Plant Extracellular Vesicles and Nanovesicles: Focus on Secondary Metabolites, Proteins and Lipids with Perspectives on Their Potential and Sources

Abstract

While human extracellular vesicles (EVs) have attracted a big deal of interest and have been extensively characterized over the last years, plant-derived EVs and nanovesicles have earned less attention and have remained poorly investigated. Although a series of investigations already revealed promising beneficial health effects and drug delivery properties, adequate (pre)clinical studies are rare. This fact might be caused by a lack of sources with appropriate qualities. Our study introduces plant cell suspension culture as a new and well controllable source for plant EVs. Plant cells, cultured in vitro, release EVs into the growth medium which could be harvested for pharmaceutical applications. In this investigation we characterized EVs and nanovesicles from distinct sources. Our findings regarding secondary metabolites indicate that these might not be packaged into EVs in an active manner but enriched in the membrane when lipophilic enough, since apparently lipophilic compounds were associated with nanovesicles while more hydrophilic structures were not consistently found. In addition, protein identification revealed a possible explanation for the mechanism of EV cell wall passage in plants, since cell wall hydrolases like 1,3-β-glucosidases, pectinesterases, polygalacturonases, β-galactosidases and β-xylosidase/α-L-arabinofuranosidase 2-like are present in plant EVs and nanovesicles which might facilitate cell wall transition. Further on, the identified proteins indicate that plant cells secrete EVs using similar mechanisms as animal cells to release exosomes and microvesicles.

Keywords: exosome-like nanoparticles; extracellular vesicles; in vitro plant cell culture; lipids; metabolomics; plant nanovesicles; proteomics.

Figure A1

Product ion spectrum of aconitine standard.

Figure A2

Product ion spectrum of putative mesaconitine.

Figure A3

Product ion spectrum of putative hypaconitine.

Figure 1

Scheme of current issues in plant nanovesicle research. The illustrated mechanisms of extracellular vesicle (EV) formation can probably be transferred from mammals [6,7] to eukaryotic cells in general. While evidence for the release of exosomes has been reported, it remains unknown whether plant cells also secrete plasma membrane derived microvesicles. Apparently, EVs from plants exert inhibitory effects against pathogenic microorganisms [8,9,10,11] and spread information to other cells, although it has not yet been conclusively elucidated how EVs pass through cell walls. Like other EVs, plant nanovesicles carry distinct nucleic acids. Proteomic analysis of plant nanovesicles revealed the repeated identification of cytosolic proteins (e.g., heat shock proteins), as well as membrane-associated proteins. It remains puzzling, whether cell wall hydrolases are membrane associated and if they enable cell wall passage. Also to be clarified are the questions why and to which extent lipid compositions can vary, as well as if secondary metabolites are packaged into nanovesicles. CHMP: charged multivesicular body protein; DUB: deubiquitinases; ESCRT: endosomal-sorting complex required for transport; EV: extracellular vesicle; MVB: multivesicular body; v-SNARE: vesicular soluble N-ethylmaleimide-sensitive-factor attachment receptor.

Figure 2

Transmission electron microscopy images of (a) extracellular vesicles (EVs) from Craterostigma plantagineum Hochst. cell culture medium. (b) Nanovesicles from C. plantagineum cells. (c) Nanovesicles from dried tubers of Aconitum napellus L. (d) EVs from Nicotiana tabacum L. cell culture medium.

Figure 3

Influence of pH on secondary metabolite lipophilicity by the example of a curcuminoid HPTLC-scan. Nanovesicles (NVs) from Curcumae zanthorrhizae rhizoma, Curcuma zanthorrhiza Roxb. (CZ) were prepared in acidic vesicle isolation buffer (VIB), as well as in alkaline Tris-buffered saline (TBS). While curcuminoids (curcuminoid mixture contained curcumin, demethoxycurcumin and bisdemethoxycurcumin *, curcumin with the highest Rf, bisdemethoxycurcumin with the lowest) dissolved in the organic extractant, when the vesicles were prepared in acidic VIB, vesicles in TBS had to be acidified, since otherwise acidic curcuminoids would not dissolve within the solvent. These results implicate that secondary metabolites likewise cannot pass membranes when the buffer is chosen to generate charges in the structures of interest. Rf: Retention factor S: Supernatant of high speed centrifugation before (I) and after (II) washing the nanovesicles. * The absorbance spectrum of the band corresponding to bisdemethoxycurcumin in lane “CZ NVs in VIB” showed severe differences to curcuminoid spectra and was possibly caused by caffeic acid derivatives.

Figure 4

HPTLC-scans of nanovesicles (NVs) isolated from dried Aconiti tuber, Aconitum napellus L. (AN), extracellular vesicles (EVs) from Nicotiana tabacum L. (NT) leaves’ apoplastic fluid and NVs from and Uvae-ursi folium, Arctostaphylos uva-ursi (L.) Spreng. (AU). While the aqueous supernatants of high-speed centrifugation contained characteristic secondary metabolites prior to the washing step (S I) in all three samples, those structures have not been found in the investigated vesicle samples. Nonetheless, the metabolites present in S I have been effectively removed from the vesicles during the washing step, as the chromatograms of the second supernatants (S II) show. Interestingly, AN S I showed no aconitine band but instead two bands giving a positive reaction with Dragendorff’s reagent and similar absorbance spectra like aconitine. LC-MS analysis of reextracted silica identified aconitine to be present in traces, besides larger amounts of hypaconitine and mesaconitine. Rf: Retention factor.

Figure 5

(a) HPTLC-chromatogram at 366 nm of Craterostigma plantagineum Hochst. (CP) nanovesicles (NVs) from in vitro cultured cells and from dried Betulae folium, Betula pubescens Ehrh. and/or Betula pendula Roth (BP) and related supernatants (S) of NV isolation before (I) and after (II) washing vesicles. Similar to the membrane dye DiOC₆, lipophilic chlorophylls appeared to be enriched in the vesicle fraction. While no blue-fluorescing zones occurred in birch NVs, those from C. plantagineum showed such zones, probably due to caffeic acid derivatives that could make their way into the vesicles because of the acidic pH of the cell suspension growth medium. (b) Influence of buffer pH on secondary metabolites in NVs at 366 nm. While phenylpropanoids were not found in NVs if isolated under alkaline conditions, preparation in an acidic environment resulted in the detection of lipophilic yellow- and blue-fluorescing substances, probably aglyca of flavonoids and caffeic acid derivatives. DiOC₆: 3,3’-dihexyloxacarbocyanine iodide; Rf: Retention factor; TBS: Tris-buffered saline; VIB: vesicle isolation buffer.

Figure 6

HPTLC phospholipid profiles of methanol-chloroform extracts from nanovesicles isolated from Uvae-ursi folium, Arctostaphylos uva-ursi (L.) Spreng. (AU), Craterostigma plantagineum Hochst. (CP) cells, Curcumae zanthorrhizae rhizoma, Curcuma zanthorrhiza Roxb. (CZ) and Zingiberis rhizoma, Zingiber officinale Roscoe (ZO) after derivatization with 10% (m/v) CuSO₄ in 8% (v/v) H₃PO₄ and 10–15 min heating to 140–145 °C, charring analytes. Rf: Retention factor.