
==== Front
Genetics
Genetics
genetics
Genetics
0016-6731
1943-2631
Oxford University Press US

38884207
10.1093/genetics/iyae088
iyae088
WormBook
Cell and Organelle Biology
AcademicSubjects/SCI01180
AcademicSubjects/SCI01140
Featured
Extracellular vesicles
Wang Juan Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ 08854, USA

https://orcid.org/0000-0003-4483-2952
Barr Maureen M Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ 08854, USA

https://orcid.org/0000-0001-9826-4132
Wehman Ann M Department of Biological Sciences, University of Denver, Denver, CO 80210, USA

Greenwald I Editor
Corresponding author: Department of Biological Sciences, University of Denver, 2101 E. Wesley Ave, Denver, CO 80208, USA. Email: awehman@alum.mit.edu
Conflicts of interest The author(s) declare no conflicts of interest.

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Abstract

Extracellular vesicles (EVs) encompass a diverse array of membrane-bound organelles released outside cells in response to developmental and physiological cell needs. EVs play important roles in remodeling the shape and content of differentiating cells and can rescue damaged cells from toxic or dysfunctional content. EVs can send signals and transfer metabolites between tissues and organisms to regulate development, respond to stress or tissue damage, or alter mating behaviors. While many EV functions have been uncovered by characterizing ex vivo EVs isolated from body fluids and cultured cells, research using the nematode Caenorhabditis elegans has provided insights into the in vivo functions, biogenesis, and uptake pathways. The C. elegans EV field has also developed methods to analyze endogenous EVs within the organismal context of development and adult physiology in free-living, behaving animals. In this review, we summarize major themes that have emerged for C. elegans EVs and their relevance to human health and disease. We also highlight the diversity of biogenesis mechanisms, locations, and functions of worm EVs and discuss open questions and unexplored topics tenable in C. elegans, given the nematode model is ideal for light and electron microscopy, genetic screens, genome engineering, and high-throughput omics.

Extracellular vesicles (EV) play increasingly diverse roles during development, physiology, and disease. In vivo studies in the nematode Caenorhabditis elegans have revealed insights into the mechanisms of EV formation and an array of EV functions for the EV-releasing cell or EV-receiving cells. This review summarizes the current state of the art and important directions for future research.

extracellular vesicle
Caenorhabditis elegans
exosome
microvesicle
cilia
spermatogenesis
midbody remnant
American Cancer Society 10.13039/100000048 Deutsche Forschungsgemeinschaft 10.13039/501100001659 WE5719/2-1 National Institutes of Health 10.13039/100000002 DK059418 DK116606 NS120745
==== Body
pmcWhat are extracellular vesicles?

Beyond the plasma membrane, cells have extracellular organelles, including the extracellular matrix and extracellular vesicles (EVs). Cells from bacteria to animals, including specialized cells like neurons, produce EVs using evolutionarily conserved molecular mechanisms (Colombo et al. 2014). EVs were originally considered a type of cell debris (Chargaff and West 1946), but studies over the past few decades have revealed a wide range of EV functions from signaling and transport to cellular remodeling or defense. See Couch et al. (2021) for a fascinating review on the history of EV research, one of the cutting edges of cell and organelle biology.

The term “extracellular vesicle”? was first used to describe membrane-bound particles secreted into culture media by the algae Ochromonas danica, as observed by negative staining transmission electron microscopy (TEM) (Aaronson et al. 1971), but human blood EVs were already observed decades earlier (Chargaff and West 1946). EVs are spherical or tubular vesicles released by cells into their environment. EVs are produced through diverse cellular processes that are active and regulated at specific locations within cells (Dixson et al. 2023; Sohal and Kasinski 2023). EVs reflect their cellular origin and are composed of a lipid bilayer membrane that encloses cytosolic organelles and macromolecules, including proteins, metabolites, and nucleic acids (such as mRNA, microRNA, and DNA).

The International Society for Extracellular Vesicles (ISEV) has community guidelines for classifying EV subtypes published in the Minimal Information for Studies of Extracellular Vesicles (MISEV) (Théry et al. 2018; Welsh et al. 2024). EVs are typically classified based on their size into small and large EVs, which is derived from the methods used to separate EVs from body fluids. Small EVs, with diameters ranging from tens to hundreds of nanometers, are further subdivided based on their subcellular origins, including exosomes and microvesicles (MVs). Exosomes originate from the endosomal system, forming when multivesicular bodies fuse with the plasma membrane to release their vesicular contents outside the cell. In contrast, MVs bud directly from the plasma membrane into the extracellular space. Despite their distinct subcellular origin, exosomes and MVs can overlap in their size range, contents, and molecular regulation, making it challenging to distinguish small EV subtypes after their release. Discriminating exosomes from MVs has been a major challenge for ex vivo EV studies, with no clear consensus on specific markers (Théry et al. 2018; Welsh et al. 2024).

Large EVs, with diameters in the micron range, encompass a variety of structures such as midbody remnants from dividing cells, apoptotic bodies from dying cells, and exophers, which may be as large as the cell body that released them. Large EV subtypes have specific cargo or biogenesis mechanisms that distinguish them from other subtypes. Large EVs can even contain membrane-bound organelles like mitochondria or multivesicular bodies and become a secondary source of small EVs. Overall, diverse small and large EV subtypes can be utilized for different cellular functions.

EVs are implicated in human health, aging, and disease progression as EVs are involved in a wide range of physiological functions, including intercellular communication and immune regulation (Berumen Sanchez et al. 2021; Buzas 2023; Polyakova et al. 2023). EVs can be hijacked by viruses for infection or exploited by parasites to evade the host immune system (Bou et al. 2023; Kuipers et al. 2018). Based on the capability of cells to take up EVs, EVs are being studied for their potential in drug delivery and other therapeutic applications (Thakur et al. 2022; Cheng and Hill 2022). EVs are also of great interest to human medicine for their diagnostic potential as cellular biomarkers. EVs containing cellular content are released into body fluids, and blood collection is typically less invasive than tissue biopsy.

Major challenges in the EV field include a limited understanding of the molecular control of EV biogenesis and difficulties tracking EV release in vivo. Observing structures with light microscopy is technically challenging for objects smaller than 200 nm, especially within the tissue context. Furthermore, it has been challenging to understand the complexity and heterogeneity of EVs and the diverse functions of EV subpopulations often found in a heterogeneous EV sample isolated from body fluids. Understanding the commonalities and intricacies of EVs will enable the field to harness EVs for diagnostic and therapeutic purposes. Caenorhabditis elegans serves as an excellent model for showcasing the diversity of EV formation and function within an intact organism due to its transparency, small size, and the extensive knowledge base established by the C. elegans community (Fig. 1, Table 1). This WormBook chapter summarizes the current state of the art in EV biology in C. elegans and highlights key features including the simplicity, tractability, and versatile genetic toolkit that make C. elegans an invaluable animal model for EV research.

Fig. 1. Caenorhabditis elegans release diverse EVs from various tissues. Distinct EV subtypes are released from diverse tissues during different developmental states in male and hermaphrodite worms, such as epidermal cells, neurons, muscle, and germ cells, as well as undifferentiated embryonic blastomeres. Each EV subtype has a characteristic size range from tens of nanometers to several microns.

Table 1. Diverse cell types release and internalize diverse EVs.

Cell type	Example cells	EV subtype	Event	References	
Undifferentiated					
Embryonic blastomeres	P0, AB, P1, etc.	Microvesicle	Release	Wehman et al. (2011), Beer et al. (2018)	
Embryonic blastomeres	P0, AB, P1, etc.	Midbody remnant	Release	Green et al. (2013), König et al. (2017)	
Embryonic blastomeres	EMS, P2, etc.	Midbody remnant	Uptake	Ou et al. (2014), Singh and Pohl (2014) , Fazeli et al. (2016), König et al. (2017), Bai et al. (2020), Fazeli et al. (2020)	
Ectodermal					
Epidermis	H1-H2, V1-V6, T	Exosome	Release	Liegeois et al. (2006), Hyenne et al. (2015)	
Epidermis	H1-H2, V1-V6, T	Microvesicle	Release	Katz et al. (2022)	
Epidermis	H1-H2, V1-V6, T	Unknown	Release	Oren-Suissa et al. (2017)	
Epidermis	Hyp7	Midbody remnant	Uptake	Chai et al. (2012)	
Epidermis	Hyp7	Exopher	Uptake	Melentijevic et al. (2017), Wang, Arnold, et al. (2023)	
Neuroblasts	Q	Midbody remnant	Release	Chai et al. (2012)	
Ciliated neurons: male	CEM, RnB (n = 1–9, not 6), HOB, IL2, Amphid, Phasmid	Microvesicle	Release	Wang et al. (2014), Maguire et al. (2015), O'Hagan et al. (2017), Silva et al. (2017), Akella and Barr (2021), Wang et al. (2021), Razzauti and Laurent (2021) , Clupper et al. (2022), Nikonorova et al. (2022)	
Ciliated neurons: hermaphrodite	IL2, Amphid, Phasmid	Microvesicle	Release	Wang et al. (2014), Maguire et al. (2015), O'Hagan et al. (2017), Silva et al. (2017), Akella et al. (2020), Wang et al. (2021), Razzauti and Laurent (2021), Clupper et al. (2022), Nikonorova et al. (2022)	
Ciliated neurons with microvilli	AFD	Microvesicle	Release	Raiders et al. (2021)	
Mechanosensory touch neurons	AVM, ALM, PVM, PLM	Exopher	Release	Melentijevic et al. (2017), Arnold et al. (2020), Cooper et al. (2021), Arnold et al. (2023)	
Mechanosensory touch neurons	PVM	Microvesicle	Release	Linton et al. (2019)	
Nociceptive mechanosensory neuron	PVD	Unknown	Fusion?	Oren-Suissa et al. (2017)	
Dopaminergic neurons	CEP	Microvesicle?	Release	Ke (2020a, 2020b)	
Glia	AMsh, AMso	Microvesicle	Uptake	Razzauti and Laurent (2021), Raiders et al. (2021)	
Mesodermal					
Body wall muscle		Exopher	Release	Turek et al. (2021)	
Gonadal sheath cells	SS	Residual body	Uptake	Huang et al. (2012)	
Endodermal					
Intestinal cells	E	Lobes	Uptake	Abdu et al. (2016)	
Germ cells					
Primordial germ cell	Z2, Z3	Lobes	Release	Abdu et al. (2016), Maniscalco et al. (2020), Schwartz et al. (2022)	
Spermatocyte		Residual Body	Release	Roberts et al. (1986), Kelleher et al. (2000), Hu et al. (2019)	
Spermatid		MV (Mitopher)	Release	Liu et al. (2023)	
Spermatids and spermatozoa		Double-membrane MV	Release	Kosinski et al. (2005)	
Oocyte		Exopher	Uptake	Turek et al. (2021)	
Dying cells					
Apoptotic embryonic cells		Microvesicle	Release	Mapes et al. (2012)	

How are EVs formed?

Exosome formation from endolysosomes

The biogenesis mechanisms of exosomes have been illuminated by decades of research on endosomal trafficking pathways conserved from yeast to mammals. Many excellent reviews summarize exosome biogenesis pathways (Buzas 2023; Dixson et al. 2023), which typically start with the formation of intraluminal vesicles (ILVs) within the limiting membrane of an endosome to form a multivesicular body (MVB). The MVBs then fuse with the plasma membrane via exocytosis, releasing their ILVs outside the cell as exosomes (Fig. 2).

Fig. 2. Exosome biogenesis pathways. Cargo proteins can be endocytosed and sorted into the late endosome to localize to the surface of ILVs within the MVB. Cargo proteins can also be incorporated into the lumen of ILVs by budding of the limiting endosomal membrane. RAL-1, actin crosslinkers, and the ESCRT machinery are involved in ILV formation in C. elegans. The MVB is then transported proximally to the plasma membrane, where the MVB docks and releases its contents into the extracellular space. MVBs can also mature into lysosomes for degradation of ILV contents.

The Endosomal Sorting Complex Required for Transport (ESCRT) is well established to form ILVs by budding endosomal membranes away from the cytoplasm. The ESCRT pathway comprises four subcomplexes along with essential accessory proteins like the ATPase VPS-4. The ESCRT-0, I, and II subcomplexes play a crucial role in clustering ubiquitinated proteins into ILV buds, a critical step for sorting cargo into exosomes. ESCRT-III proteins form spiraling filaments that promote membrane curvature to start budding the ILVs. VPS-4 ATPase activity remodels the ESCRT-III filaments to constrict the bud neck for scission and release the ILV inside the MVB (Fig. 2).

In C. elegans embryos, ESCRT proteins localize to nascent MVBs after fertilization and are required for ILV formation (Frankel et al. 2017; Clarke et al. 2022). Notably, partial depletion of ESCRTs led to an increase in ILV diameter from ∼60 nm in wild type to ∼100 nm in ESCRT mutants. In contrast, disrupting an ESCRT-III-associated factor, ISTR-1, led to a decrease in ILV diameter to ∼35 nm (Frankel et al. 2017). As ILVs can give rise to exosomes, ESCRT perturbations would alter exosome diameter as well as quantity, which could also change exosome contents.

The ESCRT machinery is important for exosome biogenesis from epidermal cells, which are essential for alae formation in C. elegans (Fig. 1). Alae are ridge-like structures found on the cuticle of C. elegans, particularly visible in the L1 larval stage and adults. A genetic screen for alae defects identified 73 genes that may regulate exosome biogenesis (Fig. 2), including ESCRT-0 component HGRS-1, ESCRT-I components TSG-101, VPS-28, and VPS-37, ESCRT-II components VPS-22, VPS-25, and VPS-36, ESCRT-III components VPS-20 and VPS-32, as well as the ATPase VPS-4 (Hyenne et al. 2015). This suggests a role for the ESCRT complex in ILV formation for exosome secretion by epidermal cells to form alae.

MVB formation and maturation in the epidermis are also promoted by actin and the filamin FLN-2 (Shi et al. 2022). Using electron microscopy (EM), MVBs can be subdivided into electron-light MVBs and electron-dark MVBs (Fig. 2). Electron-light MVBs are thought to be either exosomal precursors for secretion or early lysosomal precursors, while electron-dark MVBs are thought to have matured and already begun lysosomal degradation (Liégeois et al. 2006). In fln-2 mutants, electron-light MVBs are reduced with fewer ILVs, while electron-dark MVBs are almost absent (Shi et al. 2022), suggesting that FLN-2 promotes both ILV formation and MVB maturation. Filamins crosslink F-actin to organize the cytoskeleton and other proteins on membranes, and FLN-2 appears to bridge MVBs to actin filaments. Inhibiting actin polymerization with latrunculin A also reduced the number of light and dark MVBs, suggesting that FLN-2 regulates the actin cytoskeleton to promote ILV budding on endosomes.

ILV budding is also promoted by the small GTPase RAL-1, which localizes to the surface of epidermal MVBs (Hyenne et al. 2015). There are fewer light or dark MVBs in ral-1 mutants and fewer ILVs in ral-1 MVBs. Intriguingly, the human RAL-1 homologs RalA and RalB are required for exosome release in cancer cells (Ghoroghi et al. 2021), suggesting that the Ral family GTPases have conserved roles regulating MVBs for exosome release. RalA and RalB interact with the phospholipase PLD1 (Ghoroghi et al. 2021), which can convert cylindrical phosphatidylcholine (PC) lipids to conical phosphatidic acid (PA) lipids in endosomal membranes (Egea-Jimenez and Zimmermann 2018). Lipid shape is thought to influence membrane curvature to aid membrane budding, but lipid conversions can also regulate lipid–protein interactions to regulate protein localization to endosomes.

RAL-1 is also required for MVB fusion to release ILVs as exosomes (Hyenne et al. 2015), which have been visualized using EM (Table 2). The constitutively active form of the RAL-1 GTPase colocalizes with the t-SNARE protein SYX-5, suggesting that RAL-1 interacts with factors that regulate MVB fusion. RAL-1 colocalizes with the V-ATPase subunit VHA-5 (Hyenne et al. 2015), whose V0 sector is required for MVB fusion with the plasma membrane (Liégeois et al. 2006).

Table 2. Established reporters for EV subtypes in different cell types.

Source Tissue	EV reporter	Method	EV subtype	Notes	Reference	
Mitotic cells						
Dividing cells	FP::ZEN-4	Fluorescence	Midbody remnant	Spindle midbody	Green et al. (2013)	
Dividing cells	NMY-2::FP	Fluorescence	Midbody remnant	Contractile ring component	Green et al. (2013)	
Dividing cells	FP::CYK-7	Fluorescence	Midbody remnant	Contractile ring component	Green et al. (2013)	
Dividing cells	FP::MVB-12	Fluorescence	Midbody remnant	ESCRT-I subunit	Green et al. (2013)	
Dividing cells	GRP-1 antibody	Antibody staining	Midbody remnant	Arf GEF associated with contractile ring	Teuliere (2014)	
Early Embryo	FP::ZF1::SYX-4	Fluorescent degron protection	Multiple subtypes	Transmembrane protein, somatic EVs after six-cell stage	Wehman et al. (2011)	
Early Embryo	pie-1::
FP::PH::ZF1	Fluorescent degron protection	Microvesicle	Lipid-binding domain, somatic MVs after six-cell stage	Beer et al. (2018)	
Early Embryo	pie-1::
FP::PH::CTPD	Fluorescent degron protection	Microvesicle	Lipid-binding domain, MV-specific after two-cell stage	Beer et al. (2019)	
Germ cells						
Spermatids and spermatozoa	MSP	Immunofluorescence, Immuno-EM	Double-membraned microvesicle	Cytosolic cytoskeletal protein	Kosinski et al. (2005)	
Spermatocytes	LifeAct::FP	Fluorescence	Residual Body	F-actin-binding domain	Huang et al. (2012)	
Ectodermal						
Epidermis	VHA-5::FP	Fluorescence, Immuno-EM	Exosome	Transmembrane protein	Liegeois (2006)	
Epidermis	FP::WRT-2/8	Fluorescence, Immuno-EM	Exosome	Lipid-modified secreted protein	Liegeois (2006)	
Male-specific EV-releasing neurons	PKD-2::FP	Fluorescence	Microvesicle	Transmembrane protein	Wang et al. (2014)	
Male-specific EV-releasing neurons	LOV-1::FP	Fluorescence, Immuno-EM	Microvesicle	Transmembrane protein	Wang et al. (2014), Walsh et al. (2022)	
Male-specific EV-releasing neurons	MCM-3::FP	Fluorescence	Microvesicle	Cytosolic and nuclear protein	Nikonorova et al. (2022)	
Male-specific EV-releasing neurons	SID-2::FP	Fluorescence	Microvesicle	Transmembrane protein	Nikonorova et al. (2022)	
IL2, Male-specific EV-releasing neurons	CWP-1::FP	Fluorescence	Microvesicle	Cytosolic protein	Wang et al. (2014)	
IL2, Male-specific EV-releasing neurons	DDN-3::FP	Fluorescence	Microvesicle	Secreted or transmembrane protein	Wang et al. (2015)	
IL2, Male-specific EV-releasing neurons	CIL-7::FP	Fluorescence	Microvesicle	Myristoylated protein	Maguire et al. (2015)	
IL2, Male-specific EV-releasing neurons	TTLL-11b::FP	Fluorescence	Microvesicle	Cytosolic protein	O’Hagan et al. (2017)	
IL2, Male-specific EV-releasing neurons	ENPP-1::FP	Fluorescence	Microvesicle	Transmembrane protein	Nikonorova et al. (2022)	
IL2	ASIC-2::FP	Fluorescence	Microvesicle	Transmembrane protein	Wang et al. (2015)	
Ciliated neurons	FP	Fluorescence	Microvesicle	Cytosolic reporters	Razzauti and Laurent (2021)	
Ciliated neurons	TSP-6::FP	Fluorescence	Microvesicle	Transmembrane protein	Razzauti and Laurent (2021), Nikonorova et al. (2022)	
Ciliated neurons	GCY-22::FP	Fluorescence	Microvesicle	Transmembrane protein	Razzauti and Laurent (2021)	
Ciliated neurons	CHLM-1::FP	Fluorescence	Microvesicle	Transmembrane protein	Clupper et al. (2022)	
Ciliated neurons with microvilli	GCY-8::FP	Fluorescence	Microvesicle	Transmembrane protein	Raiders et al. (2021)	
Ciliated neurons with microvilli	GCY-18::FP	Fluorescence	Microvesicle	Transmembrane protein	Raiders et al. (2021)	
Ciliated neurons with microvilli	GCY-23::FP	Fluorescence	Microvesicle	Transmembrane protein	Raiders et al. (2021)	
Ciliated neurons with microvilli	SRTX-1::FP	Fluorescence	Microvesicle	Transmembrane protein	Raiders et al. (2021)	
Touch neurons	mCherry	Fluorescence	Exopher	Aggregation-prone cytosolic reporter	Melentijevic et al. (2017)	
Mesodermal						
Body wall muscle	myo-3::
RPN-5::FP	Fluorescence	Exopher	Proteasome subunit	Turek et al. (2021)	
Body wall muscle	myo-3::
PAS-7::FP	Fluorescence	Exopher	Proteasome subunit	Turek et al. (2021)	
Body wall muscle	VIT-2::FP	Fluorescence	Exopher	Vitellogenin yolk protein	Turek et al. (2021)	
Dying cells						
Apoptotic embryonic cells	sAnxV::FP	Immuno-EM	Microvesicle	Secreted phosphatidylserine-binding reporter	Mapes et al. (2012)	
Apoptotic embryonic cells	sFP::LactC1C2	Immuno-EM	Microvesicle	Secreted phosphatidylserine-binding reporter	Mapes et al. (2012)	
Note that overexpression of reporter proteins can alter EV size and frequency (Razzauti and Laurent 2021).

MV formation from the plasma membrane

MVs are also known as ectosomes because MVs are produced by ectocytosis, a process in which the plasma membrane buds cellular content away from the rest of the cytoplasm to form vesicles in the extracellular space (van Niel et al. 2022; Dixson et al. 2023). In humans, MVs modulate important processes like coagulation, inflammation, cancer, and the immune response (Hu et al. 2023; Kalluri and McAndrews 2023). In C. elegans, embryonic blastomeres shed MVs that are confined by the eggshell or between cells. Embryonic MVs can be labeled with fluorescent reporter proteins that localize to the plasma membrane (Table 2), providing a useful system for observing MV biogenesis and uptake in live cells (Fig. 3, Wehman et al. 2011; Beer et al. 2018).

Fig. 3. Biogenesis of MV EVs in embryos. MVs are formed by ectocytosis, the progressive budding, and scission of the plasma membrane. TAT-5 is a PE flippase that transports PE lipids to the inner leaflet of the plasma membrane to maintain lipid asymmetry. TAT-5 is activated by PAD-1 to maintain membrane homeostasis. During ectocytosis, other proteins cause the exposure of cone-shaped PE, which may contribute to the curvature necessary for budding. The ESCRT machinery is recruited to the plasma membrane to create the bud and release the MV.

The mechanisms of MV formation from the plasma membrane are less well studied than ILV formation from endosomal membranes, but there are commonalities between the two topologically equivalent budding events. ESCRT subunits can be recruited to the plasma membrane and promote MV biogenesis in C. elegans embryos (Wehman et al. 2011). The ESCRT machinery is thought to bud the membrane away from the cytoplasm (Fig. 3), similar to ILV formation from endosomal membranes (Fig. 2). In addition, the small GTPase RAB-11 can promote MV biogenesis and was also identified in the alae screen for exosome mutants (Wehman et al. 2011; Hyenne et al. 2015). RAB-11 regulates endosomal recycling (reviewed in Sato, 2014b), but the specific role of RAB-11 in either exosome or MV biogenesis is unclear. Thus, MV budding and ILV budding share some biogenesis mechanisms in common and it will be important to determine how RAB-11 and ESCRTs are regulated to promote EV budding from different organelles.

There are also mechanistic differences between MV budding and ILV budding. A loss of plasma membrane phosphatidylethanolamine (PE) asymmetry consistently correlates with increased MV release without altering the number of ILVs in MVBs (Wehman et al. 2011; Beer et al. 2018). PE is normally maintained in the cytosolic face of the plasma membrane by the P4-ATPase TAT-5, which functions as an aminophospholipid translocase to flip PE from the exofacial leaflet to the cytofacial leaflet of the membrane bilayer and create lipid asymmetry (Fig. 3). TAT-5 is activated by the Dopey homolog PAD-1, and their essential interaction is conserved from yeast to mammals (Barbosa et al. 2010; McGough et al. 2018). In tat-5 or pad-1 mutant germ cells and embryos, PE is exposed on the outer leaflet of the plasma membrane and 100–400-nm spherical or tubular MVs accumulate between cells or along the embryo surface. Excessive MV biogenesis disrupts gastrulation movements and cell shape, leading to embryonic lethality and sterility in strong loss-of-function tat-5 or pad-1 mutants (Guipponi et al. 2000; Wehman et al. 2011). Excessive EV biogenesis also occurred after the human TAT-5 homolog ATP9A was depleted in cell lines (Naik et al. 2019; Xu et al. 2020), suggesting that TAT-5 and ATP9A have conserved roles inhibiting EV release.

Which proteins or processes act in opposition to TAT-5 to expose PE and initiate MV release are currently unknown. Likely candidates for PE exposure include lipid scramblases that act as channels to destroy lipid asymmetry (Fig. 3). Indeed, the lipid scramblase TMEM16F regulates inducible MV release from platelets (Fujii et al. 2015). Scott syndrome is caused by mutations in TMEM16F (Suzuki et al. 2010), which is characterized by a lack of induced MV release and defects in blood coagulation (Zwaal et al. 2004). However, scramblases alter the asymmetry of multiple lipids, consistent with the observed exposure of both PE and phosphatidylserine (PS) in gain-of-function TMEM16F mutant cells (Suzuki et al. 2010). Consistently, a C. elegans homolog of TMEM16F, ANOH-1, promotes PS exposure in dying neurons (Li et al. 2015), but its effect on PE has not been examined. Therefore, it will be important to determine which scramblases are activated to promote MV budding or whether there are other cellular processes capable of disrupting lipid asymmetry.

The effect of lipid symmetry on MV budding seems specific to PE, because PS asymmetry maintained by the TAT-1 flippase influences large EV uptake, not MV budding (Fazeli et al. 2020). PE asymmetry and MV biogenesis are indirectly regulated by endosomal recycling regulators that traffic TAT-5 back to the plasma membrane, such as PI3Kinase VPS-34, Beclin1 homolog BEC-1, DnaJ protein RME-8, and sorting nexins SNX-1/6 and SNX-3 (Beer et al. 2018). However, it is unclear whether these recycling proteins also traffic lipid scramblases. Furthermore, it remains to be determined which biophysical properties of PE lipids influence MV budding, such as conical shape, neutral charge, abundance, or saturation. It is also unclear how PE symmetry leads to ESCRT recruitment to the plasma membrane (Fig. 3), as ESCRT subunits bind phosphatidylinositol (PI) lipids (Katzmann et al. 2003).

During spermatogenesis, spermatids also release 0.5–1 µm MVs (Liu et al. 2023). These large EVs are formed from plasma membrane buds and induced by protease signaling pathways feeding into the tyrosine kinase SPE-8. In spermatids, MV formation depends on actin filaments and the myosin VI motor SPE-15. The actin dependence of MV formation was surprising, given the apparent depletion of actin from spermatids that occurs during residual body formation (reviewed in L’Hernault 2006). However, actin dynamics have previously been linked to MV biogenesis from the plasma membrane of chondrocytes (Hale and Wuthier 1987), suggesting that actin dynamics may play conserved roles in MV formation.

MV formation from the cell body of mechanosensory touch neurons has been linked to regulation of the fusogen EFF-1 by the small GTPase RAB-5 (Linton et al. 2019). EFF-1 is homologous to viral class-II fusion proteins and remodels membranes for fusion events (Mohler et al. 2002). Intriguingly, transfection of worm EFF-1 into hamster kidney cells led to EV shedding into the culture media (Zeev-Ben-Mordehai et al. 2014). MVs carrying EFF-1 can also be released from the soma of PLM neurons after the expression of a dominant-negative RAB-5 mutant (Linton et al. 2019). Disrupting GTP exchange on RAB-5 caused EFF-1 accumulation in the plasma membrane and increased membrane protrusion and EV shedding, suggesting that RAB-5 promotes EFF-1 endocytosis on the plasma membrane to maintain EFF-1 in intracellular endosomes and prevent fusogen-mediated EV budding. However, it is unclear how overexpression or mislocalization of EFF-1 caused the plasma membrane to protrude and induce EV budding, as EFF-1 was the EV cargo being studied.

EV formation from membrane protrusions

EVs can be formed from specialized organelles at the plasma membrane, which depends on organelle formation and structure. Both stable protrusions like cilia and dynamic protrusions like filopodia are common sites of EV production (D’Angelo et al. 2023).

MVs from cilia

Cilia are microtubule-based membrane protrusions that serve as conserved sites for EV shedding from unicellular and multicellular organisms (Ojeda Naharros and Nachury 2022, Ma et al. 2024 ). Cilia can form EVs from the ciliary tip, base, and along the ciliary membrane, ranging from whole cilia shedding at the transition zone (Gogendeau et al. 2020; Mirvis et al. 2019), to ciliary tip decapitation or ectocytosis (Nager et al. 2017; Phua et al. 2017), and the formation of nanotubes from the ciliary membrane that later bead into EVs (Szempruch et al. 2016).

In C. elegans, cilia are located at the dendritic ends of 60 sex-shared and 52 male-specific sensory neurons (Fig. 4a, reviewed in Inglis et al. 2007). The main sensory organs, the amphid and phasmid cilia, release EVs from both the ciliary tip and base in hermaphrodites (Razzauti and Laurent 2021). Furthermore, a set of six IL2 and 21 male-specific sensory neurons have been specially classified as EV-releasing neurons (EVNs) due to their prolific production of 50–200-nm ciliary EVs (Wang et al. 2014). Electron tomography data showed ciliary EVs budding from the ciliary membrane (Wang et al. 2014), and multivesicular bodies have not been observed inside C. elegans cilia by TEM. Therefore, ciliary EVs are MVs derived from the cell membrane and not exosomes derived from internal membranes. Ciliary EVs can be visualized using ciliary membrane proteins or cytosolic proteins that localize in cilia and become EV cargos. Fluorescent tagging of ciliary EV cargo (Table 2) allows the observation of ciliary EVs in and around intact, live animals.

Fig. 4. EV biogenesis from cilia. a) Sensory cilia release EVs from both the ciliary tip and base. The ciliary transport system promotes EV release at the tip while inhibiting EV release at the base. EVs released from the ciliary base can be taken up by neighboring amphid glia. b) Male-specific EVNs utilize cell type-specific ciliary kinesin (KLP-6) to traffic cargos to the tip for sorting into EVs. KLP-6 and the tubulin TBA-6, along with the post-translational modification enzyme polyglutamylase TTLL-11 and its counteracting enzyme, CCPP-1, specialize the axoneme to facilitate abundant EV release.

The release of EVs from the ciliary tip relies on intraflagellar transport (IFT) (Wang et al. 2014; Clupper et al. 2022). IFT is important for ciliary structure and the dynamic localization of ciliary proteins (reviewed in Inglis et al. 2007). Anterograde transport of IFT complexes toward the ciliary tip is driven by ciliary kinesins, while retrograde transport away from the ciliary tip is driven by dynein motor proteins (Fig. 4). IFT complex components important for ciliogenesis, such as DAF-10 and OSM-5, are required for EV release from the ciliary tip (Bae et al. 2006; Wang et al. 2014). In IFT mutants, ciliary proteins accumulate along the ciliary membrane but are not released on EVs, suggesting that the enrichment of EV cargo at the ciliary tip is essential for ciliary EV release (Wang et al. 2021). In C. elegans cilia, anterograde IFT is driven by two pan-ciliary kinesin-2 complexes (Fig. 4a): the homodimeric kinesin OSM-3 and the heterotrimeric kinesin comprised of KLP-11, KLP-20, and KAP-1 (Ou et al. 2005). The homodimeric and the heterotrimeric ciliary kinesins act redundantly in ciliary tip EV release (Wang et al. 2014; Clupper et al. 2022). Furthermore, the CHE-3 dynein heavy chain is essential for the release of ciliary tip EVs (Wicks et al. 2000; Wang et al. 2014), demonstrating the importance of both anterograde and retrograde IFT for EV release from the ciliary tip. However, EV release independent of the redundant kinesin-2 complexes has been observed for an EV cargo that does not localize to the tips of cilia (Clupper et al. 2022), suggesting the existence of molecularly distinct biogenesis mechanisms for EVs derived from different parts of the cilium.

EVN cilia extend their tips out of the cuticular pore, which allows EVs shed from the ciliary tip to be released into the worm's environment. TEM analysis also revealed EVs in the lumen of labial and cephalic sensilla (Doroquez et al. 2014; Wang et al. 2014), indicating that EVNs also release abundant EVs from the base or along the cilium. Furthermore, EVs from the ciliary tip and base are released in an antagonistic manner; disrupting the release of ciliary tip EVs causes an excessive release of ciliary base EVs (Wang et al. 2014; Maguire et al. 2015; Wang et al. 2021).

EVN cilia are specialized in microtubule structure, post-translational modifications, and motor proteins that support EV shedding from the ciliary tip. For example, the centrosome-derived, 9-fold symmetric microtubule core of cilia known as the axoneme differs in cephalic male (CEM) cilia. The doublet microtubules splay and fuse to create a curved axoneme, which requires a unique alpha tubulin TBA-6 (Silva et al. 2017). In tba-6 mutants, half of the CEM cilia have inward-curved tips instead of extending out the ciliary pore and produce excessive small EVs alongside the cilia. Similarly, environmental ciliary EV release requires the long isoform of the polyglutamylation enzyme TTLL-11b, which post-translationally modifies microtubule axonemes and is only expressed in EVNs (O'Hagan et al. 2017). The counteracting enzyme responsible for deglutamylation of the microtubules, CCPP-1, has cell-specific regulation in the EVNs and is required for efficient environmental EV shedding. Furthermore, the EVN-specific kinesin-3 protein, KLP-6, and the myristoylated coiled-coil domain protein, CIL-7, further enrich ciliary EV cargos at the ciliary tip for environmental EV release (Wang et al. 2014; Maguire et al. 2015; Wang et al. 2021).

Amphid and phasmid cilia release fewer EVs from the ciliary tip and ciliary base than EVNs, and it is untested whether all amphid and phasmid cilia release EVs. EVs were not observed in the lumen of the amphid channel in adult hermaphrodites by TEM (Doroquez et al. 2014), likely due to EV uptake by the surrounding glia (Razzauti and Laurent 2021). EV shedding from amphid and phasmid cilia can be increased by disrupting ciliary transport through ciliary gene mutations, overexpression of ciliary proteins, or exposure to lipophilic dyes (O'Hagan et al. 2017; Akella et al. 2020; Razzauti and Laurent 2021; Nikonorova et al. 2022). Notably, amphid and phasmid ciliary EVs are taken up by the surrounding glia in a process that depends on ATP production (Ohkura and Bürglin 2011; Razzauti and Laurent 2021), providing an in vivo system to observe the dynamics of EV uptake.

MVs pruned from neuronal microvilli

Caenorhabditis elegans sensory neurons can also shed EVs from other protrusions, including actin-based microvilli. Adult AFD microvilli shed 0.5 µm EVs during phagocytosis by AMsh glial sheath cells (Raiders et al. 2021). The glial cell responds to decreases in neuronal activity by pruning neuronal microvilli by phagocytosis. PS exposure on microvillar membranes is recognized by glial cell using classic phagocytic pathways, including PS-binding TTR-52, PS receptor PSR-1, integrin PAT-2, and the CED-2/5/12 pathway that culminates in the actin-regulating Rac GTPase CED-10 (Fig. 5). The activity-dependent mechanisms that regulate PS exposure specifically on microvilli remain to be determined, but at least one lipid scramblase, SCRM-1, helps promote PS exposure. If PS exposure is not restricted to microvilli, excessive pruning occurs, such as in the PS flippase mutant, tat-1. Thus, organelles can also be fragmented to form EVs through interactions with neighboring cells.

Fig. 5. EV biogenesis from microvilli. Glial phagocytosis regulates the sensitivity of AFD neurons by engulfing the ends of AFD microvilli as EVs. This process involves the glial sheath recognizing “eat-me”? signals on microvilli, such as PS lipids (red PS) after their exposure by lipid scramblases, including SCRM-1. The PS flippase TAT-1 maintains the normal asymmetry of PS lipids in the rest of the cell. PS exposure is recognized by phagocytic pathways for the selective pruning of microvilli to become EVs.

Double-membrane EVs from protrusions

Not all EVs are wrapped in a single membrane bilayer. In C. elegans, spermatids and spermatozoa produce small EVs with two membrane bilayers (Fig. 6, Kosinski et al. 2005). The outer membrane of these EVs appears scalloped after high-pressure freezing and TEM, characterized by a wavy surface. The major sperm protein (MSP) is sandwiched between the two membranes, while the inner core appears less electron dense. MSP is a cytoskeletal protein predominantly found inside extensions like the pseudopod of ameboid sperm (reviewed in L’Hernault 2006). This led to the proposal that membrane protrusions would extend around extracellular material and the folding back of the protrusions could seal themselves to release EVs with two membranes and a hollow core (Fig. 6, Kosinski et al. 2005). Given the advances in microscopy over the past two decades, it would be interesting to revisit these unusual EVs with super-resolution or tomographic approaches to determine whether the double-membrane EVs are donut-shaped toroids or concentric spheres to better understand the diversity of EV shapes.

Fig. 6. Biogenesis of double membrane EVs in sperm. The main cytoskeletal protein MSP (star) drives the formation of membrane protrusions in spermatids and spermatozoa. Protrusions can fold back to form double-membrane vesicles (adapted from Kosinski et al. 2005).

EV formation involving contractile rings

Large EVs from mitotic cell division

Micron-sized EVs called midbody remnants are formed at the end of cytokinesis after dividing animal cells separate (Skop et al. 2004; Elia et al. 2011; Chai et al. 2012; Crowell et al. 2014). Cytokinesis begins with furrow ingression, where an actomyosin ring contracts to pull the plasma membrane toward the spindle midbody (Fig. 7a). Ingression creates a transient organelle that forms a thin connection between the cells termed the intercellular bridge. This bridge is then constricted on both sides of the midbody, with one side sealing first to complete abscission and separate the two cells. Both sides of the bridge soon seal to release the midbody remnant as a large EV. The cargo of midbody remnants is enriched in spindle midbody proteins, actin, and non-muscle myosin, making these large EVs relatively easy to label and track.

Fig. 7. Large EV biogenesis involving contractile rings. a) During cell division, an actomyosin ring contracts to pull the plasma membrane between the two cells toward the spindle midbody. Cytokinesis forms an intercellular bridge, which undergoes active constriction and membrane remodeling to release the midbody remnant as an EV. b) During meiosis II, spermatocytes use two types of myosin motors to polarize their contents and bud off from a shared cytoplasm, resulting in the release of a large EV known as a residual body. c) Mid-embryogenesis, primordial germ cells reduce their cell volume in half using an actomyosin ring to form a large lobe. The lobe is engulfed by the neighboring endodermal cell to non-autonomously form a large EV.

Caenorhabditis elegans midbody remnants were first reported in the context of larval Q cells (Chai et al. 2012). Midbody remnants from the Q neuroblasts were labeled with centralspindlin ZEN-4 reporters and engulfed by the neighboring epidermal cell hyp7, indicating that the midbody remnants had been released extracellularly as EVs. The Q midbody remnants exposed the lipid PS on the surface of their membrane to signal for phagocytosis by hyp7. Q midbody remnant uptake depended on the cell death (CED) engulfment pathways (Chai et al. 2012), which were identified for regulating cell corpse clearance by phagocytosis but regulate many physiological engulfment events (reviewed in Conradt et al. 2016; Ghose and Wehman 2021).

Given their relatively large cell size, early embryos are particularly useful for dissecting the mechanisms of midbody remnant biogenesis. Midbody remnants can be labeled with reporters that accumulate in the spindle midbody like ZEN-4, reporters that become enriched in the contractile ring like non-muscle myosin NMY-2, or ESCRT reporters that accumulate in the bridge (Fig. 7a, Table 2). After furrow ingression, microtubules are disassembled from the midbody and septins create a cytoplasmic barrier in the intercellular bridge (Green et al. 2013; König et al. 2017). The ESCRT machinery accumulates in the bridge and pulls the membranes closer together (Fig. 7a), similar to ILV and MV formation (Figs. 2 and 3). Membrane fusion on one side of the bridge leads to abscission (König et al. 2017), which allows the membranes to physically separate the two cells (Fig. 7a). This is quickly followed by a second membrane fusion event on the other side of the bridge to release the midbody remnant as a ∼1-µm EV. The dynamin DYN-1 also promotes abscission for midbody remnant release (König et al. 2017). Thus, midbody remnant biogenesis depends on furrow ingression and mechanisms involved in ILV and MV budding but on a larger scale.

Early C. elegans embryos are also excellent for tracing the release and uptake of midbody remnant vesicles (Ou et al. 2014; Singh and Pohl 2014; Fazeli et al. 2016; König et al. 2017; Bai et al. 2020), given the stereotyped pattern of embryonic cell divisions (reviewed in Gönczy and Rose 2005). Embryonic midbody remnants can be taken up by one of the daughter cells that produced the remnant or by a neighboring cell (Ou et al. 2014; Singh and Pohl 2014; Fazeli et al. 2016; Bai et al. 2020). Midbody remnant uptake uses CED corpse engulfment pathways in early embryos (Ou et al. 2014; Fazeli et al. 2016), similar to Q midbody remnant uptake by hyp7 (Chai et al. 2012).

Large EVs from meiotic cell division

Germ cells also use cell division machinery to release large EVs. In C. elegans, spermatocytes use non-muscle myosin NMY-2 and actin to form a pseudocleavage furrow that partially separates the future spermatids during meiosis II (Fig. 7b) (Hu et al. 2019). This syncytium uses actomyosin motors to grow an anuclear region in its center with distinct cargo from the spermatids. Then, the myosin VI motor SPE-15 uses its cargo adaptors GIPC-1 and GIPC2 to complete cytokinesis and separate the four spermatids from the anuclear region, forming a ∼4-µm EV known as a residual body. Actin and myosin VI/GIPC-mediated cleavage are essential for residual body formation, but residual bodies still form after the loss of NMY-2-mediated pseudocleavage. Thus, multiple types of actomyosin contractile rings can contribute to EV formation.

Large EVs from pseudocleavage and phagocytosis

Mid-embryogenesis, primordial germ cells use pseudocleavage to form a lobe and phagocytic signaling to reduce cell size and organelle content in large EVs (Fig. 7c) (Abdu et al. 2016; Maniscalco et al. 2020). In bean stage embryos, the primordial germ cells Z2 and Z3 elongate and NMY-2 forms a pseudocleavage furrow around the middle of the cells along with the anillin ANI-1 and septin UNC-59 (Maniscalco et al. 2020). Actin polymerization regulated by the formin CYK-1 is also required for lobe furrow ingression to start separating the lobe from the rest of the cell. Actin polymerization is activated by the small GTPase RHO-1 and its GEF ECT-2. However, unlike cytokinesis, components of the centralspindlin complex are not required for lobe formation, consistent with the absence of a spindle midbody. Instead, a novel protein NOP-1 is thought to activate ECT-2 after nuclear migration to allow anuclear lobe formation. However, NMY-2 is also required to keep the contractile ring from releasing the lobe as an EV. Instead, the lobe is released as an EV after engulfment by neighboring intestinal precursors, as the lobes fail to detach in endoderm-less mutants (Abdu et al. 2016). The small Rac1-like GTPase CED-10 promotes actin polymerization in the endoderm for lobe engulfment, but unlike most phagocytic events, CED-10 was not activated by the CED-1/6/7 or CED-2/5/12 pathways. However, other factors required for apoptotic corpse clearance, namely the SNX9-like sorting nexin LST-4 and its binding partner dynamin DYN-1 (Kinchen et al. 2008), are required for lobe engulfment (Abdu et al. 2016). Thus, non-muscle-myosin-mediated contractile rings initiate several types of large EV formation, but scission to release the EVs is completed by other intrinsic or extrinsic factors (Fig. 7).

EV formation from stressed or dying cells

Exophers from stressed cells

Stressed neurons and muscle cells in C. elegans and mammals can produce large extensions containing organelles that are cast off as large EVs, including exophers (Fig. 8a, Melentijevic et al. 2017; Nicolás-Ávila et al. 2020; Turek et al. 2021). Caenorhabditis elegans neurons, including the mechanosensory touch receptor neurons, produce exophers ranging from 1 to 8 µm in diameter, sometimes larger than the neuronal cell body (5–8 µm) (Melentijevic et al. 2017). Exophers first appear as an outward bud of the cell membrane and are not formed by reentry into the cell cycle, as exophers form independent of nuclear replication. The budding exopher later constricts to be attached only by a thin nanotube (Fig. 8b); however, it is unknown whether contractile rings are involved in exopher constriction.

Fig. 8. Caenorhabditis elegans extrudes exophers from stressed neurons and body wall muscle cells. a) Adult hermaphrodites release exophers from stressed neurons, with ALMR neurons releasing the most. Under starved conditions, body wall muscles release exophers to feed oocytes. b) Stressed neurons accumulate protein aggregates in aggresomes near the nucleus and dysfunctional organelles, including mitochondria. Exopher formation begins with membrane bud formation, followed by enlargement and constriction. Aggresomes caged by intermediate filaments promote exopher formation. The exopher remains connected to the cell body by a nanotube, which facilitates exopher growth. Exophers are released as EVs after phagocytosis by a neighboring epidermal cell. Mito, mitochondria; Agg, aggresome.

Organismal and cellular stress appears to be major regulatory stimuli for exopher biogenesis, as starvation, osmotic stress, and oxidative stress can increase exopher production (Cooper et al. 2021; Turek et al. 2021). Disrupting macroautophagy, proteasomal degradation, or the mitochondrial unfolded protein response increases the number of exophers, indicating that proteostress upregulates exophergenesis (Melentijevic et al. 2017). However, while most autophagy proteins reduce the cellular need for exophers, ATG-16.2 promotes exopher production (Yang et al. 2024). Atg16 homologs lipidate Atg8/LC3 family proteins to attach the protein to phagophore membranes during macroautophagy and to endosomes and phagosomes during non-canonical forms of autophagy (reviewed in Durgan and Florey 2022). The non-canonical role of ATG-16.2, dependent on its C-terminal WD40 domain, promotes exopher production (Yang et al. 2024), suggesting that a non-canonical form of autophagy promotes exophergenesis.

Exophers contain organelles, such as Golgi, lysosomes, ER, and mitochondria, as well as protein aggregates. Neurotoxic proteins, such as the polyglutamine expansion protein huntingtin Q128 or the amyloid-forming human Alzheimer's disease fragment Aβ1–42, accumulate in exophers and increase their formation. Exopher formation can even be induced by the red fluorescent reporter protein mCherry (Melentijevic et al. 2017), which can form aggregates (Shemiakina et al. 2012). These observations suggest that exopher formation is driven by cell stress-inducing cargos.

Different cargos engage distinct organelles for extrusion in exophers. Cargo proteins such as Q128 first aggregate and coalesce to form aggresomes near the nucleus (Fig. 8b). Q128 trafficking into exophers depends on intermediate filament proteins IFD-1 and IFD-2, which associate with aggresomes, as well as adaptors for dynein microtubule motors (Arnold et al. 2023). In contrast, mCherry is concentrated into LMP-1-positive late endosomes or lysosomes, expanding the endolysosome. Through an unknown mechanism, expanded endolysosomes and aggresomes are then extruded into an exopher. Although the connection between the cell body and exopher thins to a nanotube, additional cargo continues to be trafficked into the growing exopher (Fig. 8b), including calcium and protein aggregates (Melentijevic et al. 2017).

Exopher release depends on actin and Arp2/3-dependent engulfment by a neighboring epidermal cell (Wang, Arnold, et al. 2023). Indeed, phagocytic receptors and membrane trafficking regulators implicated in phagocytic engulfment, including the receptor CED-1, the small GTPase RAB-35, and the Arf GAP CNT-1, are required in the epidermis for exopher engulfment through inhibition of the small GTPase ARF-6. ARF-6 effectors like the exocyst and lipid kinase PPK-1 also regulate exopher engulfment, likely through their roles in bringing additional membrane and protein regulators to the growing phagocytic cup. If engulfment is disrupted in the epidermis, exophers often fuse back with the neuronal cell body (Wang, Arnold, et al. 2023), emphasizing the importance of the neighboring cell for exopher release.

Apoptotic EVs from dying cells

Apoptotic cells create multiple types of EVs as the cell dies, including small EVs and large apoptotic bodies, only the latter of which are visible with light microscopy (Santavanond et al. 2021). In C. elegans, 50–150-nm EVs were observed around embryonic cell corpses using EM (Mapes et al. 2012). Corpse EVs had the lipid PS exposed on the outer leaflet of the lipid bilayer, similar to the plasma membrane of apoptotic corpses. Both the ATP-binding cassette (ABC) transporter CED-7 and the PS-binding transthyretin TTR-52 promoted EV release from embryonic apoptotic corpses. A related ABC transporter, ABCA1, promotes PS translocation across membrane bilayers and is required for MV release from apoptotic cells in mammals (Hamon et al. 2000). Therefore, ABCA1 and CED-7 are hypothesized to directly cause PS exposure as bidirectional lipid scramblases or unidirectional lipid floppases. ABCA1 also alters the localization of cholesterol and sphingomyelin and changes lipid packing within the plasma membrane (Landry et al. 2006), which may indicate a broader lipid specificity than PS. In C. elegans, PS exposure did not increase EV release from viable embryonic cells in tat-1 flippase mutants (Fazeli et al. 2020), suggesting that CED-7 may also alter another lipid or protein to regulate EV release from apoptotic cells in embryos. Secreted TTR-52 binds to the surface of apoptotic cells and EVs (Mapes et al. 2012), but how TTR-52 increases EV release requires further investigation.

Open question: how is cargo sorted into EVs?

The content of EVs is not identical to cell content; specific proteins, RNAs, and lipids are selectively incorporated into EVs during EV biogenesis. Indeed, proteins involved in EV biogenesis like ESCRT subunits are enriched in EVs (Nikonorova et al. 2022). Cargos that contribute to EV functions are also sorted, such as protein aggregates specifically enriched in neuronal exophers compared to the cell body to promote their secretion in EVs (Melentijevic et al. 2017). Specific cargos can also be excluded from EVs during EV biogenesis. Abundant cortical proteins like clathrin can be selectively excluded from MVs to prevent their secretion in EVs (Beer et al. 2019). The sorting of cargos is also specific to EV subtypes. Myristoylated plasma membrane markers and cytosolic reporters can be specifically excluded from RAB-5DN touch neuron EVs (Linton et al. 2019), while being preferentially included in touch neuron exophers and ciliary EVs (Maguire et al. 2015; Melentijevic et al. 2017). How C. elegans cells regulate selective sorting events on endolysosomal and plasma membranes is unclear, but membranes are thought to act as a platform for the assembly and sorting of EV cargo into EVs.

Cargo sorting into ILVs for release in exosomes

Cargo sorting into ILVs for degradation often occurs via ubiquitin-mediated sorting. The E2 ubiquitin ligases UBC-13 and UEV-1 promote the polyubiquitination and sorting of cargo proteins to ILVs for degradation inside lysosomes (Sato, Konuma, et al. 2014). Similarly, ESCRT proteins recognize and cluster ubiquitinated cargo for sorting into ILVs (Frankel et al. 2017). Although ESCRTs are used to form both secretory and degradative MVBs, it is unclear how often ubiquitination is used to sort cargos into exosomes.

Non-ubiquitinated cargos can also be sorted into ILVs for secretion in EVs, and several ILV budding pathways have been characterized in mammalian systems. The syntenin-Alix pathway uses the PDZ domain protein syntenin to connect ESCRT-mediated ILV budding to sorting non-ubiquitinated cargos into exosomes (Baietti et al. 2012). Neutral sphingomyelinases hydrolyze the lipid sphingomyelin to produce conical ceramide lipids for cargo sorting into ILVs and exosome biogenesis (Trajkovic et al. 2008). Some tetraspanins (e.g. CD63, CD81) can form membrane nanodomains that sort cargos into ILVs independent of ESCRT pathways (van Niel et al. 2011).

Cargo sorting into cilia for release in MVs

Specialized projections like cilia can also be used to sort EV cargo. Cilia serve as sensory organelles for the cell, housing signal transduction modules that are segregated from the rest of the cell (Mohieldin et al. 2021). Asymmetrical localization of membrane proteins along the cilium may serve as a scaffold to presort ciliary EV cargo. The conserved ciliary EV cargo PKD-2 is localized in two rows on opposite sides of the Chlamydomonas ciliary membrane and the dorsal side of mouse nodal cilia (Liu et al. 2020; Katoh et al. 2023). In C. elegans, PKD-2 and CIL-7 are sequentially enriched at the distal tip of EVN cilia, a process coordinated by three ciliary kinesins: the kinesin-3 protein KLP-6 and two pan-ciliary kinesin-2 motors, the homodimeric OSM-3 kinesin and the heterotrimeric kinesin of KLP-11, KLP-20, and KAP-1(Wang et al. 2021). Another ciliary EV cargo CLHM-1 is localized to the proximal ciliary membrane independently of the pan-ciliary kinesins (Clupper et al. 2022). Thus, EVNs engage multiple kinesin motors to temporally and spatially separate ciliary EV cargos, allowing EVNs to shed distinct EV subtypes from a single cilium. When exposed to hermaphrodites, male EVN cilia can increase the ratio of PKD-2 EVs to CIL-7 or CLHM-1 EVs (Wang et al. 2021; Clupper et al. 2022), demonstrating that sensory signals can regulate the production of EV subtypes by altering EV biogenesis or cargo sorting.

Open question: can we specifically control EV formation?

EV biogenesis involves a complex interplay of genetic, metabolic, and environmental factors. This complexity raises pivotal questions about the specificity and universality of EV production mechanisms. Are there conserved pathways across different cell types and organisms that can be targeted to better understand EV functions or to apply therapeutic interventions? Our current understanding from C. elegans suggests that cells produce EVs in response to a range of stimuli, with each EV subtype containing distinct cargoes and utilizing a mix of overlapping and distinct molecular machinery. Unraveling EV biogenesis mechanisms may enable the manipulation of EV cargo loading and production, marking a new era in biomedical research and therapeutic strategies.

Some trends in EV regulation suggest commonalities across disparate types of EV biogenesis. Lipid pathways play a key role in EV regulation, suggesting that alterations to biophysical properties of the membrane bilayer could broadly regulate EV formation. In mammalian cells, EV biogenesis is promoted after the creation of high-curvature lipid species by reducing the size of lipid headgroups. PLD1 converts the headgroup of PC lipids to smaller PA lipids, and neutral sphingomyelinases convert sphingomyelins to smaller ceramides on the cytosolic face of the endosome membrane to promote ILV budding (Trajkovic et al. 2008, Egea-Jimenez and Zimmermann 2018). However, decreasing the amount of PE in the cytosolic face of the plasma membrane correlates with increased EV production in C. elegans embryos with disrupted PE asymmetry (Wehman et al. 2011, Beer et al. 2018), making it difficult to draw conclusions on the effect of cytosolic small lipid headgroups on EV release. Meanwhile, lipids with large headgroups play signaling roles for EV uptake. PS exposure enhances phagocytosis-aided EV formation (Raiders et al. 2021, Wang, Arnold, et al. 2023). Investigating the fundamental role of lipids in EV regulation is key to comprehending the physiological roles of EVs.

Mechanical regulation of EV production is also significant. In cancer cells, stiff ECMs significantly enhance exosome secretion (Wu et al. 2023), underscoring the intricate relationship between mechanical forces and EV release. In C. elegans, mechanical pressure induces and sustains ciliary PKD-2 EV release from EVNs (Wang et al. 2020; Wang et al. 2024b) and may regulate exopher production from mechanosensory touch receptor neurons (Wang et al. 2024a). Imbalanced proteostasis drives exopher production (Melentijevic et al. 2017; Cooper et al. 2021), and proteostasis regulates and is regulated by cellular mechanics (Evers et al. 2021). Indeed, exopher production from both neurons and body wall muscles peaks when the uterus is expanded by embryos and is decreased when embryonic development is disrupted (Turek et al. 2021; Wang et al. 2024). Thus, biomechanics could be a driving force behind EV shedding, warranting further investigation.

What can EVs do?

EVs have important functions for the cell that releases the EV, for a nearby or distant cell that receives the EV, or for inter-organismal communication. Cells can release EVs to change the composition of their membrane, cytoplasm, or organelles. Packaging cargos in EVs allows the releasing cell to rapidly terminate signaling pathways, remove dysfunctional cell components, or outsource the slower degradation of EV cargos to other cells. EV-receiving cells can respond to the signals carried by EVs or benefit structurally or metabolically from EV cargo. The EV membrane provides a protective shell for its cargos compared to simple macromolecules, allowing EV-mediated functions to be long-range, occurring between distant tissues or different organisms, including different species. Below we highlight specific examples for different modes of EV function in C. elegans and beyond.

Functions for EV-releasing cells

EVs sculpt cell membranes to terminate cell division

Cells can use EVs to alter their shape. At the end of cell division, the intercellular bridge is enriched with proteins regulating the actin and microtubule cytoskeletons, endocytosis, and ectocytosis. To allow the cytoskeleton and membrane trafficking to return to homeostasis, the bridge is released as a large EV after the connections to both cells are constricted for scission (Fig. 9a). Notably, midbody remnants are not spherical by electron tomography, but have many protrusions and folds, leading to a complex structure (König et al. 2017; Fazeli et al. 2020). ESCRT-like spirals are seen in electron tomograms forming thin protrusions from the intercellular bridge, and small fragments of the bridge can be taken up before the large midbody remnant (König et al. 2017). These data suggest that ESCRT-dependent ectocytosis promotes the release of small MVs from the intercellular bridge, which may be used to further break up these micron-sized structures or to initiate phagocytic signaling for midbody remnant uptake. Thus, cells may use both large and small EVs to remodel their membranes to terminate the signaling pathways that regulate cell division. Severing both sides of the bridge could also provide redundancy to ensure cellularization before differentiation.

Fig. 9. Functional roles of EVs for EV-releasing cells. a) At the end of cell division, cells release the intercellular bridge full of cytoskeletal and membrane trafficking regulators as a large EV known as the midbody remnant (MBR). The bridge also uses the ESCRT machinery to bud off tubules that can be taken up by neighboring cells as small EVs. b) Sensory cilia release EVs from the ciliary tip and base to maintain the balance of ciliary proteins. c) Protease-activated spermatids bud mitochondria in EVs called mitophers to decrease their mitochondrial content as they differentiate into spermatozoa. d) Neurons concentrate toxic protein aggregates and dysfunctional organelles in exophers to remove them from the cell body and improve neuronal function.

EVs regulate sensory homeostasis and plasticity

Neuronal EV shedding plays a conserved role in regulating both ciliary proteostasis and structural stability of neuronal protrusions. In mammalian cells, primary cilia eliminate accumulating G protein-coupled receptors (GPCRs) in EVs via actin-mediated ciliary tip ectocytosis (Nager et al. 2017). In C. elegans, ciliary EV shedding facilitates signal transduction by removing excess proteins and membrane components at both the ciliary base and tip (Fig. 9b, Wang et al. 2014; Maguire et al. 2015; Wang et al. 2021; Razzauti and Laurent 2021). Ciliary EVs also contribute to the regulation of ciliary stability, being released during ciliary degeneration and resorption. The shedding of the ciliary tip as an EV hastens the removal of the cilium during cilia resorption in cultured mammalian cells (Phua et al. 2017). Septins are crucial for maintaining the structure of the ciliary tip, and defects in septins can lead to the release of ciliary EVs through ectocytosis (Kanamaru et al. 2022). In C. elegans, the carboxypeptidase CCPP-1 is essential for ciliary axoneme stability; mutants of ccpp-1 exhibit EVs in the amphid channel and cephalic sensillum alongside degenerating cilia (O’Hagan et al. 2011). The balanced shedding of EVs from both the ciliary tip and base, observed in EVN, amphid, and phasmid cilia, suggests that the shedding of ciliary EVs is a conserved mechanism for cilia to maintain membrane composition and structure (Ojeda Naharros and Nachury 2022; Ma et al. 2024). Moreover, modulation of neuronal sensitivity is achieved through EV formation during the glial pruning of sensory microvilli (Raiders et al. 2021). In summary, EV shedding maintains neuronal homeostasis and plasticity, regulating both the protein composition and sculpting the structure of cilia and microvilli.

EVs remodel organelle content during differentiation

Cells can use EVs to alter their organelle content, including during developmental processes like differentiation. At the end of meiosis II, C. elegans spermatocytes rearrange their contents to form haploid spermatids enriched in mitochondria and a large EV known as a residual body, which is anuclear and enriched in actin, tubulin, and ribosomes (Roberts et al. 1986; Kelleher et al. 2000). The myosin VI motor SPE-15 migrates cortical actin from spermatids toward the expanding residual body to reduce the actin content of spermatids, as well as to enrich tubulin and ribosomes in residual bodies (Kelleher et al. 2000; Hu et al. 2019). Simultaneously, the myosin II motor moves toward the spermatids and helps enrich mitochondria in spermatids. In nmy-2 mutants, more mitochondria remain in the residual body (Hu et al. 2019). Thus, these two myosin motors coordinate the movement of multiple organelles to create functional spermatids. Further study of asymmetric trafficking events between spermatids and residual bodies is likely to provide insights into mechanisms of EV cargo sorting.

At a later stage during spermatogenesis, spermatids decrease their mitochondrial quantity by one-third by rapidly releasing functional mitochondria in 0.5–1-µm MVs (Fig. 9c) (Liu et al. 2023). These large EVs carry a single mitochondrion and are named mitophers. Mitopherogenesis correlates with sperm motility and fertility, suggesting that spermatids use EV release to reduce their mitochondrial content to become functional spermatozoa.

Primordial germ cells also use EVs to remodel their size and organelle content during development. In C. elegans, the primordial germ cells Z2 and Z3 extend ∼3-µm lobes into the neighboring endoderm containing most of their mitochondria (Abdu et al. 2016). Lobe scission by the endoderm results in primordial germ cells losing half their cell volume, in addition to half their mitochondrial DNA (Abdu et al. 2016; Schwartz et al. 2022). This provides a complementary method to mitophagy for the clearance of germ cell mitochondria (Schwartz et al. 2022), allowing the germ cell to outsource mitochondrial degradation to the neighboring endoderm. This depletion of nutrients could be advantageous for a cell type that will be quiescent until nutritional signaling induces germ cell proliferation (reviewed in Kimble and Crittenden 2005; Hubbard et al. 2013).

EVs remove toxic proteins or dysfunctional organelles

Cells can shed EVs to respond to the buildup of toxic proteins or dysfunctional organelles after physiological or foreign insults. EVs are often a result of stress signaling, both autonomous and cell non-autonomous. In C. elegans, the stress-activated p38 MAP kinase PMK-1 is significantly enriched in EVNs and promotes PKD-2 ciliary EV biogenesis (Wang et al. 2015). Proteostatic stress directly influences exopher formation in touch receptor neurons (Melentijevic et al. 2017), and systemic stress signaling affects exopher production (Cooper et al. 2021). EV release bears similarity to retinal photoreceptor renewal, which involves the nightly shedding of the outer segment after light-induced proteotoxic stress (Spencer 2023). Therefore, EV release is a key cellular response to stress pathways.

Exophers provide a great example of EV formation to concentrate protein aggregates or dysfunctional organelles and release them for contained degradation in other cells (Fig. 9d). Exophers have been observed in C. elegans neurons, mouse cardiac muscles, and renal epithelial cells (Melentijevic et al. 2017; Arnold et al. 2023; Nicolás-Ávila et al. 2020; Huang et al. 2023). In C. elegans touch neurons, exophers accumulate damaged mitochondria, protein aggregates, and toxic proteins, which are then released as large EVs as a neuroprotective measure (Melentijevic et al. 2017). Exophergenesis in the anterior lateral microtubule (ALM) neuron plays a role in neuronal protection as inhibiting exophergenesis compromises touch sensitivity in neurons expressing neurotoxic proteins. Exopher uptake and endolysosomal degradation by the neighboring epidermis likely allow the neuron to focus on its signaling function by outsourcing degradation (Wang , Arnold, et al. 2023). However, the transfer of aggregates in exophers also supports the idea that EVs could contribute to the spread of protein aggregates in neurodegenerative diseases.

EVs can also remove toxic protein aggregates after exposure to toxins. A low concentration of methylmercury (MeHg, < 0.5 µM) increases the number of EVs seen around the dendrites of cephalic (CEP) neurons (Ke et al. 2020b), but a higher concentration (5 µM) could also decrease the number of EVs (Ke et al. 2020a). The size of the toxin-induced neuronal EVs ranged from 1–3 µm, as measured by confocal fluorescence microscopy. However, fluorescence intensity can skew apparent sizes and the diameters would be better evaluated by EM. The observation that dopaminergic neurons can respond to environmental toxins by releasing EVs may be relevant to EV contributions to the degenerative propensity of dopaminergic neurons in Parkinson's disease.

Functions for EV-receiving cells

EVs carry morphogens and regulate patterning

Signaling is the most cited function for EVs, especially carrying signaling molecules that would not easily leave cells through classical secretion pathways. Morphogens are long-range signaling proteins that regulate developmental patterning as well as cancer but can be limited in movement due to hydrophobic lipid modifications (Parchure et al. 2018). For example, Wnts are palmitoylated in the ER before secretion, while Hedgehogs (Hhs) are both palmitoylated and cholesterylated, which would limit morphogen diffusion if the lipid groups were not embedded in a mobile hydrophobic structure like an EV membrane. Wnts and Hhs pattern Drosophila imaginal discs through EV-based signaling (Gross et al. 2012; Matusek et al. 2014; Hurbain et al. 2022).

In C. elegans, the Wnt EGL-20 travels long distances extracellularly (Pani and Goldstein 2018), but whether EGL-20 is found on an EV or another type of particle is unclear. In contrast, the Hh-related proteins WRT-2 and WRT-8 are carried on exosomes (Liégeois et al. 2006). Epidermal cells secrete exosomes that regulate cuticle formation during development (Fig. 10a). The epidermis plays a crucial role in secreting extracellular matrix and organizing the ridge shape of the alae. Mutants that disrupt the Hedgehog pathway or exosome release exhibit defects in alae ridges in L1 larvae and adults, suggesting that exosomes carry Hedgehog morphogens through the extracellular space to pattern the overlying cuticle. Indeed, small and large EVs are most abundant mid-L4 stage, while the adult cuticle is being formed (Katz et al. 2022). However, it is unclear whether Hedgehog-related proteins on exosomes are providing signals to receiving cells or supporting the extracellular matrix.

Fig. 10. Functional roles of EVs for EV-receiving cells. a) In L1 larvae and adults, two lateral rows of cuticle fold into three ridges, known as alae, formed by the deposition of structural extracellular matrix. The underlying epidermal cells release exosomes carrying morphogens, such as hedgehog-like proteins WRT-2 and WRT-8, to instruct alae formation. b) PVD neurons form extensive dendritic arbors to cover the body area beneath the cuticle and sense harsh touch. Dendritic damage stimulates seam cells to release EVs containing fusogens like AFF-1 that repair dendritic connections. c) Embryos emit signals inducing the body wall muscle to convert muscle mass into yolk nutrition, released in large exophers to feed oocytes for embryogenesis and larval growth. d) Apoptotic cells expose PS on the outer leaflet of the plasma membrane and release small PS-positive EVs that facilitate PS exposure on neighboring cells and promote phagocytic clearance.

The first embryonic midbody remnant has also been implicated in dorsoventral patterning (Singh and Pohl 2014). Cortical flows move the P0 midbody remnant toward the future ventral side of the two-cell embryo. Astral microtubules in the dividing P1 spindle contact the cell cortex neighboring the P0 midbody remnant. The contact to the midbody remnant appears to instruct P1 spindle orientation, as ablating a region around the midbody remnant with a UV laser leads to P1 division along the D–V axis instead of the A–P axis, disrupting the normal positioning of cells in the four-cell embryo. However, whether the midbody remnant contributes to dorsoventral patterning using signaling molecules or mechanical deformation on the P1 membrane remains to be determined.

EVs carry fusogens to repair dendritic arbors

Damaged cells can induce neighboring cells to release EVs to repair injuries. In C. elegans, the nociceptive mechanosensory PVD neuron exhibits extensive dendritic arborization covering most of the worm's body (Tsalik et al. 2003). After damage to dendrites, such as the removal of branches (dendrotomy), PVD neurons can repair by fusing intact dendrite branches together (Fig. 10b) (Oren-Suissa et al. 2010). AFF-1 and EFF-1 are transmembrane proteins that act as homotypic or heterotypic fusogens during developmental cell–cell fusion events (Mohler et al. 2002; Sapir et al. 2007). While EFF-1 acts cell autonomously in PVD, AFF-1 is expressed in epidermal seam cells, and its expression is upregulated in response to dendrotomy (Oren-Suissa et al. 2017). Seam cells release EVs containing AFF-1, although it is unclear whether the AFF-1 EVs are derived from the plasma membrane, filopodia, or endolysosomal organelles. Intriguingly, AFF-1 EVs can fuse with cells expressing EFF-1 (Avinoam et al. 2011), suggesting that the AFF-1 EVs mediate membrane fusion between the dendritic branches (Fig. 10b). However, it is unclear whether AFF-1 on EVs interacts with low levels of AFF-1 that are not detectable on PVD or with EFF-1 on the plasma membrane of PVD. Furthermore, it will be interesting to determine whether the EVs only provide the fusogen protein for dendritic branch fusion or whether the fusogen allows the EVs to fuse with the arbors to provide additional membrane for repair.

EVs carry nutrition to feed developing cells

EVs can transfer metabolic cargo to other cells (Buzas 2023). In zebrafish, exosomes transfer nutrients from the yolk syncytial layer to distant tissues through the circulatory system (Verweij et al. 2019). In C. elegans hermaphrodites, embryos induce exopher production from body wall muscle cells to deliver yolk to oocytes (Fig. 10c) (Turek et al. 2021). RNAi knockdown of vitellogenin vit-1, the principal intestinal yolk protein (reviewed in Perez and Lehner 2019), doubles the number of muscle-released exophers (Turek et al. 2021), suggesting that starved embryos can signal to body wall muscles to compensate for reduced nutrition secreted by the intestine. Increased exopher release results in faster progeny growth, correlating a developmental advantage with EV-mediated cargo transfer.

EVs aid the recognition of dying cells

Dying cells can use EVs to promote their clearance by neighboring cells, which is important to avoid damage from cells lysing in tissues. In C. elegans embryos, apoptotic cells produce small EVs with the lipid PS exposed on the surface of their membrane (Fig. 10d) (Mapes et al. 2012). The PS-positive EVs are correlated with the appearance of low levels of PS on the surface of the engulfing cell and successful clearance of the cell corpse by phagocytosis. Expression of the PS-binding probe LactC1C2 can increase the number of PS-positive EVs while simultaneously blocking PS exposure on the engulfing cell and disrupting corpse clearance, suggesting that the PS-binding probe blocks both PS function and EV clearance. This implies that PS needs to be recognized by the engulfing cell for the cell to expose low levels of PS and phagocytose the cell corpse. Indeed, increasing PS exposure on engulfing cells by disrupting the PS flippase TAT-1 can increase phagocytosis of living cells, cell corpses, and midbody remnants (Darland-Ransom et al. 2008; Mapes et al. 2012; Fazeli et al. 2020). Furthermore, the observed increase in EVs after expression of the PS-binding probe LactC1C2 suggests that PS signaling promotes the clearance of the PS-positive EVs (Mapes et al. 2012), although it is unclear whether the PS-positive EVs fuse with the receiving cell or are taken up by endocytosis. The apoptotic PS-positive EVs are therefore thought to help initiate phagocytic signaling for corpse engulfment, although it is difficult to disentangle the roles of PS on the surface of the cell corpse from the PS on the surface of the apoptotic EVs.

EV functions for inter-organismal communication

EVs regulate male locomotory and mating behavior

Caenorhabditis elegans release EVs into the environment and transfer EVs among individuals, thus serving as a useful model for illustrating the fundamental functions and mechanisms of EVs in mediating inter-organismal communication. Caenorhabditis elegans males possess 27 EVNs with cilia whose tips prolifically shed signaling EVs into the environment (Fig. 11b; Wang et al. 2014; Silva et al. 2017; Wang et al. 2020). EVs isolated from wild-type male-enriched cultures trigger tail-chasing behavior in EV-recipient males (Fig. 11a), while environmental EVs isolated from ciliary EV-defective klp-6 or tba-6 mutant cultures fail to elicit male tail-chasing behavior (Wang et al. 2014; Silva et al. 2017). These results suggest that environmentally released ciliary EVs contain specific signaling cargos that stimulate locomotory changes in males. Male tail-chasing behavior shares a signature motif with male mating behavior, persistent backward movement (Liu and Sternberg 1995). Another mating-related behavior is male clumping, where males aggregate together (Gems and Riddle 2000). Male clumping behavior depends on LOV-1 and PKD-2 (Kaletta et al. 2003), cargos of male ciliary EVs (Wang et al. 2014; Walsh et al. 2022), raising the possibility that additional male behaviors could be regulated by EVs. EVN cilia are likely specialized devices for the release of signaling EVs, a secretory function of cilia that has yet to be fully recognized (Luxmi and King 2024).

Fig. 11. EVs mediate inter-organismal communication. a) Isolated environmental EVs spotted on a plate induce males to switch from normal sinusoidal movement to backward tail-chasing behavior. b) Males transfer multiple types of EVs to hermaphrodites during mating, including ciliary EVs, seminal fluid containing ENPP-1, and sperm-derived EVs carrying MSP that induce ovulation.

Males can transfer EVs during mating and ejaculation. In many animals, EVs produced by the epididymis or found in the seminal fluid can alter the physiology of female reproductive tract cells (Foot and Kumar 2021; Tamessar et al. 2021). In Drosophila, male EV transfer can even regulate female remating behavior (Corrigan et al. 2014). In C. elegans, males deposit ciliary PKD-2 EVs on the hermaphrodite cuticle surrounding the vulva during mating and deliver the ectonucleotide pyrophosphatase/phosphodiesterase I (ENPP-1) from the cuboidal epithelia of the vas deferens into the uterus (Fig. 11b, Wang et al. 2020; Nikonorova et al. 2022). Additionally, males transfer double-membrane MVs derived from spermatozoa (Fig. 6, Fig. 11b). Once inside the spermatheca, spermatozoa release MVs containing MSP to induce ovulation in females (Kosinski et al. 2005). Thus, C. elegans males release distinct types of EVs that may optimize reproductive capacity, although the mechanisms remain to be determined.

Open question: parasitic helminth EVs

EVs facilitate the transfer of proteins, lipids, and nucleic acids integral to the secretome of parasitic helminths, which include nematode and flatworm species (Britton et al. 2020; Hoffmann et al. 2020; Sotillo et al. 2020; Drurey and Maizels 2021. Parasitic helminths are thought to infect more than a quarter of the world population (Jourdan et al. 2018; GBD 2017 Disease and Injury Incidence and Prevalence Collaborators 2018), in addition to infecting livestock and crops (Montarry et al. 2021), making helminths a major economic and health burden. Parasitic EV cargos can modify host biological processes, including immune responses and cellular metabolism (Eichenberger et al. 2018; Gazzinelli-Guimaraes and Nutman 2018; Ressel et al. 2019). Consequently, understanding the molecular composition, biogenesis, and cargo sorting mechanisms of helminth EVs holds significant potential for therapeutic applications (Sánchez-López et al. 2021). Analyzing the composition of helminth EVs could also serve as a biomarker to reveal critical insights into the status and severity of infections (Mu et al. 2021). Moreover, employing biocontrol strategies and harnessing helminth EVs present a promising avenue for treating parasitic, allergic, and autoimmune diseases (Siles-Lucas et al. 2015; Eichenberger et al. 2018; Sánchez-López et al. 2021).

Research in parasitic helminths is constrained by a lack of genomic and proteomic knowledge, advanced molecular and genetic manipulation techniques, standardized methods for EV isolation, and consistent culture conditions (Al-Jawabreh et al. 2024). The nematode C. elegans can therefore serve as a valuable model for helminth EV research, particularly for parasitic nematode EVs (Duguet et al. 2020). Given the high degree of conservation in the nervous systems among nematodes (Schafer 2016), understanding ciliary EVs released by C. elegans males could provide vital clues about how sensory cilia in parasitic nematodes release signaling EVs (Wang and Barr 2018; Akella et al. 2020). Furthermore, the investigation of EV cargoes unique to nematode species but highly conserved among nematodes, such as hedgehog-like proteins (Liégeois et al. 2006; Bürglin 2008), could lead to novel methods for disrupting EV-based signaling. Therefore, fostering interdisciplinary collaboration among nematologists could significantly advance EV science and human health.

Open question: what else can EVs do?

The studies discussed here provide only a glimpse into the EV landscape in C. elegans. Proteomic profiling has identified >3,000 proteins in environmental EVs (Duguet et al. 2020; Russell, Kim, et al. 2020; Nikonorova et al. 2022), which reveal that the cargo of environmental EVs is likely to be derived from virtually every tissue and cell type. In addition, EVs can remain within the animal, suggesting that many potential external and internal functions of EVs and their underlying principles remain to be discovered. For example, migrating cells in other animals leave behind trails of EVs known as migrasomes that contribute to polarized cell movement (Zhai et al. 2024). Whether migrasomes are produced by migrating cells in C. elegans or regulate cell movement during embryonic or larval development remains unexplored.

The intestine is the largest organ in C. elegans and is likely to be a primary source of environmental EVs based on proteomic studies (McGhee 2007; Nikonorova et al. 2022). Intestinal microvilli shed EVs in mammals and EV cargoes originate predominantly from the gastrointestinal tract in parasitic helminths (McConnell et al. 2009; Carrera-Bravo et al. 2021). Intriguingly, the dsRNA transporter SID-2 localizes to intestinal microvilli in C. elegans and is a cargo of ciliary EVs (Winston et al. 2007; McEwan et al. 2012; Nikonorova et al. 2022), making it plausible that SID-2 could also be shed in microvilli-derived intestinal EVs. In other systems, EVs can transfer small RNA between cells to mediate cellular defenses (Claycomb et al. 2017; Cai et al. 2019; Ressel et al. 2019; Duguet et al. 2020; Munhoz da Rocha et al. 2020; O'Brien et al. 2020). In C. elegans, small RNA from pathogenic bacteria triggers germline expression of the Cer1 retrotransposon, activating neurons to avoid the bacteria in a form of transgenerational memory transfer (Moore et al. 2021). Whether EV-mediated RNA transfer plays a role in Cer1-mediated defense is unclear, but it is likely that intestinal EVs have important functions in C. elegans.

In addition to potential bacterial regulation of intestinal EVs, the interaction between C. elegans and microbial EVs in its microbiome or environment is poorly understood. The most common proteins identified in C. elegans EV proteomic profiles are from outer membrane vesicles of their laboratory food source, Escherichia coli (Russell, Kim, et al. 2020; Nikonorova et al. 2022), suggesting potential interplay between bacterial EVs and C. elegans. Furthermore, C. elegans can take up EVs from microalgae and alter metabolic gene expression in response to EVs isolated from rumen fluids (Picciotto et al. 2022; Choi et al. 2023). More broadly, C. elegans could be capable of responding to ecological EVs from predator and prey species.

In addition to physiological roles of EVs, C. elegans could also serve as an ideal model for studying engineered EVs, particularly in understanding EV uptake pathways and drug efficacy. For example, EVs carrying superoxide dismutase can extend lifespan in C. elegans (Shao et al. 2023), building on the strengths of C. elegans in aging research (reviewed in Collins et al. 2008). In summary, C. elegans has emerged as an exemplary model for EV research.

Which techniques can be used to visualize and/or analyze EVs?

ISEV has published guidelines for EV research (Théry et al. 2018; Welsh et al. 2024), recommending the use of complementary approaches to confirm the presence and activity of EVs. The MISEV guidelines also summarize common challenges in EV research, including challenges in labeling EVs and confusion of non-EV particles with EVs. In this section, we highlight the features and limitations of different techniques for EV research using C. elegans examples.

EM of EVs

EM is considered the gold standard for EV research (Verweij et al. 2021), enabling the visualization of the enclosing lipid bilayer to confirm the vesicular nature of EVs. EM also reveals intricate EV structures, such as vesicles within vesicles, multiple layered membranes, and tubulovesicular organelles (Verweij et al. 2021). Caenorhabditis elegans is well suited to EM studies, because all stages of the worm can be high-pressure frozen to preserve the ultrastructure (Kosinski et al. 2005, Liégeois et al. 2006; O’Hagan et al. 2011; Wang et al. 2014; Maguire et al. 2015; Silva et al. 2017; Melentijivec et al. 2017; Akella et al. 2020; Turek et al. 2021, Wang, Arnold, et al. 2023; Arnold et al. 2023; Liu et al. 2023). Classical fixation methods can also be effective for observing worm EVs in situ (Wehman et al. 2011), which was also used for the WormAtlas.org anatomical database. EVs can be observed using scanning electron microscopy (SEM) or TEM (Verweij et al. 2021). SEM allows the observation of intact C. elegans or combination with fluorescent approaches for correlated light and electron microscopy (CLEM) approaches, but SEM has lower resolution (∼5 nm) than TEM (<0.5 nm). TEM approaches require thinning tissue to 50–100 nm (Fig. 12a), which is the diameter of small EVs. To accurately determine the size of EVs, many sections or serial sections should be analyzed by TEM to compensate for catching fragments of EVs in ultrathin sections. There are also 3D approaches like electron tomography that help distinguish EVs from plasma membrane projections (Fig. 12c-d) (Wehman et al. 2011; Wang et al. 2014; Silva et al. 2017). Tomography allows thicker 200–400-nm sections to be analyzed by TEM using tilt series and back projection to compute a virtual image stack (Markert et al. 2017). Tomography can also be performed serially to enable the analysis of micron-sized large EVs like midbody remnants in situ (König et al. 2017; Fazeli et al. 2020).

Fig. 12. Visualization of EVs using electron microscopy. a) TEM of the cell contract from a wild-type two-cell embryo after high-pressure freezing. Data were collected for Wehman et al. (2011). b) Negative staining of environmental EVs isolated from C. elegans culture. Data were collected for Wang et al. (2014). c), d) Computed section from a 3D electron tomogram of a 200-nm section of the cell–cell contact (C) from a two-cell embryo that overproduces EVs by ectocytosis. The magnified region in d shows the cross section of a plasma membrane bud. Data were collected for Wehman et al. (2011).

Furthermore, EM is compatible with different stains that can provide contrast to the EV membrane or to EV contents. For example, negative staining of isolated EVs can be used to judge the purity of an EV preparation (Fig. 12b) (Wang et al. 2014; Verweij et al. 2021; Nikonorova et al. 2022). EM can also be combined with immunostaining to visualize the localization of EV cargos in situ (Kosinski et al. 2005, Liégeois et al. 2006; Wang et al. 2014), bridging the high resolution of TEM with the specific localization of cargos on EVs. Immuno-EM thereby allows the characterization of cargos on a subset of EVs within a heterogeneous population (Wang et al. 2014). Additionally, immunostaining can be useful for tracing the trafficking routes of EV cargoes along different organelles, which can help elucidate the mechanisms of EV biogenesis (Kosinski et al. 2005, Liégeois et al. 2006). However, a major limitation of EM studies is that EM can only be performed on fixed tissues; the gain in spatial insights comes at the cost of temporal dynamics.

Fluorescent imaging of EVs

EVs are often first suspected through fluorescent imaging approaches, where punctate fluorescence of a reporter protein is observed to move away from the source cell (Table 2). Fluorescent reporters allow the tracking of EV dynamics, including EV targeting to specific tissues, EV uptake, and intracellular breakdown of EV cargos (Verweij et al. 2021). EVs are often labeled with fluorescent transmembrane proteins found in the plasma membrane or endolysosomal membranes. In C. elegans, tetraspanins TSP-6 and TSP-7 are ciliary EV cargos and have been used to track EV uptake into neighboring glial cells and EV release into the environment (Razzauti and Laurent 2021; Nikonorova et al. 2022; Razzauti et al. 2023). GCY-8, GCY-18, GCY-23, and SRTX-1 are microvillar cargos phagocytosed as EVs by glia (Raiders et al. 2021). Similarly, LOV-1, PKD-2, and CLHM-1 are transmembrane EV cargos used to track environmental release of different subpopulations of ciliary EVs (Fig. 13a-b) (Wang et al. 2014; Clupper et al. 2022; Walsh et al. 2022). However, overexpression of fluorescent transmembrane proteins can alter EV size and abundance (Razzauti and Laurent 2021), making it important to consider using endogenously tagged proteins or single-copy reporters (reviewed in Nance and Frøkjær-Jensen 2019). It is also important to verify that the fluorescent tag does not alter the function of the cargo proteins, the EV size, or abundance using functional assays, EM, or additional reporters.

Fig. 13. Visualization, characterization, and tracking of EVs using fluorescence microscopy. a), b) RnB neurons (1–5, 7–9) in the male tail release many EVs into the environment from their cilia. The transmembrane cargos, LOV-1 (magenta) and PKD-2 (green), are conserved markers for ciliary EVs (arrows and arrowheads). Data were collected for Walsh et al. (2022). Midbody remnants (arrowheads) are labeled with a plasma membrane reporter (PH, cyan) and a contractile ring reporter (NMY-2, yellow) before (c) and after (d) phagocytosis, enabling their tracking. Data were collected for Fazeli et al. (2016). e), f) A plasma membrane-localized degron reporter (PH::ZF1, yellow) allows the specific labeling of MVs in vivo after degradation is initiated in somatic cells (left side of embryo in e). More MVs (arrowheads) are observed between the eggshell and cell surface in partial loss-of-function pad-1 mutant embryos. PB, polar body. Data were collected for Fazeli et al. (2020).

Tagged cytosolic cargos can also be used to label EVs. Markers of the spindle midbody or contractile ring, such as ZEN-4 and NMY-2 reporters (Fig. 13c-d), allow tracking of midbody remnant formation and endolysosomal clearance by LC3-associated phagocytosis (Chai et al. 2012; Green et al. 2013; Ou et al. 2014; Singh and Pohl 2014; Fazeli et al. 2016; König et al. 2017). Similarly, actin reporters have been used to track the in vivo clearance of residual bodies by phagocytosis (Huang et al. 2012). However, not every secretion event occurs in EVs, so additional methods such as EM are necessary to demonstrate that the puncta labeled by a fluorescent reporter are in fact EVs. Care should also be taken to distinguish reporter fluorescence from autofluorescence, which can arise from the worm, debris from microscope slides, culture media, or bacterial food sources. Small EVs are rarely visible under bright-field imaging, which can help distinguish EVs from larger autofluorescent debris or bacteria.

One challenge with fluorescent labeling of EVs is the limit in spatial resolution. Fluorescent reporters will change their apparent width based on abundance and the resulting changes in fluorescence intensity, making it difficult to determine EV size based on fluorescent puncta, especially for EVs with diameters below the ∼0.25-µm diffraction limit of light. Furthermore, EVs found next to the releasing cell can be hidden in the fluorescence of the plasma membrane if both are labeled. For example, the PH domain of PLC is a common plasma membrane reporter that binds to the lipid PI4,5P2 (Szentpetery et al. 2009). EVs are only detectable with PH reporters when EVs move away from the releasing cell or accumulate to form large clusters of EVs (Wehman et al. 2011). One approach to circumvent this limitation is to degrade the reporter in the releasing cell after EV release using a degron protection assay (Beer et al. 2019). Degrons are motifs recognized by ubiquitin ligases for regulated polyubiquitination and proteasomal or lysosomal degradation and are commonly used in C. elegans for loss-of-function approaches (reviewed in Nance and Frøkjær-Jensen 2019). However, once a reporter is released in an EV, the reporter is protected from degradation, allowing degron-tagged reporters to selectively label EVs after degradation is induced (Fig. 13e-f). Endogenous C. elegans degrons such as the ZF1 domain of PIE-1 and the C-terminal phosphodegrons (CTPD) of OMA-1 have been used to observe embryonic EVs based on the developmental timing of degradation initiation (Beer et al. 2019), but this selective labeling approach is likely to work at other stages and with other degrons such as the auxin-inducible degron (AID2) (Negishi et al. 2022).

Another approach to overcome the diffraction limit of fluorescent EV reporters involves the use of super-resolution imaging modalities, such as structured illumination, array detectors, stimulated emission depletion (STED), and stochastic optical reconstruction microscopy (STORM) (Verweij et al. 2021). Super-resolution using array detectors like Airyscan offers a 2D resolution of 120 nm, compared to the 200–300 nm of standard confocal microscopy, and a Z-axis resolution of 350–400 nm, in contrast to the 500–600-nm Z-axis resolution achieved with conventional confocal microscopy (Verweij et al. 2021; Razzauti et al. 2023). Super-resolution imaging significantly enhances the visualization of EVs in 3D or 4D imaging (timelapse imaging with Z-stacks), allowing better separation of C. elegans EVs from their source or recipient cells (Razzauti and Laurent 2021; Wang et al. 2021; Nikonorova et al. 2022; Walsh et al. 2022; Razzauti et al. 2023; Wang et al. 2024b).

EV isolation and characterization

C. elegans culture for EV collection

As a complement to in vivo EV studies, EVs can also be isolated and characterized from C. elegans cultures (Fig. 14a). EVs released into the environment can be collected directly from standard agar plate cultures (Wang et al. 2014; Silva et al. 2017; Nikonorova et al. 2022). However, standard culture includes co-culture with E. coli bacteria as a food source, leading to co-isolation and contamination of C. elegans EV preps with E. coli outer membrane vesicles (Nikonorova et al. 2022). It is straightforward to computationally distinguish worm proteins and RNAs from bacterial sequences after analysis; this requires increased depth to capture sufficient C. elegans data from MassSpec or sequencing data of co-isolated EVs.

Fig. 14. Methods for EV isolation from c. elegans cultures. a) EVs are harvested by washing culture plates, which contain worms, bacteria, different EV subtypes, and other secreted macromolecules. EVs can also be collected from liquid culture. b) Differential centrifugation sequentially pellets worms and bacteria from the mixed sample. c) EVs can be isolated from the cleaned supernatant using ultracentrifugation on a cushion. Different speeds can be used to pellet large and small EVs. d) For increased purity, EVs can be pelleted from the cleaned supernatant by ultracentrifugation on a cushion and then loaded onto a gradient solution. Ultracentrifugation allows gradient fractionation to separate dense and light EVs. e) EVs can be isolated from cleaned supernatants using size exclusion chromatography, which allows the separation of EVs based on size.

To limit bacterial EV contamination, C. elegans have been grown without bacteria in liquid cultures (Duguet et al. 2020; Russell, Kim, et al. 2020). However, differences were observed in EVs isolated after long periods in axenic M9 buffer; the abundant cytosolic protein actin was only found in the 5-hour EV samples, not the 24-hour EV samples (Duguet et al. 2020). Given the nutritive roles of EVs and changes to EV biogenesis caused by starvation or cell stress, it is worth considering that different culture conditions can influence EV subtypes, abundance, cargos, stability, and thereby downstream functions. To facilitate comparisons between EV studies, it is important to include detailed methods about food sources (bacteria, lipids, carbohydrates, proteins, etc.), worm density, temperature, antibiotics, as well as potential shear stress from oxygenating liquid cultures.

Vesicles can also be isolated from homogenized C. elegans samples (Thomas et al. 2023; Ma et al. 2023), which can improve access to EVs within tissues. However, homogenization disrupts cell membranes, making vesicles isolated from homogenized samples likely to contain a mix of EVs, intracellular vesicles, and cell fragments induced by tissue damage. Therefore, while homogenization can increase vesicle collection, homogenization will also increase vesicle heterogeneity beyond EVs.

EV isolation

After EV collection, several methods have been applied to isolate and concentrate EVs from liquid media (Fig. 14) (Théry et al. 2018; Welsh et al. 2024). EV isolation is normally a balancing act between achieving sufficient purity from co-isolates and obtaining an adequate amount of material for downstream assays. Differential ultracentrifugation is a widely used method for EV isolation, involving a series of centrifugation steps at increasing speeds. For C. elegans samples, the slower speeds using a centrifuge remove whole worms, bacteria, and debris from the media (Fig. 14b), while faster speeds using an ultracentrifuge pellet large and small EV subpopulations (Fig. 14c) (Wang et al. 2014; Nikonorova et al. 2022). Ultracentrifugation is performed on a cushion of sucrose or iodixanol to avoid damaging EVs during high-speed spins (Nikonorova et al. 2022). Differential ultracentrifugation can lead to vesicle aggregation and co-isolate non-EV particles, such as lipoprotein particles and protein aggregates (Théry et al. 2018; Welsh et al. 2024).

The separation of EVs from co-isolates can be improved using density gradient centrifugation. EVs are separated based on their buoyant density in sucrose or iodixanol gradients (Nikonorova et al. 2022). Density gradient centrifugation facilitates the separation of distinct subpopulations of EVs (Fig. 14d), as well as enabling the separation of EVs from both soluble proteins and dense particles. The choice of gradient fraction for further study can be guided by imaging a fluorescently labeled EV subpopulation (Table 2), EM to observe vesicles, or another characterization method. Although density gradient centrifugation is time-consuming, it has proven effective at identifying endogenous EV cargos such as SID-2, MCM-3, and ENPP-1.

Another approach to isolate EVs is size exclusion chromatography, which separates EVs based on size without an ultracentrifuge. EVs are passed through a column with a porous matrix, allowing smaller molecules to be trapped in the pores, while larger particles like EVs elute earlier (Fig. 14e). Size exclusion chromatography preserves vesicle integrity while separating EVs from bulk proteins (Russell, 2020a; Russell, Kim, et al. 2020; Duguet et al. 2020). Methods isolating EVs based on size can separate small EVs from large EVs but are likely to contain a mixture of EVs derived from different organelles, cell types, and organisms.

Characterization of isolated EVs

Given the broad diversity of EVs and non-vesicular particles secreted by cells, it is important to characterize the purity of isolated EVs before functional assays (Théry et al. 2018; Welsh et al. 2024). Our bodies secrete many types of extracellular materials into body fluids, including lipoprotein particles, protein aggregates, soluble proteins, different types of RNAs, and DNAs (Sódar et al. 2016; Jeppesen et al. 2023). In addition, C. elegans cultures can include outer membrane vesicles from bacteria and particles such as viral capsids (Moore et al. 2021; Nikonorova et al. 2022). To confirm the presence of EVs, samples can be characterized by negative staining TEM, which reveals the shape and size of the EVs (Fig. 12b, Wang et al. 2014; Nikonorova et al. 2022). Immunostaining combined with negative staining TEM can further characterize the morphology of EVs containing the cargo of interest (Wang et al. 2014). Fluorescence microscopy can also be used on EV fractions to isolate the EVs carrying specific fluorescent cargos or other membrane markers (Wang et al. 2014; Nikonorova et al. 2022). Western blots could also be used to characterize EVs, as in mammalian systems where tetraspanins CD9, CD63, and CD81 are popular EV markers (Théry et al. 2018; Welsh et al. 2024). However, it is challenging to generate specific antibodies for C. elegans proteins (reviewed in Duerr 2006), and there are currently no validated EV antibodies for western blots.

Once the desired EV fraction has been isolated, proteomic profiling and RNA sequencing can be applied to further characterize the composition of the isolated EVs (Fig. 15; Duguet et al. 2020; Russell, Kim, et al. 2020; Nikonorova et al. 2022). Proteomic datasets can be a starting point for studies on EV biogenesis, cargo sorting, function, and clearance, as EV cargos can include proteins involved in each step of the EV life cycle. Analyses of small bioactive molecules and lipidomics have yet to be performed on C. elegans EVs, but EVs can also be artificially loaded with desired cargos (Shao et al. 2023), allowing their study for therapeutic delivery. In-depth characterization and exploitation of EV cargos could yield further insights into EV biogenesis and function.

Fig. 15. MyEVome, a tool for identifying EV cargo candidates for c. elegans cell types. MyEVome combines single-cell transcriptomic data with environmental EV proteomics to plot likely cell-specific EV cargos (Cao et al. 2017; Nikonorova et al. 2022). MyEVome is currently available at https://myevome.shinyapps.io/evome-app.

Functional assays with isolated EVs

Isolated EV samples can be applied to C. elegans in different ways for functional assays. Spotting EV samples onto culture plates has been effective at altering locomotory behaviors, transcription, stress resistance, and lifespan (Wang et al. 2014; Silva et al. 2017; Shao et al. 2023; Choi et al. 2023). This approach can also allow a side-by-side comparison of EV-treated and untreated worms on the same plate (Fig. 11a). Alternatively, worms can be incubated in an EV-containing buffer or EVs can be injected into the pseudocoelom (Picciotto et al. 2022), which can allow EVs to interact with different tissues. It is recommended to use a range of EV dosages to test whether a consistent effect is observed across different concentrations (Théry et al. 2018; Welsh et al. 2024). Dosage can be critical, as vesicles isolated from homogenized dauers extended lifespan at an intermediate dose but not at a 10-fold lower or higher dose (Ma et al. 2023). EV quantities can be normalized using their protein concentrations (Wang et al. 2014; Silva et al. 2017) or particle counting instruments (Théry et al. 2018; Welsh et al. 2024).

Additional control experiments using EVs isolated from mutant strains or pre-treating EVs can also help identify the EV subpopulation or cargo that is having a functional impact. For example, environmental EVs isolated from klp-6 and tba-6 mutants fail to alter male locomotory behavior, allowing the function of ciliary EVs to be separated from other EVs in the sample (Wang et al. 2014; Silva et al. 2017). It is best practice to isolate EVs from wild-type and mutant cultures concurrently to ensure their comparability. Another common control is to use detergent treatment to disrupt the lipid membrane of EVs and confirm whether the function of an EV sample is lost. Detergent treatment can also be used in combination with protease or nuclease protection assays to determine whether an EV-associated function is due to an internal EV protein, DNA, or RNA cargo (Théry et al. 2018; Welsh et al. 2024). However, EV functions can also be carried out by cargos on the surface of EVs, known as the EV corona (Buzas 2022), making it more challenging to distinguish functional co-isolates from EV surface proteins. In summary, standardizing methods for C. elegans culture, EV isolation, and EV application holds great potential for understanding the functions of environmentally released EVs.

Open question: overcoming gaps in EV techniques

One challenge for EV research compared to most intracellular organelles is the lack of a pan-EV marker to enable EV labeling from any tissue. EVs typically reflect the composition of their source organelle and cell type, which means that EV studies often necessitate the generation of new markers for each tissue or EV subtype of interest. Table 1 lists current examples of C. elegans tissues where EV release or uptake has been observed, and Table 2 lists markers that have been applied to different EV subtypes. Moreover, combining information from the proteomic characterization of environmental EVs with single-cell proteomic studies is likely to identify new reporters that can be applied to new EVs, an approach simplified by the online data mining tool MyEVome (Fig. 15, Cao et al. 2017; Nikonorova et al. 2022). The EV proteome data used in MyEVome originate from mixed-stage cultures containing male and hermaphrodite C. elegans, while the single-cell transcriptomic profiling dataset was obtained from a synchronized L2 hermaphrodite population. Consequently, candidate EV cargos should be validated further. Developing additional EV markers from different tissues will greatly enhance our understanding of EV biogenesis, diversity, and function in C. elegans.

Another challenge with EVs is obtaining adequate fluorescent signal from EV markers to monitor EV biology in the context of organismal fluorescence. While approaches have been developed to decrease cellular background, like degron protections assays (Beer et al. 2019), approaches that boost EV signal without disrupting EV function need to be explored further, especially for small EVs with limited cargos. Single-molecule localization has been performed on tumor cell exosomes made larger using expansion microscopy (Wei et al. 2023), and C. elegans samples can also be expanded (Yu et al. 2020). Possible solutions include tandem fluorescent protein tags on EV reporters, with the caveat that mCherry aggregation can induce neuronal exopher formation (Melentijivec et al. 2017). Improving EV signals would help trace cargo trafficking routes into EVs, the release and movement of EVs, EV–cell interaction modes, and the modalities of EV uptake and clearance.

A further challenge for EV characterization is obtaining a pure EV population from C. elegans culture, which would facilitate deeper profiling of the EV proteome and transcriptome. One approach to avoid contamination with bacterial outer membrane vesicles could be feeding C. elegans with E. coli mutant strains that are defective in shedding outer membrane vesicles (Premjani et al. 2014; Murase et al. 2016). However, given that EVs can signal between species, it is prudent to first investigate whether bacterial outer membrane vesicles (OMVs) influence C. elegans physiology, behavior, or EV production. Alternatively, chemically defined culture media that support normal C. elegans development without bacteria would also be an option (Zečić et al. 2019), which could also allow the dissection of metabolite influence on EV biogenesis. It may also be possible to use microfluidic devices to characterize EV diversity without isolating C. elegans EVs from cells and bacterial EVs, similar to studies of whole blood (Davies et al. 2012; Wu et al. 2017; Chen et al. 2019; Meng et al. 2023).

Alternatively, in vivo labeling of EV cargos before ex vivo characterization could be a promising strategy to avoid characterizing co-isolates like bacterial EVs while maintaining standard culture conditions. Proximity labeling approaches fuse an EV cargo protein to a biotin ligase to post-translationally modify nearby proteins (Löf et al. 2017; Kaneda et al. 2021; Li et al. 2023). The biotinylated proteins can then be isolated through biotin–streptavidin interaction, reducing the number of non-EV proteins to be identified. As different EV cargos can be labeled, this allows the characterization of specific EV subtypes, as well as differentiating EV cargo inside the lumen from cargo on the EV surface or corona. As proximity labeling has been applied successfully in C. elegans (Branon et al. 2018; Sanchez and Feldman 2021), the technique is likely to help identify EV protein cargos.

Conclusions

Caenorhabditis elegans serves as a model organism for both in vivo studies of EVs within a physiological context and the roles of environmental EVs. C. elegans EV research has illustrated conserved pathways in EV biogenesis, uptake, and clearance in addition to EV functions regulating tissue homeostasis, regeneration, developmental signaling, and social interactions. Studies to date highlight the power of the C. elegans model in elucidating fundamental principles of EV biology. The discovery that EV biogenesis is dependent on cellular activity, development, and cross-tissue signaling provides a mechanistic understanding of EV heterogeneity across the whole organism. At the same time, conserved themes in EV biogenesis pathways reveal that EVs are integral extracellular organelles with key roles in cellular processes. C. elegans is poised to lead the way in exciting areas of EV research, including the fundamental mechanisms governing how, when, and what types of EVs cells produce, as well as unearthing new functions for EVs. Research on C. elegans EVs is making big strides toward understanding how EVs contribute to health and disease.

Acknowledgements

The authors would like to thank Drs. Gholamreza Fazeli, Katharina Beer, and Inna Nikonorova and the New York Structural Biology Center for providing images. The authors thank Monica Driscoll, Barth Grant, our anonymous reviewers, and members of the Barr and Wehman labs for valuable feedback on the manuscript.

Funding

Funding was provided by the American Cancer Society and Deutsche Forschungsgemeinschaft (DFG) grant WE5719/2-1 (A.M.W.) and by the National Institute of Health (NIH) grants DK059418, DK116606, and NS120745 (M.M.B).
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