Plant Exosomal Vesicles: Comparison
View Latest Version
Please note this is a comparison between Version 2 by Dean Liu and Version 1 by Shkryl Yury.

Exosomal nanoparticles (exosomes or nanovesicles) are biogenic membrane vesicles secreted by various cell types and represent a conservative mechanism of intercellular and interspecies communication in pro- and eukaryotic organisms.

  • exosomes
  • targeted drug delivery
  • multivesicular bodies

1. Introduction

The discovery and development of new effective pharmaceuticals are considered as priority outcomes of biomedical science, aimed to improve the quality of human life. In recent years, significant advances have been made in this field, which have led to the development of many new drugs, including low-molecular-weight compounds of natural and synthetic origin, therapeutic proteins, and nucleic acids [1,2,3,4][1][2][3][4]. However, an important issue of therapy is the delivery effectiveness of these molecules to target cells or tissues. The targeted delivery makes it possible to create the required dose of an effector in the right location, protects it from degradation, reduces the toxic impact, and makes the therapy process more economical [5,6][5][6].
Many materials and approaches for delivery have been developed to date by the means of inorganic and organic platforms. Inorganic carriers include various nanoparticles, mainly based on metals and silica, as well as various combinations thereof [7,8][7][8]. The particles are functionalized with active groups, such as amine, carboxyl, and thiol, that allow them to chemically bind bioactive substances [9]. In particular, the mesoporous silica nanoparticles attract a lot of attention as promising carriers due to their high surface area and pore volume [10]. The carbon-based nanomaterials, especially graphene and multi-walled carbon nanotubes, also have a great potential for encapsulation and controlled release of bioactive compounds [11]. Still, limitations, such as particle aggregation in biological fluids and high cost of production, may restrict their use in practical applications.
Organic nanocarriers based on proteins (e.g., zein, gliadin, albumin) and polysaccharides (e.g., chitosan, alginate, cellulose) are promising candidates for targeted drug delivery [12]. The natural biopolymers used for their fabrication ensure the improved biological compatibility of such platforms, along with other advantages of nanosized materials. However, the low stability may limit their medical application in particular cases. Liposomes are simple vesicular structures and have been widely used as transporters for about 40 years [13]. Due to the use of physiological lipids as building blocks, liposomes have many advantages in transdermal, topical, and pulmonary applications [5]. However, the short half-life and production cost of such systems limit their therapeutic use [14]. The lipid nanoparticles represent a new delivery method, with improved properties compared to liposomes, including the ease of fabrication processes and stability [15]. They became particularly well known worldwide due to their use as the mRNA-containing vaccines for coronavirus (COVID-19) developed by Moderna and BioNTech/Pfizer. Such problems as the low drug-loading efficiency and lack of sufficient clinical observation should be solved to promote further the benefits of these carriers [16]. In addition to the above-mentioned delivery methods, the biological vectors derived from bacteriophages, mammalian, and plant viruses are also clinically used [17].
Exosomal nanoparticles from various biological objects, including mammals, plants, fungi, and bacteria have emerged as a new category of membrane vectors [17]. The practical application of plant exosomes requires a solid scientific basis and understanding of their biogenesis, molecular composition, physical characteristics, and biological properties as well as safety and effectiveness in disease treatment. Actually, thorough research in this field has only recently begun and thus many questions remain unsolved.

2. General Characteristics

Membrane vesicles (exosomes) are biogenic nano-formations with a characteristic size of 30–200 nm, which are released from the cell by fusion of the multivesicular body with the plasma membrane [18]. Other varieties of extracellular membrane structures differ from the exosomes by their size, origin, and function. For example, the particles of 200–1000 nm, formed by budding of the plasma membrane, are related to microvesicles, while the structures of more than 1000 nm are related to apoptotic bodies, the products of the cell decay via programmed cell death [19]. The existence of nanovesicles in plants was questioned until numerous studies proving their presence in all plant organs were carried out in the last decade [20]. Morphologically, the plant nanovesicles have a rounded shape formed by a phospholipid bilayer with an average thickness of 5.3 nm [21]. The exosomal membrane protects their molecular contents from enzymatic degradation, as well as from environmental influences (e.g., high and low temperatures, extreme pH, high salinity, moisture, and sunlight) [22,23,24][22][23][24]. Any parts of plants can serve as a source of nanovesicles for biomedical purposes, with the most preferred being the leaves, fruits, and apoplastotic fluid. It should be noted that the different plant organs produce different amounts of nanovesicles with unique compositions and properties, which may reflect their specializations in intercellular communications [25].

3. Functions of Plant Nanovesicles

The extracellular vesicles were previously believed to be mainly needed for cellular waste removal and do not have any important function. However, recent studies have shown that exosomes mediate intercellular communication by transporting various biologically active molecules in the organism. This issue is currently actively investigated in various animal and human models. Their exosomes play important roles in the regulation of developmental processes, activation of the immune system, and protective mechanisms in response to stresses, maintaining the pluripotency of embryonic stem cells, and in many other functions [63][26]. The functions of nanovesicles in plants are mostly considered in terms of their protective function in the system of plant-pathogen interactions. The ability of plants to quickly respond to various pathogens is essential for their survival. Restructuring the signaling pathways, cytoskeleton, and cell wall, as well as increased synthesis of the defense compounds, lead to the formation of physical (modification of plant cell wall) and biochemical (defense-related molecules) barriers that are designed to resist infection [64][27]. Such changes can be carried out through the rapid and targeted delivery of the necessary molecules by vesicles. It has been shown that the fungal infection enhances the rapid accumulation of exosomes between the plasma membrane and cell wall in plant cells, indicating their important role in the immune response [41,65][28][29]. For example, the vesicular structures containing polyphenolic metabolites and hydrogen peroxide prevented infection of the barley leaves by the powdery mildew fungus Blumeria graminis [66][30]. During the infection of Arabidopsis by a biotrophic fungus Golovinomyces orontii, the PEN1 and PEN3 proteins were transported by the exosomes and incorporated into the cell wall, acting as a protective barrier [67][31]. The transfer of miRNAs from the host plant to the pathogen, causing silencing of virulence genes, has also been described in the plant pathosystems, such as cotton/Verticillium dahliae and wheat/Fusarium graminearum [57,58][32][33]. At the same time, pathogens can transport their own miRNAs in the invaded area, contributing to the suppression of the immune response and defense systems of their host plant [39][34]. The grain yellow rust pathogen Puccinia striiformis produces miRNA-like RNA that suppresses the expression of defense genes in wheat [68][35]. The transfer of specific miRNAs of Hyaloperonospora arabidopsidis and Botrytis cinerea into the cells of Arabidopsis thaliana suppresses the expression of AGO1 protein, thus disrupting the loading of the plant nanovesicles with protective miRNAs [69][36]. Notably, the plant pathogenic fungi also widely utilize the extracellular vesicles to transport the effectors through host barriers aiming to suppress the plant immunity [40][37].

4. Characterization of Exosomes

The characterization of plant nanovesicles includes a wide spectrum of morphological, physical, and biochemical analytical approaches. One of the most common imaging techniques used is scanning and transmission electronic microscopy (SEM and TEM, respectively) [83][38]. SEM provides information about the three-dimensional structure of the exosome surface, however, as soon as drying is applied during sample preparation, their natural morphology could be changed, leading to the formation of cup-shaped structures [84][39]. In turn, TEM is a more accurate method, as nanovesicles are not deformed during processing. Moreover, staining with heavy metals, such as osmium tetroxide and uranyl acetate, creates lipid membrane contrasts allowing distinguishing exosomes from impurities [85][40]. Cryo-SEM, or cryomicroscopy, implies freezing and sample analysis at a very low temperature (below −100 °C) [86][41]. It is used to assess the morphology of nanovesicles in a state close to the native one due to their perfect preservation, as well as the absence of pre-fixation operations or the addition of heavy metals [21,83][21][38]. Atomic force microscopy can also be used as an additional indirect method for assessing the morphological and physical properties (e.g., adhesion and stiffness) of nanovesicles [87][42]. The method does not require extensive sample preparation: exosomes could be rapidly adsorbed and dried on glass or mica surfaces. The hydrodynamic sizes of exosomes are evaluated by using dynamic light scattering (DLS) or nanoparticle tracking analysis (NTA). DLS measures the volumetric scattered light from nanovesicles when illuminated with a monochromatic light source. Since the particles are in Brownian motion, scattered light from all particles interferes and intensity fluctuates with time, and information about the particles is obtained from the autocorrelation of the oscillation intensity recorded during the experiment [88][43]. At the same time, NTA is based on the use of a concentrated beam of light to illuminate the particles in the sample. As the particles scatter light and undergo Brownian motion, the camera records the path of each individual particle to determine the average speed and size [85][40]. As the measurement of nanoparticles using the latter method is more accurate when analyzing polydispersed samples [89[44][45],90], NTA is now considered a gold standard for exosome characterization [91][46]. The identification of specific protein markers is one of the most important biochemical characteristics of exosomes. Usually, Western blotting with antibodies specific to such proteins as PEN3, TET8, and HSP70 indicates the plant-derived nanovesicle fraction [92][47]. However, in the case of exosomes from the non-model plants, the commercially available antibodies for Arabidopsis thaliana may have a low affinity to the proteins from evolutionarily distant species. Another versatile way to identify proteins, including marker ones, is mass spectrometry [93][48]. In addition to the purification, the peptide fractionation prior to mass spectrometric analysis is considered an important prerequisite for the identification of vesicular proteins with high confidence. In terms of detection sensitivity, mass spectrometry is not as sensitive as antibody-based methods, but it allows analysis of a whole spectrum of exosomal proteins at once [94][49]. As mentioned above, the purity of the exosomes must be checked very strictly, since the identification of many hundreds of proteins in exosomes is probably due to contamination [42][50]. Mass spectrometry can also be used to analyze the low molecular weight components, such as secondary metabolites, fatty acids, and sugars [95,96][51][52]. New generation sequencing (NGS) methods are used to identify the nucleic acid fractions, primarily the small RNAs [97][53]. Taking into account that a huge fraction of small RNAs is coprecipitated with exosomes in the complexes with proteins, protease trypsin digestion should precede RNase treatment to ensure degradation of RNAs located outside vesicles [48][54]. For low-scale analysis, quantitative real-time PCR, applying a stem-loop approach [98][55] or poly(A)-tailing [99][56] could be used to verify the individual miRNA representatives. In this term, an interesting question is whether there are inherent miRNAs that can be used as markers along with protein components.

References

  1. Sridharan, K.; Gogtay, N.J. Therapeutic nucleic acids: Current clinical status. Br. J. Clin. Pharmacol. 2016, 82, 659–672.
  2. Wang, L.; Wang, N.; Zhang, W.; Cheng, X.; Yan, Z.; Shao, G.; Wang, X.; Wang, R.; Fu, C. Therapeutic peptides: Current applications and future directions. Signal Transduct. Target. Ther. 2022, 7, 48.
  3. Dzobo, K. The role of natural products as sources of therapeutic agents for innovative drug discovery. Compr. Pharmacol. 2022, 408–422.
  4. Gerry, C.; Schreiber, S. Chemical probes and drug leads from advances in synthetic planning and methodology. Nat. Rev. Drug Discov. 2018, 17, 333–352.
  5. Tewabe, A.; Abate, A.; Tamrie, M.; Seyfu, A.; Abdela Siraj, E. Targeted drug delivery—From magic bullet to nanomedicine: Principles, challenges, and future perspectives. J. Multidiscip. Healthc. 2021, 5, 1711–1724.
  6. Manzari, M.T.; Shamay, Y.; Kiguchi, H.; Rosen, N.; Scaltriti, M.; Heller, D.A. Targeted drug delivery strategies for precision medicines. Nat. Rev. Mater. 2021, 6, 351–370.
  7. Desai, N.; Momin, M.; Khan, T.; Gharat, S.; Ningthoujam, R.S.; Omri, A. Metallic nanoparticles as drug delivery system for the treatment of cancer. Expert Opin. Drug Deliv. 2021, 18, 1261–1290.
  8. De Oliveira, L.F.; Bouchmella, K.; De Almeida Gonçalves, K.; Bettini, J.; Kobarg, J.; Borba Cardoso, M. Functionalized silica nanoparticles as an alternative platform for targeted drug-delivery of water insoluble drugs. Langmuir 2016, 32, 3217–3225.
  9. Seidu, T.A.; Kutoka, P.T.; Asante, D.O.; Farooq, M.A.; Alolga, R.N.; Bo, W. Functionalization of nanoparticulate drug delivery systems and its influence in cancer therapy. Pharmaceuticals 2022, 14, 1113.
  10. Bharti, C.; Nagaich, U.; Pal, A.K.; Gulati, N. Mesoporous silica nanoparticles in target drug delivery system: A review. Int. J. Pharm. Investig. 2015, 5, 124–133.
  11. Maiti, D.; Tong, X.; Mou, X.; Yang, K. Carbon-based nanomaterials for biomedical applications: A recent study. Front. Pharmacol. 2019, 9, 1401.
  12. Patra, J.K.; Das, G.; Fraceto, L.F.; Campos, E.V.R.; del Pilar Rodriguez-Torres, M.; Acosta-Torres, L.S.; Diaz-Torres, L.A.; Grillo, R.; Swamy, M.K.; Sharma, S.; et al. Nano based drug delivery systems: Recent developments and future prospects. J. Nanobiotechnol. 2018, 16, 71.
  13. 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.
  14. Sercombe, L.; Veerati, T.; Moheimani, F.; Wu, S.Y.; Sood, A.K.; Hua, S. Advances and challenges of liposome assisted drug delivery. Front. Pharmacol. 2015, 6, 286.
  15. Dhiman, N.; Awasthi, R.; Sharma, B.; Kharkwal, H.; Kulkarni, G.T. Lipid nanoparticles as carriers for bioactive delivery. Front. Chem. 2021, 9, 580118.
  16. Ghasemiyeh, P.; Mohammadi-Samani, S. Solid lipid nanoparticles and nanostructured lipid carriers as novel drug delivery systems: Applications, advantages and disadvantages. Res. Pharm. Sci. 2018, 13, 288–303.
  17. Chen, L.; Hong, W.; Ren, W.; Xu, T.; Qian, Z.; He, Z. Recent progress in targeted delivery vectors based on biomimetic nanoparticles. Signal Transduct. Target. Ther. 2021, 6, 225.
  18. Yang, M.; Liu, X.; Luo, Q.; Xu, L.; Chen, F. An efficient method to isolate lemon derived extracellular vesicles for gastric cancer therapy. J. Nanobiotechnol. 2020, 18, 100.
  19. Harding, C.; Heuser, J.; Stahl, P. Endocytosis and intracellular processing of transferrin and colloidal gold-transferrin in rat reticulocytes: Demonstration of a pathway for receptor shedding. Eur. J. Cell Biol. 1984, 35, 256–263.
  20. An, Q.; van Bel, A.J.; Huckelhoven, R. Do plant cells secrete exosomes derived from multivesicular bodies? Plant Signal. Behav. 2007, 2, 4–7.
  21. Garaeva, L.; Kamyshinsky, R.; Kil, Y.; Varfolomeeva, E.; Verlov, N.; Komarova, E.; Garmay, Y.; Landa, S.; Burdakov, V.; Myasnikov, A.; et al. Delivery of functional exogenous proteins by plant-derived vesicles to human cells in vitro. Sci. Rep. 2021, 11, 6489.
  22. Perez-Bermudez, P.; Blesa, J.; Soriano, J.M.; Marcilla, A. Extracellular vesicles in food: Experimental evidence of their secretion in grape fruits. Eur. J. Pharm. Sci. 2017, 98, 40–50.
  23. Kim, K.; Park, J.; Sohn, Y.; Oh, C.E.; Park, J.H.; Yuk, J.M.; Yeon, J.H. Stability of Plant Leaf-Derived Extracellular Vesicles According to Preservative and Storage Temperature. Pharmaceutics 2022, 14, 457.
  24. Yuan, F.; Li, Y.M.; Wang, Z. Preserving extracellular vesicles for biomedical applications: Consideration of storage stability before and after isolation. Drug Deliv. 2021, 28, 1501–1509.
  25. Patil, A.A.; Rhee, W.J. Exosomes: Biogenesis, Composition, Functions, and Their Role in Pre-metastatic Niche Formation. Biotechnol. Bioprocess Eng. 2019, 24, 689–701.
  26. Tkach, M.; Thery, C. Communication by Extracellular Vesicles: Where We Are and Where We Need to Go. Cell 2016, 164, 1226–1232.
  27. Frei dit Frey, N.; Robatzek, S. Trafficking vesicles: Pro or contra pathogens? Curr. Opin. Plant Biol. 2009, 12, 437–443.
  28. An, Q.; Ehlers, K.; Kogel, K.H.; Van Bel, A.J.E.; Huckelhoven, R. Multivesicular compartments proliferate in susceptible and resistant MLA12-barley leaves in response to infection by the biotrophic powdery mildew fungus. New Phytol. 2006, 172, 563–576.
  29. Wang, F.; Shang, Y.; Fan, B.; Yu, J.Q.; Chen, Z. Arabidopsis LIP5, a positive regulator of multivesicular body biogenesis, is a critical target of pathogen-responsive MAPK cascade in plant basal defense. PLoS Pathog. 2014, 10, e1004243.
  30. Mustafa, G.; Khong, N.G.; Tisserant, B.; Randoux, B.; Fontaine, J.; Magnin-Robert, M.; Reignault, P.; Sahraoui, A.L. Defence mechanisms associated with mycorrhiza-induced resistance in wheat against powdery mildew. Funct. Plant Biol. 2017, 44, 443–454.
  31. Meyer, D.; Pajonk, S.; Micali, C.; O’Connell, R.; Schulze-Lefert, P. Extracellular transport and integration of plant secretory proteins into pathogen-induced cell wall compartments. Plant J. 2009, 57, 986–999.
  32. Wang, B.; Zhuang, X.; Deng, Z.B.; Jiang, H.; Mu, J.; Wang, Q.; Xiang, X.; Guo, H.; Zhang, L.; Dryden, G.; et al. Targeted drug delivery to intestinal macrophages by bioactive nanovesicles released from grapefruit. Mol. Ther. 2014, 22, 522–534.
  33. Campo, S.; Peris-Peris, C.; Sire, C.; Moreno, A.B.; Donaire, L.; Zytnicki, M.; Notredame, C.; Llave, C.; San Segundo, B. Identification of a novel microRNA (miRNA) from rice that targets an alternatively spliced transcript of the Nramp6 (Natural resistance-associated macrophage protein 6) gene involved in pathogen resistance. New Phytol. 2013, 199, 212–227.
  34. Nielsen, M.E.; Feechan, A.; Bohlenius, H.; Ueda, T.; Thordal-Christensen, H. Arabidopsis ARF-GTP exchange factor, GNOM, mediates transport required for innate immunity and focal accumulation of syntaxin PEN1. Proc. Natl. Acad. Sci. USA 2012, 109, 11443–11448.
  35. Wang, B.; Sun, Y.; Song, N.; Zhao, M.; Liu, R.; Feng, H.; Wang, X.; Kang, Z. Puccinia striiformis f. sp. tritici microRNA-like RNA 1 (Pst-milR1), an important pathogenicity factor of Pst, impairs wheat resistance to Pst by suppressing the wheat pathogenesis-related 2 gene. New Phytol. 2017, 215, 338–350.
  36. Weiberg, A.; Wang, M.; Lin, F.M.; Zhao, H.; Zhang, Z.; Kaloshian, I.; Huang, H.D.; Jin, H. Fungal small RNAs suppress plant immunity by hijacking host RNA interference pathways. Science 2013, 342, 118–123.
  37. Samuel, M.; Bleackley, M.; Anderson, M.; Mathivanan, S. Extracellular vesicles including exosomes in cross kingdom regulation: A viewpoint from plant-fungal interactions. Front. Plant Sci. 2015, 6, 766.
  38. Echlin, P. Low-temperature scanning electron microscopy. In Low-Temperature Microscopy and Analysis; Plenum Press: New York, NY, USA, 1992; pp. 349–411.
  39. Noble, J.M.; Roberts, L.D.M.; Vidavsky, N.; Chiou, A.E.; Fischbach, C.; Paszek, M.J.; Estroff, L.A.; Kourkoutis, L.F. Direct comparison of optical and electron microscopy methods for structural characterization of extracellular vesicles. J. Struct. Biol. 2020, 210, 107474.
  40. Shao, H.; Chung, J.; Balaj, L.; Charest, A.; Bigner, D.D.; Carter, B.S.; Hochberg, F.H.; Breakefield, X.O.; Weissleder, R.; Lee, H. Protein typing of circulating microvesicles allows real-time monitoring of glioblastoma therapy. Nat. Med. 2012, 18, 1835–1840.
  41. Tatischeff, I.; Larquet, E.; Falcon-Perez, J.M.; Turpin, P.Y.; Kruglik, S.G. Fast characterisation of cell-derived extracellular vesicles by nanoparticles tracking analysis, cryo-electron microscopy, and raman tweezers microspectroscopy. J. Extracell. Vesicles 2012, 1, 19179.
  42. Sharma, S.; Rasool, H.I.; Palanisamy, V.; Mathisen, C.; Schmidt, M.; Wong, D.T.; Gimzewski, J.K. Structural-mechanical characterization of nanoparticle exosomes in human saliva, using correlative afm, fesem, and force spectroscopy. ACS Nano 2010, 4, 1921–1926.
  43. Sitar, S.; Kejzar, A.; Pahovnik, D.; Kogej, K.; Tusek-Znidaric, M.; Lenassi, M.; Zagar, E. Size characterization and quantification of exosomes by asymmetrical-flow field-flow fractionation. Anal. Chem. 2015, 87, 9225–9233.
  44. Hou, J.; Ci, H.; Wang, P.; Wang, C.; Lv, B.; Miao, L.; You, G. Nanoparticle tracking analysis versus dynamic light scattering: Case study on the effect of Ca2+ and alginate on the aggregation of cerium oxide nanoparticles. J. Hazard. Mater. 2018, 360, 319–328.
  45. Filipe, V.; Hawe, A.; Jiskoot, W. Critical evaluation of nanoparticle tracking analysis (nta) by nanosight for the measurement of nanoparticles and protein aggregates. Pharm. Res. 2010, 27, 796–810.
  46. Szatanek, R.; Baj-Krzyworzeka, M.; Zimoch, J.; Lekka, M.; Siedlar, M.; Baran, J. The methods of choice for extracellular vesicles (evs) characterization. Int. J. Mol. Sci. 2017, 18, 1153.
  47. Rutter, B.D.; Innes, R.W. Growing pains: Addressing the pitfalls of plant extracellular vesicle research. New Phytol. 2020, 228, 1505–1510.
  48. Thongboonkerd, V. Practical points in urinary proteomics. J. Proteome Res. 2007, 6, 3881–3890.
  49. Kreimer, S.; Belov, A.M.; Ghiran, I.; Murthy, S.K.; Frank, D.A.; Ivanov, A.R. Mass-spectrometry-based molecular characterization of extracellular vesicles: Lipidomics and proteomics. J. Proteome Res. 2015, 14, 2367–2384.
  50. Burkova, E.E.; Grigoreva, A.E.; Bulgakov, D.V.; Dmitrenok, P.S.; Vlassov, V.V.; Ryabchikova, E.I.; Sedykh, S.E.; Nevinsky, G.A. Extra Purified Exosomes from Human Placenta Contain an Unpredictable Small Number of Different Major Proteins. Int. J. Mol. Sci. 2019, 20, 2434.
  51. Kedjouar, B.; De Medina, P.; Oulad-Abdelghani, M.; Payre, B.; Silvente-Poirot, S.; Favre, G.; Faye, J.C.; Poirot, M. Molecular characterization of the microsomal tamoxifen binding site. J. Biol. Chem. 2004, 279, 34048–34061.
  52. Mayr, M.; Grainger, D.; Mayr, U.; Leroyer, A.S.; Leseche, G.; Sidibe, A.; Herbin, O.; Yin, X.; Gomes, A.; Madhu, B.; et al. Proteomics, metabolomics, and immunomics on microparticles derived from human atherosclerotic plaques. Circ. Cardiovasc. Genet. 2009, 2, 379–388.
  53. Yagi, Y.; Ohkubo, T.; Kawaji, H.; Machida, A.; Miyata, H.; Goda, S.; Roy, S.; Hayashizaki, Y.; Suzuki, H.; Yokota, T. Next-generation sequencing-based small RNA profiling of cerebrospinal fluid exosomes. Neurosci. Lett. 2017, 636, 48–57.
  54. Zand Karimi, H.; Baldrich, P.; Rutter, B.D.; Borniego, L.; Zajt, K.K.; Meyers, B.C.; Innes, R.W. Arabidopsis apoplastic fluid contains sRNA- and circular RNA–protein complexes that are located outside extracellular vesicles. Plant Cell 2022, 34, 1863–1881.
  55. Lin, Q.; Mao, W.; Shu, Y.; Lin, F.; Liu, S.; Shen, H.; Gao, W.; Li, S.; Shen, D. A cluster of specified microRNAs in peripheral blood as biomarkers for metastatic non-small-cell lung cancer by stem-loop RT-PCR. J. Cancer Res. Clin. Oncol. 2012, 138, 85–93.
  56. Li, X.; Chen, C.; Wang, Z.; Liu, J.; Sun, W.; Shen, K.; Lv, Y.; Zhu, S.; Zhan, P.; Lv, T.; et al. Elevated exosome-derived miRNAs predict osimertinib resistance in non-small cell lung cancer. Cancer Cell Int. 2021, 21, 428.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)
Video Production Service
Feedback