–1.15 g/ml, corresponding to the 8%–30% (w/v) sucrose interfaces, whereas protein aggregates and membrane fragments localize to higher‐density fractions. The use of a discontinuous sucrose gradient therefore enhances selective enrichment of PDNV‐containing fractions and substantially reduces non‐vesicular contamination (Greening et al., 2015; Kowal et al., 2016; Rutter & Innes, 2017).

Proper gradient preparation and standardized fraction collection are essential for reliable outcomes. Sucrose solutions should be freshly prepared, sterile‐filtered, and carefully layered to maintain sharp interfaces. PDNV‐enriched fractions are typically recovered from the 8%–15% and 15%–30% interfaces, depending on the intended downstream application. Because residual sucrose can interfere with subsequent analyses, an additional washing step is commonly performed prior to PDNV characterization. Clear reporting of gradient composition, centrifugation parameters, and fraction selection criteria is essential for reproducibility and aligns with the rigor and transparency emphasized in the MISEV2018 and MISEV2023 guidelines (Théry et al., 2018; Welsh et al., 2024).

Although sucrose density gradient ultracentrifugation is widely used for PDNV enrichment, researchers should be aware that high sucrose concentrations (above 30% w/v) can cause osmotic shrinkage of vesicles, producing cup‐shaped or collapsed morphologies in TEM images from high‐density fractions (>1.20 g/ml). Fractions above this density range were therefore excluded from NTA and TEM analysis in this study. As an iso‐osmotic alternative, iodixanol (OptiPrep™) density gradient centrifugation is recommended for applications where preservation of native vesicle morphology is critical, as iodixanol maintains constant osmolarity across all density ranges, eliminates osmotic deformation artifacts, and can provide sharper band separation (Greening et al., 2015; van Deun et al., 2017).

Troubleshooting Table

Table 2 presents some common problems encountered when performing this protocol and possible solutions.

Table 2.

Troubleshooting Guide for the Enrichment of Plant‐derived Nanovesicles (PDNVs) from Wolffia Globosa

Problem	Possible Cause	Solution
Low PDNV yield	Poor quality or stressed W. globosa starting material	Use fresh, healthy plants with uniform green coloration. Avoid senescent or damaged biomass. Process samples immediately after harvesting.
Low PDNV yield	Insufficient homogenization	Increase homogenization time slightly while avoiding over‐homogenization. Optimize conditions for the specific blender or homogenizer used.
Excessive green coloration in PDNV preparation	Over‐homogenization leading to chloroplast rupture	Reduce homogenization intensity and duration. Add or repeat clarification centrifugation steps (e.g., 10,000 × g for 30 min).
High protein background	Co‐enrichment of soluble plant proteins	Include an additional PBS wash after ultracentrifugation if appropriate.
Broad or multimodal particle size distribution by NTA	Presence of non‐vesicular nanoparticles or aggregates	Ensure all buffers are freshly prepared and passed through a 0.22‐µm filter. Avoid vigorous pipetting or vortexing of PDNV suspensions.
Apparent PDNVs detected in buffer‐only controls	Particulate contamination from buffers or plasticware	Filter all buffers immediately before using. Use low‐bind tubes and run buffer‐only controls to identify background signals.
Pellet not visible after ultracentrifugation	Small pellet size or incorrect tube orientation	Mark the expected pellet side before centrifugation. Aspirate supernatant slowly from the opposite side of the tube.
PDNV aggregation after ultracentrifugation	High centrifugal force and/or harsh deceleration	Use gentle acceleration/deceleration settings. Consider resuspending pellets slowly by pipetting rather than vortexing.
Poor reproducibility between experiments	Inconsistent centrifugation conditions or rotor differences	Report and maintain centrifugal force (× g), rotor type, and acceleration/deceleration settings consistently across experiments.
Reduced PDNV integrity after storage	Repeated freeze–thaw cycles	Aliquot PDNV preparations immediately after enrichment and avoid multiple freeze–thaw events. Store long‐term samples at −80°C.

Understanding Results

Successful execution of the protocols described here yields PDNV preparations from W. globosa that can be evaluated using complementary physical and morphological analyses. Representative data generated in our laboratory—including NTA size distribution profiles of crude and sucrose gradient–enriched PDNVs (Fig. 4 and Fig. 5), corresponding NTA video clips illustrating particle motion and tracking behavior in crude and enriched preparations (Video 1 and Video 2), and TEM images confirming vesicular morphology and membrane integrity (Fig. 4)—serve as practical benchmarks for users to assess whether the protocols have been accurately reproduced.

Video 1.

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Representative nanoparticle tracking analysis video clips of crude plant‐derived nanovesicles isolated from Wolffia globosa.

Video 2.

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Representative nanoparticle tracking analysis video clips of sucrose gradient–enriched plant‐derived nanovesicles isolated from Wolffia globosa.

Crude PDNVs

Following differential ultracentrifugation, crude PDNV preparations are expected to exhibit a heterogeneous particle population with a broad size distribution. NTA analysis typically reveals particles spanning a wide nanoscale range, with a mean diameter of approximately 180.8 nm and a modal size of 160 nm (Fig. 4A, 4B). This broad distribution reflects the mixed vesicle population and residual non‐vesicular components commonly present in crude fractions obtained from whole‐plant homogenates.

Sucrose gradient–enriched PDNVs

In contrast, PDNVs enriched from the sucrose density gradient are expected to display markedly improved uniformity and purity. NTA analysis of sucrose gradient–enriched PDNVs demonstrates a predominantly nanoscale particle population with a narrow size distribution, characterized by a mean diameter of 105.3 nm and a modal size of 100 nm (Fig. 5A, 5B). Particle concentrations of approximately 10¹⁰ particles/ml are typically obtained and are consistent across technical replicates. TEM analysis of these enriched fractions reveals well‐preserved, membrane‐bound vesicles with characteristic round‐to‐cup‐shaped morphology and minimal background contamination (Fig. 6A–6D), supporting effective removal of non‐vesicular material during density gradient separation.

Figure 6.

Transmission electron microscopy (TEM) of enriched plant‐derived nanovesicles enriched from Wolffia globosa following sucrose density gradient ultracentrifugation. TEM images represent vesicles collected from the 8%–30% sucrose gradient interface (density 1.10–1.18 g/ml). Fractions above 1.20 g/ml were excluded due to potential osmotic deformation at high sucrose concentrations. Iodixanol (OptiPrep™) gradient centrifugation is recommended as an iso‐osmotic alternative that eliminates osmotic deformation artifacts (Greening et al., 2015; van Deun et al., 2017).

Overall interpretation

Collectively, these results illustrate the progressive improvement in the quality of PDNV preparation achieved through sucrose density gradient enrichment. Comparison of crude and enriched fractions highlights the expected trade‐off between heterogeneity and purity inherent to plant PDNV enrichment workflows. When the protocol is performed correctly, users should observe a clear transition from a broad, heterogeneous crude PDNV population to a relatively homogeneous population of small PDNVs with preserved morphology and reduced background. Such preparations are suitable for downstream physicochemical characterization, functional assays, and mechanistic studies that require high confidence in PDNV‐specific effects.

Time Considerations

The total time required to complete the protocols described in this article depends on the level of enrichment selected and the availability of equipment. Approximate time estimates for each protocol are provided below to assist with experimental planning. Enrichment of PDNVs from W. globosa using sucrose density gradient ultracentrifugation is typically completed over 1.5–2 days, depending on sample number, centrifuge availability, and operator experience.

On Day 1, preparation of plant material, including washing, homogenization, coarse filtration, and sequential low‐ and medium‐speed centrifugation steps, requires approximately 4–6 h. This is followed by an initial ultracentrifugation step to concentrate PDNV‐containing material, which requires ∼1 h, in addition to time for tube balancing and sample handling. Preparation of sucrose solutions and assembly of the step density gradient typically require an additional 1–2 h, particularly when multiple gradients are prepared in parallel. The density gradient ultracentrifugation step requires ∼1 h. Depending on laboratory scheduling, this step can be performed at the end of Day 1.

On Day 2, careful fraction collection from the sucrose gradient, pooling of PDNV‐enriched fractions (typically from the 8%–30% interfaces), and optional addition of protease inhibitors require approximately 1–2 h. Aliquoting and storage of enriched PDNVs require an additional 15–30 min.

Overall, most hands‐on time is concentrated in sample preparation and fraction collection, whereas centrifugation steps constitute the majority of total elapsed time. Preparing reagents in advance and coordinating centrifugation schedules can substantially improve workflow efficiency and reduce total processing time.

Author Contributions

Arunya Jiraviriyakul: Conceptualization; methodology; supervision; writing—review and editing; writing—original draft; project administration; investigation; funding acquisition; formal analysis; resources. Manatsanan Piamano: Investigation; validation; formal analysis; data curation; writing—original draft. Korawut Pimpa: Investigation; writing—original draft; validation; formal analysis; data curation. Nonthawan Sangprasert: Investigation; writing—original draft; validation; formal analysis; data curation. Manasanan Wongchaiya: Investigation; writing—original draft; validation; formal analysis; data curation. Sachittha Arin: Investigation; validation; formal analysis; data curation; writing—original draft. Chatchai Nensat: Visualization; writing—review and editing. Worawat Songjang: Writing—review and editing; visualization.

Conflict of Interest

The authors declare no conflict of interest.

Jiraviriyakul, A. , Piamano, M. , Pimpa, K. , Sangprasert, N. , Wongchaiya, M. , Arin, S. , Nensat, C. , & Songjang, W. (2026). Enrichment and characterization protocol for plant‐derived nanovesicles from the aquatic plant Wolffia globosa . Current Protocols, 6, e70427. doi: 10.1002/cpz1.70427

Data Availability Statement

Original files for TEM, NTA, and protein concentration data are available upon request.