Abstract
Extracellular vesicles (EVs) play an important role in intercellular communication by transporting proteins and RNA. While plant cells secrete EVs, they have only recently been isolated and questions regarding their biogenesis, release, uptake and function remain unanswered. Here, we present a detailed protocol for isolating EVs from the apoplastic wash of Arabidopsis thaliana leaves. The isolated EVs can be quantified using a fluorometric dye to assess total membrane content.
Keywords: Arabidopsis thaliana, Extracellular vesicles, EVs, Apoplastic wash, DiOC6, Fluorometric quantification
Background
Extracellular vesicles (EVs) are membrane-bound structures that mediate the cell-to-cell transfer of proteins, lipids and genetic material. Interest in mammalian EVs has grown over the years due to their ability to transfer RNA and modulate immune responses. Mammalian EVs are routinely isolated for study from the medium of cultured cells, as well as a growing list of biological fluids (Colombo et al., 2014). Plant EVs are also thought to have a role in the immune response but are comparatively understudied (An et al., 2007; Davis et al., 2016). This is due, in large part, to the absence of a method of isolation. While plant EVs have been observed since 1967 using transmission electron microscopy, methods for their isolation were not developed until 2009 (Halperin and Jensen, 1967). Regente et al. (2009) isolated small (50-200 nm in diameter) vesicle-like structures from water-imbibed sunflower (Helianthus annuus) seeds. We modified the methods presented in Regente et al. (2009) to isolate vesicles from the apoplastic wash of Arabidopsis thaliana rosettes. To determine which conditions induce or impair EV secretion, we also designed a method for staining the EV pellet with 3,3’-dihexyloxacarbocyanine iodide (DIOC6(3)), a fluorescent lipophilic dye. In the absence of sophisticated forms of nanoparticle tracking, this relatively simple approach quantifies the total membrane content and can be used to indirectly measure the concentration of EVs (Rutter and Innes, 2017). For more precise measurements, and to assess the size distributions of EVs, nanoparticle tracking can be used. Our protocols enable the study of plant EV content and composition, as well as the pathways and conditions that mediate EV biogenesis and release.
Materials and Reagents
Equipment
Procedure
Data analysis
Table 1 and Figure 5 show a typical data analysis for DiOC6 fluorescence, which includes the following steps:
Notes
Recipes
Acknowledgments
This work was supported by a grant from the United States National Science Foundation (IOS-1645745) to R.W.I. A condensed version of this protocol was presented in Rutter and Innes (2017).
References
If you have any questions/comments about this protocol, you are highly recommended to post here. We will invite the authors of this protocol as well as some of its users to address your questions/comments. To make it easier for them to help you, you are encouraged to post your data including images for the troubleshooting.
Hi Yongfang,We chose this particular French press because:1) Its lid and plunger allow us to keep the rosettes submerged in the buffer during the vacuum infiltration,2) it fits inside our vacuum chamber,3) it is the perfect diameter to hold 6-week old Arabidopsis rosettes and4) and it is easy to clean.Any container fulfilling these requirements could be used in place of the French press we recommend.Best of luck,Brian
That’s correct. The 10 mM stock of DiOC6 should be dissolved in DMSO. Then, dilute it in Tris-HCl to make a 100 uM solution.
Hi Chuan Shen,Vacuum infiltration will very depending on the strength of you vacuum pump. I would say as long as your plants are being uniformly infiltrated and aren't leaking chloroplast into your wash fluids (as indicated by a green color) it's probably okay.It shouldn't have an effect on your results whether you use 2*C or 4*C. You merely want to keep the EVs stable in a cool temperature while centrifuging.I use MES (pH 6) when initially isolating EVs or handling them in general. This buffer is isotonic and was designed to match the pH of the apoplast, which is slightly acidic. I use Tris-HCl (pH 7.5) when staining with DiOC6, just in case an acidic pH affects DiOC6 fluorescence. The EVs are stable in either buffer.I think your problem is probably in your technique and the equipment you're using during the ultracentrifuge steps. I use a tabletop ultracentrifuge and 3.5 ml thick wall, polycarbonate tubes with a fixed-angle rotor. I always make sure the tube contains its maximum volume. This prevents cracking of the tubes and washes away impurities. I also decant the supernatant. If you're using a swinging bucket rotor, this creates a loose pellet that can be easily lost. The shape of the tube can also affect the quality of your pellet, as well as pipetting out too much of the supernatant. Unfortunately, the pellet is invisible except when using very high amounts of apoplastic wash, so you'll have to guess where it is.I would recommend going over you technique. Try looking at Thery et al. (2006). It has lots of helpful tips on technique for isolating EVs: Théry, Clotilde, et al. "Isolation and characterization of exosomes from cell culture supernatants and biological fluids." Current protocols in cell biology 30.1 (2006): 3-22.If you're losing the majority of your EVs through your technique, you might also try using higher amounts of wash fluid.Best of luck,Brian
Hi Chuan Shen,I hope you got my email addressing you question.I’d like to help you, but I need more information. There are lots of reasons why you may be getting low fluorescence. It could be that you aren’t collecting enough vesicles, and you need to use more apoplastic wash in you experiments. It could be that you are losing your pellet during the centrifugation steps by using a different kind of tube or pipetting technique. Maybe your fluorometer has a different level of sensitivity or your DIOC has gone bad.If you share your protocol with me. Maybe I can see what could be the problem.Best of luck,Brian