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Extracellular vesicles
Juan Wang,
1
Maureen M. Barr ,
1
Ann M. Wehman
2,
*
1
Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ 08854, USA
2
Department of Biological Sciences, University of Denver, Denver, CO 80210, USA
*Corresponding author: Department of Biological Sciences, University of Denver, 2101 E. Wesley Ave, Denver, CO 80208, USA. Email: awehman@alum.mit.edu
Extracellular vesicles (EVs) encompass a diverse array of membrane-bound organelles released outside cells in response to developmen-
tal 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 unex-
plored topics tenable in C. elegans, given the nematode model is ideal for light and electron microscopy, genetic screens, genome en-
gineering, and high-throughput omics.
Keywords: extracellular vesicle; Caenorhabditis elegans; exosome; microvesicle; cilia; spermatogenesis; midbody remnant
Received on 21 January 2024; accepted on 21 May 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of The Genetics Society of America.
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What are extracellular vesicles?
Beyond the plasma membrane, cells have extracellular orga-
nelles, including the extracellular matrix and extracellular vesi-
cles (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 organ-
elle biology.
The term “extracellular vesicle”? was rst 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 specic locations within cells
(Dixson et al. 2023; Sohal and Kasinski 2023). EVs reect their cellular
origin and are composed of a lipid bilayer membrane that encloses
cytosolic organelles and macromolecules, including proteins, meta-
bolites, 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 clas-
sied based on their size into small and large EVs, which is derived
from the methods used to separate EVs from body uids. Small
EVs, with diameters ranging from tens to hundreds of nan-
ometers, 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 con-
tents outside the cell. In contrast, MVs bud directly from the plas-
ma 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 challen-
ging 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 specic 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 sub-
types have specic cargo or biogenesis mechanisms that distin-
guish 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, di-
verse small and large EV subtypes can be utilized for different cel-
lular functions.
GENETICS, 2024, 227(4), iyae088
https://doi.org/10.1093/genetics/iyae088
Advance Access Publication Date: 17 June 2024
WormBook
EVs are implicated in human health, aging, and disease pro-
gression as EVs are involved in a wide range of physiological func-
tions, 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 deliv-
ery 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 contain-
ing cellular content are released into body uids, and blood collec-
tion is typically less invasive than tissue biopsy.
Major challenges in the EV eld include a limited understand-
ing of the molecular control of EV biogenesis and difculties track-
ing EV release in vivo. Observing structures with light microscopy
is technically challenging for objects smaller than 200 nm, espe-
cially within the tissue context. Furthermore, it has been challen-
ging to understand the complexity and heterogeneity of EVs and
the diverse functions of EV subpopulations often found in a het-
erogeneous EV sample isolated from body uids. Understanding
the commonalities and intricacies of EVs will enable the eld
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 organ-
ism due to its transparency, small size, and the extensive knowl-
edge 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.
How are EVs formed?
Exosome formation from endolysosomes
The biogenesis mechanisms of exosomes have been illuminated
by decades of research on endosomal trafcking pathways con-
served from yeast to mammals. Many excellent reviews summar-
ize 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).
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 com-
prises 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 laments that promote membrane curvature to
start budding the ILVs. VPS-4 ATPase activity remodels the
ESCRT-III laments to constrict the bud neck for scission and re-
lease 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 per-
turbations would alter exosome diameter as well as quantity,
which could also change exosome contents.
Spermatid
mitopher
0.5-1 µm
Spermatozoa
double membrane MV
150-300 nm
Neuronal and muscle
exophers
1-15 µm
Ciliary
EVs
50-200 nm
Neuronal
dendrite
MV
1-3 µm
Midbody
remnant
1-2 µm
Embryo
MV
100-400 nm
Epidermis
exosome
50-100 nm
Residual
body
~4 µm
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.
2 | J. Wang et al.
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 cu-
ticle of C. elegans, particularly visible in the L1 larval stage and
adults. A genetic screen for alae defects identied 73 genes that
may regulate exosome biogenesis (Fig. 2), including ESCRT-0 com-
ponent 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 pro-
moted by actin and the lamin FLN-2 (Shi et al. 2022). Using elec-
tron 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 n-2 mutants, electron-light MVBs are re-
duced with fewer ILVs, while electron-dark MVBs are almost ab-
sent (Shi et al. 2022), suggesting that FLN-2 promotes both ILV
formation and MVB maturation. Filamins crosslink F-actin to or-
ganize the cytoskeleton and other proteins on membranes, and
FLN-2 appears to bridge MVBs to actin laments. 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
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)
Extracellular vesicles | 3
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 mem-
branes (Egea-Jimenez and Zimmermann 2018). Lipid shape is
thought to inuence membrane curvature to aid membrane bud-
ding, but lipid conversions can also regulate lipid–protein interac-
tions to regulate protein localization to endosomes.
RAL-1 is also required for MVB fusion to release ILVs as exo-
somes (Hyenne et al. 2015), which have been visualized using EM
(Table 2). The constitutively active form of the RAL-1 GTPase colo-
calizes with the t-SNARE protein SYX-5, suggesting that RAL-1 in-
teracts 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).
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 cellu-
lar 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, in-
ammation, cancer, and the immune response (Hu et al. 2023;
Kalluri and McAndrews 2023). In C. elegans, embryonic blasto-
meres shed MVs that are conned by the eggshell or between cells.
Embryonic MVs can be labeled with uorescent 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).
The mechanisms of MV formation from the plasma membrane
are less well studied than ILV formation from endosomal mem-
branes, 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 forma-
tion from endosomal membranes (Fig. 2). In addition, the small
GTPase RAB-11 can promote MV biogenesis and was also identied
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 specic 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 phosphatidylethano-
lamine (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 cytosol-
ic face of the plasma membrane by the P4-ATPase TAT-5, which
functions as an aminophospholipid translocase to ip PE from
the exofacial leaet to the cytofacial leaet of the membrane bi-
layer and create lipid asymmetry (Fig. 3). TAT-5 is activated by
the Dopey homolog PAD-1, and their essential interaction is con-
served 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 leaet of the plasma membrane and 100–
400-nm spherical or tubular MVs accumulate between cells or
along the embryo surface. Excessive MV biogenesis disrupts gastru-
lation 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 ex-
pose 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 plate-
lets (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 phos-
phatidylserine (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
Endocytosis
ILV formation
Sorting
Transport
Degradation
Docking
Fusion and release
Exosomes
RAL-1
SYX-5
VHA-5
MVB maturation
RAL-1 ESCRT
Cargo
Cargo
Actin
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.
4 | J. Wang et al.
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-specic after two-cell
stage
Beer et al. (2019)
Germ cells
Spermatids and
spermatozoa
MSP Immunouorescence,
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-modied secreted protein Liegeois (2006)
Male-specic
EV-releasing
neurons
PKD-2::FP Fluorescence Microvesicle Transmembrane protein Wang et al. (2014)
Male-specic
EV-releasing
neurons
LOV-1::FP Fluorescence,
Immuno-EM
Microvesicle Transmembrane protein Wang et al. (2014),
Walsh et al.
(2022)
Male-specic
EV-releasing
neurons
MCM-3::FP Fluorescence Microvesicle Cytosolic and nuclear protein Nikonorova et al.
(2022)
Male-specic
EV-releasing
neurons
SID-2::FP Fluorescence Microvesicle Transmembrane protein Nikonorova et al.
(2022)
IL2, Male-specic
EV-releasing
neurons
CWP-1::FP Fluorescence Microvesicle Cytosolic protein Wang et al. (2014)
IL2, Male-specic
EV-releasing
neurons
DDN-3::FP Fluorescence Microvesicle Secreted or transmembrane
protein
Wang et al. (2015)
IL2, Male-specic
EV-releasing
neurons
CIL-7::FP Fluorescence Microvesicle Myristoylated protein Maguire et al.
(2015)
IL2, Male-specic
EV-releasing
neurons
TTLL-11b::FP Fluorescence Microvesicle Cytosolic protein O’Hagan et al.
(2017)
IL2, Male-specic
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)
(continued)
Extracellular vesicles | 5
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 pro-
cesses capable of disrupting lipid asymmetry.
The effect of lipid symmetry on MV budding seems specic to
PE, because PS asymmetry maintained by the TAT-1 ippase inu-
ences large EV uptake, not MV budding (Fazeli et al. 2020). PE
asymmetry and MV biogenesis are indirectly regulated by endoso-
mal recycling regulators that trafc 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 trafc lipid scramblases. Furthermore, it remains
to be determined which biophysical properties of PE lipids inu-
ence MV budding, such as conical shape, neutral charge, abun-
dance, 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 forma-
tion depends on actin laments 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 bio-
genesis from the plasma membrane of chondrocytes (Hale and
Wuthier 1987), suggesting that actin dynamics may play con-
served 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 fu-
sion 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 accu-
mulation in the plasma membrane and increased membrane pro-
trusion and EV shedding, suggesting that RAB-5 promotes EFF-1
endocytosis on the plasma membrane to maintain EFF-1 in intra-
cellular 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.
Table 2. (continued)
Source Tissue EV reporter Method EV subtype Notes Reference
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).
ESCRT subunits
bind to PI
and constrict the bud neck
to faciliate membrane scission
Microvesicle formation by budding and scission
TAT-5 is activated by PAD-1
to maintain PE asymmetry
PE becomes
symmetric
?
Zygote
Ectocytosis
PE
Homeostasis
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
ippase that transports PE lipids to the inner leaet 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.
6 | J. Wang et al.
Both stable protrusions like cilia and dynamic protrusions like
lopodia 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 decapi-
tation or ectocytosis (Nager et al. 2017; Phua et al. 2017), and the for-
mation 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-specic 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 her-
maphrodites (Razzauti and Laurent 2021). Furthermore, a set of
six IL2 and 21 male-specic sensory neurons have been specially
classied as EV-releasing neurons (EVNs) due to their prolic pro-
duction of 50–200-nm ciliary EVs (Wang et al. 2014). Electron tom-
ography 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.
The release of EVs from the ciliary tip relies on intraagellar
transport (IFT) (Wang et al. 2014; Clupper et al. 2022). IFT is import-
ant 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 import-
ant 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 en-
richment of EV cargo at the ciliary tip is essential for ciliary EV re-
lease (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 re-
lease (Wang et al. 2014; Clupper et al. 2022). Furthermore, the CHE-
3dynein 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 cil-
iary 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 al-
lows EVs shed from the ciliary tip to be released into the worm’s
environment. TEM analysis also revealed EVs in the lumen of la-
bial 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 modications, 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 microtu-
bules 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 modies microtubule axonemes and is only ex-
pressed in EVNs (O’Hagan et al. 2017). The counteracting enzyme
b EVN ciliary EV biogenesis
EV cargo ciliary tip enrichment:
OSM-3
KLP-11, KLP-20, KAP-1
CHE-3
IFT and accessory proteins
Axoneme specialization
Tip enrichment
PKD-2
TBA-6
TTLL-11b
CCPP-1
CEM (4) RnB(16, n=1-5,7-9)
a Pan ciliary EV biogenesis
HOB (1)
Sub-tip enrichment
CIL-7
KLP-6
IL2 Amphid Phasmid
Ciliary
tip
Ciliary
base
Glia
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-specic EVNs utilize cell
type-specic ciliary kinesin (KLP-6) to trafc cargos to the tip for sorting into EVs. KLP-6 and the tubulin TBA-6, along with the post-translational
modication enzyme polyglutamylase TTLL-11 and its counteracting enzyme, CCPP-1, specialize the axoneme to facilitate abundant EV release.
Extracellular vesicles | 7
responsible for deglutamylation of the microtubules, CCPP-1, has
cell-specic regulation in the EVNs and is required for efcient en-
vironmental EV shedding. Furthermore, the EVN-specic
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 sur-
rounding 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 de-
creases in neuronal activity by pruning neuronal microvilli by
phagocytosis. PS exposure on microvillar membranes is recog-
nized 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 specically on microvilli remain to be
determined, but at least one lipid scramblase, SCRM-1, helps pro-
mote PS exposure. If PS exposure is not restricted to microvilli, ex-
cessive pruning occurs, such as in the PS ippase mutant, tat-1.
Thus, organelles can also be fragmented to form EVs through in-
teractions with neighboring cells.
Double-membrane EVs from protrusions
Not all EVs are wrapped in a single membrane bilayer. In C. ele-
gans, spermatids and spermatozoa produce small EVs with two
membrane bilayers (Fig. 6, Kosinski et al. 2005). The outer mem-
brane of these EVs appears scalloped after high-pressure freezing
and TEM, characterized by a wavy surface. The major sperm pro-
tein (MSP) is sandwiched between the two membranes, while the
inner core appears less electron dense. MSP is a cytoskeletal pro-
tein predominantly found inside extensions like the pseudopod of
ameboid sperm (reviewed in L’Hernault 2006). This led to the pro-
posal that membrane protrusions would extend around extracel-
lular 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.
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 be-
gins 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 rst 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, mak-
ing these large EVs relatively easy to label and track.
Caenorhabditis elegans midbody remnants were rst 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, in-
dicating that the midbody remnants had been released extracellu-
larly 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) en-
gulfment pathways (Chai et al. 2012), which were identied 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 particu-
larly useful for dissecting the mechanisms of midbody remnant
biogenesis. Midbody remnants can be labeled with reporters
Microvilli-derived EVs aided by phagocytosis
Microvilli
Cilium
AFD sensory cilium and microvilli SCRM-1
?
Neuron
CED-2
CED-5
CED-12
WSP-1
CED-10
Glia
TTR-52
PSR-1
PAT-2
PS
TAT-1
Actin
Fig. 5. EV biogenesis from microvilli. Glial phagocytosis regulates the sensitivity of AFD neurons by engulng 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 ippase 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.
8 | J. Wang et al.
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 disas-
sembled from the midbody and septins create a cytoplasmic bar-
rier 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 forma-
tion (Figs. 2 and 3). Membrane fusion on one side of the bridge
leads to abscission (König et al. 2017), which allows the mem-
branes 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 dy-
namin DYN-1 also promotes abscission for midbody remnant re-
lease (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 re-
lease 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 rem-
nants 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 ac-
tin 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
Double membrane microvesicle formation by membrane protrusion driven by MSP
Spermatazoa
MSP protrudes
the plasma membrane
The protrusion collapses
back on itsself
Membrane fusion releases a
vesicle containing MSP
between the two membranes
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).
Microtubule
disassembly
& membrane
remodeling
Sealing to
release the
midbody
remnant
Furrow
ingression
Actin
NMY-2
Abscission
ESCRT
Embryo
a
Midbody remnant released as a
n
EV upon conclusion of cytokinesis
Spermatocyte
b
Residual body released as an EV upon conclusion of meiosis II
Pseudo-cleavage
furrow
ingression
Actin
NMY-2
Furrow
ingression
& scission
Actin
SPE-15
Residual
Body
Spermatids
Polarized
Trafficking
Primordial
Germ Cell
c
Germ cell lobe released as an E
V
after engulfment by the endoderm
Soma
Lobe
Pseudo-cleavage
furrow
ingression
Actin
NMY-2
Nuclear
migration
Lobe
engulfment
CED-10
Endoderm
Actin
Scission
to release
the lobe
LST-4
Cell
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.
Extracellular vesicles | 9
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 organ-
elle 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 polymer-
ization 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 anu-
clear 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 in-
testinal 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 engulf-
ment, but unlike most phagocytic events, CED-10 was not acti-
vated 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 rst appear as an outward bud of the cell mem-
brane and are not formed by reentry into the cell cycle, as exo-
phers 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 in-
volved in exopher constriction.
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 in-
creases the number of exophers, indicating that proteostress up-
regulates exophergenesis (Melentijevic et al. 2017). However,
while most autophagy proteins reduce the cellular need for exo-
phers, 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 au-
tophagy (reviewed in Durgan and Florey 2022). The non-canonical
role of ATG-16.2, dependent on its C-terminal WD40 domain, pro-
motes 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 uorescent re-
porter 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 exo-
phers. Cargo proteins such as Q128 rst aggregate and coalesce to
form aggresomes near the nucleus (Fig. 8b). Q128 trafcking into
aC. elegans hermaphrodites produce exophers from touch receptor neurons and body wall muscles
ALM (L/R)
AVM
PVM
PLM (L/R)Body wall muscle
N
Mito
Agg
NNNN
Stressed neuron Membrane bud Constriction
Exopher maturation Exopher released into epidermis
Further cargo loading
through nanotube
Membrane sealed
by phagocytosis
Pre-exopher stage, plastic fate of membrane bud
b Neuronal exopher formation is driven by protein aggregates and dysfunctional organelles
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 laments 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.
10 | J. Wang et al.
exophers depends on intermediate lament 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 con-
tinues to be trafcked into the growing exopher (Fig. 8b), including
calcium and protein aggregates (Melentijevic et al. 2017).
Exopher release depends on actin and Arp2/3-dependent en-
gulfment by a neighboring epidermal cell (Wang, Arnold, et al.
2023). Indeed, phagocytic receptors and membrane trafcking
regulators implicated in phagocytic engulfment, including the re-
ceptor CED-1, the small GTPase RAB-35, and the Arf GAP CNT-1,
are required in the epidermis for exopher engulfment through in-
hibition of the small GTPase ARF-6. ARF-6 effectors like the exo-
cyst 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, includ-
ing 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 leaet of the lipid bilayer, similar to the plas-
ma membrane of apoptotic corpses. Both the ATP-binding cas-
sette (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 oppases.
ABCA1 also alters the localization of cholesterol and sphingomye-
lin and changes lipid packing within the plasma membrane
(Landry et al. 2006), which may indicate a broader lipid specicity
than PS. In C. elegans, PS exposure did not increase EV release from
viable embryonic cells in tat-1 ippase 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; specic pro-
teins, RNAs, and lipids are selectively incorporated into EVs dur-
ing 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 specically enriched in neuronal exophers
compared to the cell body to promote their secretion in EVs
(Melentijevic et al. 2017). Specic cargos can also be excluded
from EVs during EV biogenesis. Abundant cortical proteins like
clathrin can be selectively excluded from MVs to prevent their se-
cretion in EVs (Beer et al. 2019). The sorting of cargos is also specic
to EV subtypes. Myristoylated plasma membrane markers and
cytosolic reporters can be specically excluded from RAB-5DN
touch neuron EVs (Linton et al. 2019), while being preferentially in-
cluded in touch neuron exophers and ciliary EVs (Maguire et al.
2015; Melentijevic et al. 2017). How C. elegans cells regulate select-
ive 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 pro-
mote 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 car-
go 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 secre-
tion in EVs, and several ILV budding pathways have been charac-
terized 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 sort-
ing into ILVs and exosome biogenesis (Trajkovic et al. 2008).
Some tetraspanins (e.g. CD63, CD81) can form membrane nano-
domains that sort cargos into ILVs independent of ESCRT path-
ways (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 trans-
duction 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 mem-
brane 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 her-
maphrodites, 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), dem-
onstrating that sensory signals can regulate the production of EV
subtypes by altering EV biogenesis or cargo sorting.
Open question: can we specically control EV
formation?
EV biogenesis involves a complex interplay of genetic, metabolic,
and environmental factors. This complexity raises pivotal ques-
tions about the specicity and universality of EV production me-
chanisms. 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
Extracellular vesicles | 11
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 proper-
ties 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 head-
groups. PLD1 converts the headgroup of PC lipids to smaller PA li-
pids, and neutral sphingomyelinases convert sphingomyelins to
smaller ceramides on the cytosolic face of the endosome mem-
brane 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 corre-
lates with increased EV production in C. elegans embryos with dis-
rupted PE asymmetry (Wehman et al. 2011, Beer et al. 2018),
making it difcult 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 en-
hances 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 signicant. In
cancer cells, stiff ECMs signicantly enhance exosome secretion
(Wu et al. 2023), underscoring the intricate relationship between
mechanical forces and EV release. In C. elegans, mechanical pres-
sure 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 pro-
duction (Melentijevic et al. 2017; Cooper et al. 2021), and proteosta-
sis 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 termin-
ate 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
benet structurally or metabolically from EV cargo. The EV mem-
brane 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 specic 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 ectocyto-
sis. To allow the cytoskeleton and membrane trafcking to return
to homeostasis, the bridge is released as a large EV after the con-
nections 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
N
Exopher
d
Neurons extrude dysfunctional organelles and
toxic proteins in exophers
Damaged
mitochondrion
Healthy
mitochondrion
Cell body
Aggresome
Spermatozoon
Protease-activated
spermatid Mitopher
c
Spermatids extrude excess mitochondria in EVs
Actin
Filaments
Ciliary
tip
Ciliary base
b Cilia maintain their composition by shedding excessive
membrane and proteins into EVs from the tip and base
ESCRT
MBR
Cell Cell
ESCRT
EV
a Cells use small EVs to break up midbody remnants
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
trafcking 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.
12 | J. Wang et al.
electron tomograms forming thin protrusions from the intercellu-
lar bridge, and small fragments of the bridge can be taken up be-
fore 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.
EVs regulate sensory homeostasis and plasticity
Neuronal EV shedding plays a conserved role in regulating both
ciliary proteostasis and structural stability of neuronal protru-
sions. In mammalian cells, primary cilia eliminate accumulating
G protein-coupled receptors (GPCRs) in EVs via actin-mediated cil-
iary tip ectocytosis (Nager et al. 2017). In C. elegans, ciliary EV shed-
ding 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 degen-
eration and resorption. The shedding of the ciliary tip as an EV
hastens the removal of the cilium during cilia resorption in cul-
tured 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-
1is essential for ciliary axoneme stability; mutants of ccpp-1 ex-
hibit 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 compos-
ition 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 micro-
villi (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 mei-
osis 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 sperma-
tids 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 sperma-
tids and helps enrich mitochondria in spermatids. In nmy-2 mu-
tants, 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 trafcking 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 or-
ganelle 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 mito-
chondrial DNA (Abdu et al. 2016; Schwartz et al. 2022). This pro-
vides 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 neighbor-
ing endoderm. This depletion of nutrients could be advantageous
for a cell type that will be quiescent until nutritional signaling in-
duces 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 signicantly enriched in EVNs and promotes
PKD-2 ciliary EV biogenesis (Wang et al. 2015). Proteostatic stress
directly inuences exopher formation in touch receptor neurons
(Melentijevic et al. 2017), and systemic stress signaling affects exo-
pher production (Cooper et al. 2021). EV release bears similarity to
retinal photoreceptor renewal, which involves the nightly shed-
ding 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 concen-
trate 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 exopher-
genesis compromises touch sensitivity in neurons expressing
neurotoxic proteins. Exopher uptake and endolysosomal degrad-
ation by the neighboring epidermis likely allow the neuron to fo-
cus on its signaling function by outsourcing degradation (Wang ,
Arnold, et al. 2023). However, the transfer of aggregates in exo-
phers 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 mea-
sured by confocal uorescence microscopy. However, uores-
cence intensity can skew apparent sizes and the diameters
would be better evaluated by EM. The observation that dopamin-
ergic neurons can respond to environmental toxins by releasing
Extracellular vesicles | 13
EVs may be relevant to EV contributions to the degenerative pro-
pensity 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 clas-
sical 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 modi-
cations (Parchure et al. 2018). For example, Wnts are palmitoy-
lated 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 hydro-
phobic 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 extracellu-
larly (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 regu-
late cuticle formation during development (Fig. 10a). The epider-
mis 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 abun-
dant 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.
The rst embryonic midbody remnant has also been implicated
in dorsoventral patterning (Singh and Pohl 2014). Cortical ows
move the P0 midbody remnant toward the future ventral side of
the two-cell embryo. Astral microtubules in the dividing P1 spin-
dle contact the cell cortex neighboring the P0 midbody remnant.
The contact to the midbody remnant appears to instruct P1 spin-
dle 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 mechan-
ical 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 re-
pair 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 pro-
teins that act as homotypic or heterotypic fusogens during devel-
opmental cell–cell fusion events (Mohler et al. 2002; Sapir et al.
2007). While EFF-1 acts cell autonomously in PVD, AFF-1 is ex-
pressed in epidermal seam cells, and its expression is upregulated
in response to dendrotomy (Oren-Suissa et al. 2017). Seam cells re-
lease EVs containing AFF-1, although it is unclear whether the
AFF-1 EVs are derived from the plasma membrane, lopodia, or
endolysosomal organelles. Intriguingly, AFF-1 EVs can fuse with
c
Muscle exophers feed yolk to oocytes
Muscle cell
Mito
N
Mito
Yolk
VIT-2
Exopher
Oocytes
Embryos
Body wall muscle
Dying
cell
PS+
EVs
Neighboring
cell
Engulfing
cell
d
Apoptotic cells release EVs to induce phagocytosis
b
Seam cell EVs reconnect neuronal dendrites
Dendritic arbors Soma
Seam cell
PVD
AFF-1
EVs
Alae
Exosomes
Epidermal cell
Cuticle
a
EVs carry hedgehog-like proteins for
alae formation
WRT-2/8
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 leaet of the plasma membrane and release small PS-positive EVs that facilitate PS exposure on
neighboring cells and promote phagocytic clearance.
14 | J. Wang et al.
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 fu-
sogen 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 zeb-
rash, 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 princi-
pal intestinal yolk protein (reviewed in Perez and Lehner 2019), dou-
bles 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, correl-
ating 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 tis-
sues. 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 ap-
pearance of low levels of PS on the surface of the engulng 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 ex-
posure on the engulng cell and disrupting corpse clearance, sug-
gesting that the PS-binding probe blocks both PS function and EV
clearance. This implies that PS needs to be recognized by the en-
gulng cell for the cell to expose low levels of PS and phagocytose
the cell corpse. Indeed, increasing PS exposure on engulng
cells by disrupting the PS ippase TAT-1 can increase phago-
cytosis 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 pro-
motes the clearance of the PS-positive EVs (Mapes et al. 2012), al-
though 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 sig-
naling for corpse engulfment, although it is difcult 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 trans-
fer EVs among individuals, thus serving as a useful model for illus-
trating the fundamental functions and mechanisms of EVs in
mediating inter-organismal communication. Caenorhabditis ele-
gans males possess 27 EVNs with cilia whose tips prolically
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 specic 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 be-
haviors could be regulated by EVs. EVN cilia are likely specialized
devices for the release of signaling EVs, a secretory function of ci-
lia that has yet to be fully recognized (Luxmi and King 2024).
Males can transfer EVs during mating and ejaculation. In many
animals, EVs produced by the epididymis or found in the seminal
uid 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, sperm-
atozoa 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 me-
chanisms 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
b
Males transfer different types of EVs to hermaphrodites during mating
a
Isolated EVs induce changes in male locomotory behavior
MSP-carrying EVs induce ovulation
Ciliary EVs
Ciliary EVs
No EVs:
Sinusoidal movement
EVs trigger backwards
tail chasing behavior
Isolated EVs
ENPP-1 EVs
Spermatids
Spermatozoa
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 uid containing ENPP-1, and sperm-derived
EVs carrying MSP that induce ovulation.
Extracellular vesicles | 15
include nematode and atworm 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 hel-
minths a major economic and health burden. Parasitic EV cargos
can modify host biological processes, including immune re-
sponses and cellular metabolism (Eichenberger et al. 2018;
Gazzinelli-Guimaraes and Nutman 2018; Ressel et al. 2019).
Consequently, understanding the molecular composition, biogen-
esis, and cargo sorting mechanisms of helminth EVs holds signi-
cant 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 sever-
ity of infections (Mu et al. 2021). Moreover, employing biocontrol
strategies and harnessing helminth EVs present a promising av-
enue 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 gen-
omic 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), understand-
ing ciliary EVs released by C. elegans males could provide vital
clues about how sensory cilia in parasitic nematodes release sig-
naling 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 nema-
tologists could signicantly 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 proling has identied >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 tis-
sue 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 move-
ment (Zhai et al. 2024). Whether migrasomes are produced by mi-
grating 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 patho-
genic bacteria triggers germline expression of the Cer1 retrotrans-
poson, 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 func-
tions in C. elegans.
In addition to potential bacterial regulation of intestinal EVs,
the interaction between C. elegans and microbial EVs in its micro-
biome or environment is poorly understood. The most common
proteins identied in C. elegans EV proteomic proles are from out-
er 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
uids (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 efcacy. For ex-
ample, 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 ap-
proaches to conrm the presence and activity of EVs. The MISEV
guidelines also summarize common challenges in EV research, in-
cluding challenges in labeling EVs and confusion of non-EV parti-
cles 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
conrm 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 xation 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 combin-
ation with uorescent approaches for correlated light and elec-
tron 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
16 | J. Wang et al.
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 ser-
ies 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).
Furthermore, EM is compatible with different stains that can
provide contrast to the EV membrane or to EV contents. For ex-
ample, 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
tat-5 RNAi 100 nm 50 nm
EVs
Cell 1
Cell 2
c d
500 nm
EV
EV
EV
b
500 nm
EV
Cell 1
Cell 2
MVB
Mito
a
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 magnied region in d shows the cross section of a plasma membrane bud. Data were collected for Wehman et al. (2011).
Extracellular vesicles | 17
the high resolution of TEM with the specic 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
trafcking 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 xed tissues; the
gain in spatial insights comes at the cost of temporal dynamics.
Fluorescent imaging of EVs
EVs are often rst suspected through uorescent imaging ap-
proaches, where punctate uorescence of a reporter protein is ob-
served to move away from the source cell (Table 2). Fluorescent
reporters allow the tracking of EV dynamics, including EV target-
ing to specic tissues, EV uptake, and intracellular breakdown of
EV cargos (Verweij et al. 2021). EVs are often labeled with uores-
cent 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 car-
gos used to track environmental release of different subpopula-
tions of ciliary EVs (Fig. 13a-b) (Wang et al. 2014; Clupper et al.
2022; Walsh et al. 2022). However, overexpression of uorescent
10 µm
Ray 1 234567 8 9
a
Cilium tip
Cilium shaft
Cilium base
Ciliary EVs
LOV-1::mSc PKD-2::GFP
b
5 µm
3 µm
10 µm
pad-1(babIs1)
mCh::PH::ZF1 degron 15-cell
PB
e
f
NMY-2::GFP::ZF1
mCh::PH
2-cell4-cell
10 µm
c
d
Fig. 13. Visualization, characterization, and tracking of EVs using uorescence 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 specic 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).
18 | J. Wang et al.
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 (re-
viewed in Nance and Frøkjær-Jensen 2019). It is also important
to verify that the uorescent tag does not alter the function of
the cargo proteins, the EV size, or abundance using functional as-
says, EM, or additional reporters.
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 clear-
ance 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 punc-
ta labeled by a uorescent reporter are in fact EVs. Care should
also be taken to distinguish reporter uorescence from autouor-
escence, which can arise from the worm, debris from microscope
slides, culture media, or bacterial food sources. Small EVs are
rarely visible under bright-eld imaging, which can help distin-
guish EVs from larger autouorescent debris or bacteria.
One challenge with uorescent labeling of EVs is the limit in
spatial resolution. Fluorescent reporters will change their appar-
ent width based on abundance and the resulting changes in uor-
escence intensity, making it difcult to determine EV size based
on uorescent 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 uorescence of
the plasma membrane if both are labeled. For example, the PH do-
main 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 uores-
cent 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 reso-
lution 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 con-
ventional confocal microscopy (Verweij et al. 2021; Razzauti et al.
2023). Super-resolution imaging signicantly enhances the visual-
ization 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. ele-
gans EV preps with E. coli outer membrane vesicles (Nikonorova
Cushion
d
Ultracentrifugation & Gradient Fractionation
Gradient
EV fractions
Non-EV fractions
Non-EV fractions
e
Size-exclusion
chromatography
EV fractions
Non-EV fractions
Cushion
c
Differential Ultracentrifugation
a
Culture and collection
b
Differential Centrifugation
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.
Extracellular vesicles | 19
et al. 2022). It is straightforward to computationally distinguish
worm proteins and RNAs from bacterial sequences after analysis;
this requires increased depth to capture sufcient C. elegans data
from MassSpec or sequencing data of co-isolated EVs.
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 iso-
lated after long periods in axenic M9 buffer; the abundant cytosol-
ic 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 inuence EV subtypes, abundance, cargos, stability, and
thereby downstream functions. To facilitate comparisons be-
tween 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 sam-
ples (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 frag-
ments induced by tissue damage. Therefore, while homogeniza-
tion 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 sufcient purity from co-isolates and obtain-
ing 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 centri-
fuge 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 ultracentri-
fugation 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 separ-
ation of distinct subpopulations of EVs (Fig. 14d), as well as enab-
ling 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 uorescently 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 chromatog-
raphy, which separates EVs based on size without an ultracentri-
fuge. 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 chromatog-
raphy 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 de-
rived from different organelles, cell types, and organisms.
Characterization of isolated EVs
Given the broad diversity of EVs and non-vesicular particles se-
creted 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 uids, 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 in-
clude outer membrane vesicles from bacteria and particles such
as viral capsids (Moore et al. 2021; Nikonorova et al. 2022). To
conrm 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 fur-
ther 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 specic uores-
cent cargos or other membrane markers (Wang et al. 2014;
Nikonorova et al. 2022). Western blots could also be used to char-
acterize 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 specic anti-
bodies 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 pro-
ling 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 data-
sets 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 bio-
active molecules and lipidomics have yet to be performed on C. ele-
gans EVs, but EVs can also be articially loaded with desired cargos
(Shao et al. 2023), allowing their study for therapeutic delivery.
0
5
10
15
20
25
30
0 10 20 30 40
EV cargos
enriched in cell X
(IL2 neurons)
EV cargos
enriched in cell Y
(Ciliated neurons)
Environmental EV cargos
enriched in cell types X & Y
(IL2 & other ciliated neurons)
Enrichment in Sample X
Enrichment in Sample Y
tsp-6
tbb-4
clhm-1
sid-2 asic-2
cil-7 tba-6
klp-6
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-specic EV cargos
(Cao et al. 2017; Nikonorova et al. 2022). MyEVome is currently available
at https://myevome.shinyapps.io/evome-app.
20 | J. Wang et al.
In-depth characterization and exploitation of EV cargos could
yield further insights into EV biogenesis and function.
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 pseudo-
coelom (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 ex-
tended 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 mu-
tant strains or pre-treating EVs can also help identify the EV sub-
population or cargo that is having a functional impact. For
example, environmental EVs isolated from klp-6 and tba-6 mu-
tants 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 com-
parability. Another common control is to use detergent treatment
to disrupt the lipid membrane of EVs and conrm whether the
function of an EV sample is lost. Detergent treatment can also
be used in combination with protease or nuclease protection as-
says 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 environ-
mentally released EVs.
Open question: overcoming gaps in EV techniques
One challenge for EV research compared to most intracellular or-
ganelles is the lack of a pan-EV marker to enable EV labeling from
any tissue. EVs typically reect the composition of their source or-
ganelle and cell type, which means that EV studies often necessi-
tate 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 character-
ization of environmental EVs with single-cell proteomic studies is
likely to identify new reporters that can be applied to new EVs, an
approach simplied 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 con-
taining male and hermaphrodite C. elegans, while the single-cell
transcriptomic proling dataset was obtained from a synchro-
nized L2 hermaphrodite population. Consequently, candidate EV
cargos should be validated further. Developing additional EV
markers from different tissues will greatly enhance our under-
standing of EV biogenesis, diversity, and function in C. elegans.
Another challenge with EVs is obtaining adequate uorescent
signal from EV markers to monitor EV biology in the context of or-
ganismal uorescence. 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 uorescent 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 trafcking
routes into EVs, the release and movement of EVs, EV–cell inter-
action 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 dee-
per proling 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 de-
fective 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 rst investigate whether bacterial outer
membrane vesicles (OMVs) inuence C. elegans physiology, behav-
ior, or EV production. Alternatively, chemically dened culture
media that support normal C. elegans development without bac-
teria would also be an option (Zec
ic et al. 2019), which could also
allow the dissection of metabolite inuence on EV biogenesis. It
may also be possible to use microuidic devices to characterize
EV diversity without isolating C. elegans EVs from cells and bacter-
ial 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 char-
acterization 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 pro-
tein to a biotin ligase to post-translationally modify nearby pro-
teins (Löf et al. 2017; Kaneda et al. 2021; Li et al. 2023). The
biotinylated proteins can then be isolated through biotin–strepta-
vidin interaction, reducing the number of non-EV proteins to be
identied. As different EV cargos can be labeled, this allows the
characterization of specic 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 en-
vironmental EVs. C. elegans EV research has illustrated conserved
pathways in EV biogenesis, uptake, and clearance in addition to
EV functions regulating tissue homeostasis, regeneration, devel-
opmental signaling, and social interactions. Studies to date
highlight the power of the C. elegans model in elucidating funda-
mental 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 hetero-
geneity across the whole organism. At the same time, conserved
themes in EV biogenesis pathways reveal that EVs are integral
Extracellular vesicles | 21
extracellular organelles with key roles in cellular processes.
C. elegans is poised to lead the way in exciting areas of EV re-
search, including the fundamental mechanisms governing
how, when, and what types of EVs cells produce, as well as un-
earthing 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).
Conicts of interest
The author(s) declare no conicts of interest.
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Editor: I. Greenwald
28 | J. Wang et al.
