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Mechanisms Shaping the
Membranes of Cellular
Organelles
Yoko Shibata,1 Junjie Hu,2 Michael M. Kozlov,3
and Tom A. Rapoport1
1
Howard Hughes Medical Institute and Department of Cell Biology, Harvard Medical
School, Boston, Massachusetts 02115; email: yoko [email protected],
tom [email protected]
2
College of Life Sciences, Nankai University, 300071 Tianjin, China;
email: [email protected]
3
Department of Physiology and Pharmacology, Sackler Faculty of Medicine,
Tel Aviv University, 699978 Tel Aviv, Israel; email: [email protected]
Annu. Rev. Cell Dev. Biol. 2009. 25:14.1–14.26
Key Words
The Annual Review of Cell and Developmental
Biology is online at cellbio.annualreviews.org
endoplasmic reticulum, mitochondria, caveolae, tubules, reticulons,
dynamins
This article’s doi:
10.1146/annurev.cellbio.042308.113324
c 2009 by Annual Reviews.
Copyright All rights reserved
1081-0706/09/1110-0001$20.00
Abstract
Cellular organelles have characteristic morphologies that arise as a result of different local membrane curvatures. A striking example is the
endoplasmic reticulum (ER), which consists of ER tubules with high
curvature in cross-section, peripheral ER sheets with little curvature
except at their edges, and the nuclear envelope with low curvature except where the nuclear pores are inserted. The ER may be shaped by
several mechanisms. ER tubules are often generated through their association with the cytoskeleton and stabilized by two families of integral
membrane proteins, the reticulons and DP1/Yop1p. Similar to how curvature is generated in budding vesicles, these proteins may use scaffolding and hydrophobic insertion mechanisms to shape the lipid bilayer
into tubules. In addition, proteins of the dynamin family may deform
the ER membrane to generate a tubular network. Mechanisms affecting
local membrane curvature may also shape peripheral ER sheets and the
nuclear envelope, as well as mitochondria and caveoli.
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Contents
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INTRODUCTION . . . . . . . . . . . . . . . . . . 14.2
GENERATION OF CURVATURE
DURING VESICLE BUDDING . . 14.6
GENERATION OF THE TUBULAR
ER NETWORK . . . . . . . . . . . . . . . . . . 14.8
GENERATION OF MEMBRANE
SHEETS IN THE ER . . . . . . . . . . . . .14.12
SHAPING MITOCHONDRIA. . . . . . .14.14
GENERATING THE CURVATURE
OF CAVEOLAE . . . . . . . . . . . . . . . . . .14.17
CONCLUSIONS . . . . . . . . . . . . . . . . . . . .14.19
INTRODUCTION
Membrane-bound organelles in eukaryotic
cells have characteristic shapes. Some organelles, such as lysosomes and peroxisomes,
are relatively spherical, but others are more
complex. For example, mitochondria form a
tubular network, the endoplasmic reticulum
(ER) is an interconnected system of sheets and
tubules, and the Golgi apparatus consists of a
stack of perforated sheets. Cellular membrane
shapes are generally dynamic. They often
change during processes such as cell movement, division, and growth, but also under
steady state conditions. For example, many
membrane-bound compartments have the ability to bud small vesicles and to fuse with vesicles;
that is, they communicate with one another
by vesicular trafficking. Membrane shaping
relates to the generation of membrane curvature (McMahon & Gallop 2005, Zimmerberg
& Kozlov 2006). For example, high curvature
is seen in cross-sections of narrow tubules, at
the edges of closely spaced membrane sheets,
or with small transport vesicles. Conversely,
low curvature membranes include the sheet
structures of the ER or Golgi. Intracellular
membranes can change their shapes without
altering their basic topology. For example,
a flat membrane may undergo bending, or a
tubule may be drawn out of a flat membrane.
Alternatively, the membrane shape can change
ER: endoplasmic
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topologically by remodeling, a process that
occurs most often by fusion or fission. During
fusion two separate membranes merge into
one, whereas during fission a continuous membrane is divided into two separate ones. For
example, ER tubules can fuse with an existing
tubule to form a three-way junction. How the
characteristic shape of an organelle is achieved
and how membrane curvature is generated are
still only poorly understood, but these questions are now central to a new research field.
Here, we will concentrate on the mechanisms
of ER morphogenesis but point out that other
organelles are shaped by similar principles.
The mechanisms by which biological membranes are shaped and remodeled can be described by physical terms. The shape of a membrane at a given point of its surface can be described by two principle curvatures, c1 and c2 ,
which are the inverse of the radii of two perpendicular arcs lying on the membrane plane, R1
and R2 (c1 = 1/R1 and c2 = 1/R2 ; see Figure 1,
which also gives a more detailed definition of
curvature). The curvature is considered to be
small if the radii are larger than the thickness of
the membrane d, R1 d, R2 d. It is considered
to be large when one of the radii is comparable
with or somewhat larger than the membrane
thickness, R1 or R2 ≥ d. Taking into account
that the lipid bilayer thickness is d ≈ 4 nm, intracellular tubules or vesicles with radii of ∼25–
30 nm have relatively high curvatures.
Lipid bilayers tend to remain flat because the
generation of curvature requires energy. The
energy cost for bending a bilayer can be determined from the elastic model of lipid membranes (Helfrich 1973, 1990) (see Figure 1).
Generation of a fragment of a lipid tubule with
a diameter of 60 nm and a length of 60 nm
requires ≈70 kcal/mol, whereas formation of
a spherical vesicle of the same diameter requires ≈300 kcal/mol (see Figure 1 legend).
Both energies are much larger than the characteristic thermal energy (≈0.6 kcal/mol), which
means these shapes cannot be generated simply
by thermal fluctuations. However, these bending energies can be generated by the hydrolysis
of a few tens of adenosine triphosphate (ATP)
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Curvature
Curved shapes
Cylinder
Saddle
COPII-coated vesicles
Yop1p-generated
membrane tubules
Neck of budding vesicle
R2
R1
20 nm
c1 = R
c2 = R–2
–1
c1 = c2 = R
J = 2(R–1)
K = R–2
–1
100 nm
c1 = R
J = R–1
–1
c2 = 0
K=0
c1 = –c2 = R–1
J=0
K = –R–2
Figure 1
Curvature and shapes of membranes. A membrane is represented by the mid surface between the two lipid monolayers (left panel ). The
shape of every small element of the mid surface is described by two perpendicular arcs of radii R1 and R2 . The curvatures of the arcs,
c 1 = R1 and c 2 = R1 are referred to as principle curvatures of the surface and fully characterize the local shape of the membrane. More
1
2
commonly, two combinations of the principle curvatures are used: the total curvature, J = c 1 + c 2 , and the Gaussian curvature,
K = c 1 c 2 (Helfrich 1973). The other panels show three basic shapes adopted by biological membranes and electron microscopy images
of actual examples (Image of COPII vesicle, courtesy of L. Orci and R. Schekman; image of neck of a budding vesicle, from Gallop and
McMahon 2005). The formulas show the relationship between the curvatures and the radii. Generation of membrane curvature
requires energy, which can be computed using the Helfrich theory of membrane elasticity (Helfrich 1973, 1990). The bending energy
2
per unit area of the membrane, fB , is expressed through the total, J, and Gaussian, K, curvatures by f B = 12 · κ B · J + κ̄ · K , where κ B ≈
−19
Joule ≈ 10 kcal/mol is the lipid bilayer bending modulus, and κ̄ is the modulus of the Gaussian curvature (the latter has not been
10
measured directly but can be estimated to be in the range −κ B < κ̄ < 0). To compute the total bending energy, Ftot , the energy per unit
area has to be integrated over the entire membrane surface, Ftot = f B d A. It should be noted that the total curvature J determines the
energy of both membrane shaping and remodeling, whereas the energy of the Gaussian curvature K is only important for remodeling.
For example, the energy of the Gaussian curvature of a spherical vesicle does not change when the vesicle is arbitrarily deformed, but it
changes by 4π κ̄ for splitting a vesicle into two and by −4π κ̄ for fusion of a vesicle with a planar membrane. Based on the Helfrich
cyl
model, the bending energy of a cylindrical membrane of radius R and length L equals Ftot = π κ B RL , and for a spherical membrane it is
= 8π κ B + 4π κ̄.
molecules (each ≈ 10 kcal/mol), consistent with
the notion that metabolically generated energy can be used to shape biological membranes. These considerations indicate that high
membrane curvature requires molecular mechanisms that generate and stabilize these energetically unfavorable states. These mechanisms
are based on the interplay between lipids and
proteins and require regulation to achieve the
dynamic behavior of cellular membranes.
In principle, lipids alone can generate
membrane curvature if the lipid composition
of the membrane monolayers is asymmetric.
For example, a nonbilayer lipid species could be
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Midsurface element
Sphere
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Membrane mid
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between the two
monolayers of a
membrane-
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Molecular motors:
ATPases that move
associated cargo along
microtubules (includes
kinesins and
cytoplasmic dynein) or
actin filaments
(myosins)
Amphipathic helix:
has distinct
hydrophobic and
hydrophilic surfaces
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concentrated in one of the membrane monolayers. Nonbilayer lipids, such as diacylglycerol
or lysophospholipids, tend to self-assemble
into extremely bent monolayers with curvature
radii close to the lipid monolayer thickness,
R ≈ d/2 (for review, see Zimmerberg & Kozlov
2006). When mixed with regular lipids in a
membrane and asymmetrically distributed
between the membrane monolayers, these
lipids can produce high curvature of the whole
bilayer. Lipid asymmetry can also lead to a
difference in membrane area between the two
monolayers. In this case, curvature is generated
because the two monolayers are practically
nonstretchable and noncompressible, but are
coupled to one another along the membrane
mid surface [bilayer-coupling mechanism
(Sheetz & Singer 1974)]. In agreement with
these predictions, it has been found experimentally that high mole fractions of some lipids can
deform liposomes into tubules (Stowell et al.
1999), and certain lipids can segregate and lead
to buds and domains of moderate curvature
(Bacia et al. 2005, Baumgart et al. 2003).
However, theoretical calculations show that
the generation of strongly bent membranes
with curvature radii of a few tens of nanometers
requires fairly strong monolayer asymmetry
(Zimmerberg & Kozlov 2006), which is not
expected based on the lipid composition of cell
membranes (van Meer et al. 2008). Therefore,
the current consensus in the field is that high
membrane curvature in intracellular membranes is generated by proteins. Nevertheless,
lipids play an important role, modifying and facilitating the function of curvature-generating
proteins in decisive ways (Devaux et al. 2008).
Proteins could cause curvature indirectly by
generating lipid asymmetry. For example, they
could produce nonbilayer lipids in one of the
membrane monolayers, or they could transfer
lipids from one bilayer leaflet to another (flippases). However, most of the evidence to date
suggests that proteins generate membrane curvature in a direct manner, rather than indirectly
through lipids.
Proteins appear to use three major, but
not mutually exclusive, mechanisms to directly
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generate high membrane curvature (McMahon
& Gallop 2005, Zimmerberg & Kozlov 2006)
(Figure 2). The first mechanism involves proteins that exert mechanical force on a lipid bilayer (Figure 2a). Thin membrane tubules can
be pulled out of low-curvature membrane reservoirs by molecular motors as they move along
microtubules or actin filaments. Alternatively,
membrane tubules can be pulled when they
attach to polymerizing microtubules or actin
filaments. According to theoretical estimates,
tubules of 30 nm radii require a pulling force
of 20 pN, which can be generated by about
10 molecular motors (Zimmerberg & Kozlov
2006).
The second mechanism by which proteins
bend membranes is scaffolding (Figure 2b).
Here, a protein or protein network has a highly
curved surface with a strong affinity for the lipid
polar head groups. The scaffolding proteins interact with a lipid membrane along the bent
surface and force the bilayer to adopt the same
curvature. This requires that the rigidity of the
protein scaffold is high compared to the membrane bending rigidity, and that the energy of
the membrane-protein attachment exceeds the
membrane-bending energy.
The third mechanism of membrane bending
is hydrophobic insertion (wedging) (Figure 2c).
Here, proteins insert hydrophobic domains,
such as the hydrophobic surface of an amphipathic helix, into the upper part of a membrane
monolayer. This results in the perturbation of
the packing of the lipid head groups and generates local membrane curvature. According to
theoretical calculations, maximal curvature is
produced if the hydrophobic insertion is shallow, penetrating the membrane to about one
third of the lipid monolayer thickness, that is,
to approximately the interface between the polar head groups and the hydrocarbon chains
(Campelo et al. 2008).
As discussed below, some integral membrane proteins have also been proposed to generate curvature by a combination of scaffolding
and hydrophobic insertion (wedging).
Protein embedding alone could cause membrane curvature by the bilayer coupling effect
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if a membrane protein displaces lipids primarily in one monolayer. Because the effect would
be averaged over the total membrane surface, a
tubule with a curvature radius of 30 nm would
require that these proteins generate an overall area difference of 13% between the monolayers. While this suggests that most integral
membrane proteins can only cause high curvature when they are extremely abundant, it is
conceivable that some membrane proteins displace lipids specifically at a certain depth of the
lipid bilayer and thus generate local curvature
by the hydrophobic insertion mechanism, similar to how amphipathic helices act, or that they
combine hydrophobic insertion (wedging) and
scaffolding effects in a cooperative manner.
Membrane remodeling by fusion and fission is tightly related to membrane shaping (Chernomordik & Kozlov 2005, 2008;
Chernomordik et al. 2006). During fusion two
lipid bilayers come in close contact with one another, leading initially to the fusion of the approaching monolayers (hemi-fusion), followed
by fusion of the outer monolayers (Figure 3).
The first step in a fusion reaction is to bring the
two membranes into close proximity. For example, the fusion of transport vesicles with their
target membrane is caused by the association
of SNARE proteins; three helices of SNARE
proteins in one membrane, all with their Ntermini in the cytosol, zipper together with one
helix of the same orientation in the other membrane to force the bilayers into close proximity
(Sudhof & Rothman 2009). The actual fusion
process may be initiated by the generation of a
a
le
u
Tub
ule
tub t
lls amen
u
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oto ton
ar mskele
l
u
lec yto
Mo ong c
al
Tubule binds to polymerizing
cytoskeleton filament
Tubule
−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−→
Figure 2
Three major mechanisms by which proteins
generate curvature. (a) Membrane deformation
by proteins that exert mechanical force. In the cases
shown, tubules are pulled out of a flat membrane by
either being attached to molecular motors (blue) that
move along cytoskeleton fibers, or by binding to
the tip of a polymerizing cytoskeleton filament. The
directions of movement of the molecular motor,
the polymerizing microtubule, and the membrane
tubules are indicated by arrows. (b) Curvature generation by scaffolding proteins. Scaffolding proteins
have a rigid curved shape, whose concave surface
interacts with the membrane, forcing the bilayer to
adopt the same curvature. (c) Curvature generation
by the hydrophobic insertion (wedging) mechanism.
Soluble proteins insert hydrophobic regions into a
membrane monolayer. Shown in blue and red are the
hydrophilic and hydrophobic parts of an amphipathic
helix, respectively, in cross section. Note that shallow
insertion of the hydrophobic surface of the helix
generates maximal curvature. Integral membrane
proteins that occupy more space in one leaflet of the
membrane than the other could also generate membrane curvature by the bilayer-coupling mechanism.
b
Scaffolding
proteins
c
Amphipathic
helix
Integral membrane
protein
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a
Membrane fusion pathway
b
Membrane fission pathway
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Figure 3
Pathways of membrane remodeling by fusion and fission. (a) A fusion pathway starts with the generation of a strong local deformation
of one membrane. A protrusion or dimple is formed, which establishes a point-like contact with the target membrane. Relaxation of the
protrusion drives fusion stalk formation and subsequent hemifusion. Decay of the hemifusion intermediate into a fusion pore completes
membrane fusion. (b) Membrane fission begins with the splitting of the inner monolayer of a membrane neck, which results in a
hemi-fission structure similar to that of a fusion stalk. Decay of this structure drives separation of the membrane neck into two
nonconnected membranes.
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point-like protrusion or a strongly curved dimple in one of the membranes (Efrat et al. 2007,
Kozlov & Chernomordik 1998, Martens et al.
2007) (Figure 3). The generated dimples would
point to the adjacent membrane (Figure 3) and
promote close membrane contact. Fusion of the
dimple with the target membrane would be favored by the accompanying relaxation of the
strongly curved shape of the dimple.
Similarly, the first step of fission is hemifission, where only one monolayer divides,
which is then followed by splitting of the second
monolayer (Kozlov & Chernomordik 2002,
Kozlovsky & Kozlov 2003). Membrane fission
may be driven by the relaxation of a highly
curved funnel-like shape, generated in the thin
necks that connect the dividing membranes
(Kozlovsky & Kozlov 2003) (Figure 3). The
generation of protrusions and dimples in fusion
and of narrow necks in fission requires energies
of tens of kilocalories per mole (Efrat et al. 2007,
Kozlov & Chernomordik 1998, Martens et al.
2007), which must be provided by proteins.
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These considerations show that both membrane shaping and remodeling employ proteins
that generate high membrane curvature.
GENERATION OF CURVATURE
DURING VESICLE BUDDING
Most of the principles by which membranes are
shaped and remodeled were discovered in studies on clathrin-mediated endocytosis, where a
small vesicle with a highly curved membrane
is generated from the essentially flat plasma
membrane (for review, see Brodsky et al. 2001,
Higgins & McMahon 2002). Clathrin forms
cages around a budding vesicle, but it does not
form a rigid scaffold and does not interact directly with the membrane (Kirchhausen 2000).
Therefore, several other components generate
most of the curvature required for vesicle budding, employing hydrophobic insertion (wedging) and scaffolding mechanisms.
Hydrophobic insertion is often mediated
by amphipathic helices. Examples include the
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N-terminal helix of epsin (helix zero of the
ENTH domain) (Ford et al. 2002), the amphipathic N-terminal helices of the BAR domain–
containing proteins amphiphysin (Peter et al.
2004) and endophilin (Gallop et al. 2006,
Masuda et al. 2006), and the amphipathic helix between the PX and BAR domains of Sorting Nexin 9 (Pylypenko et al. 2007). The lipidinserted polypeptide segments are often flexible
and unstructured in solution and adopt a helical
conformation only upon binding to a lipid bilayer (Gallop et al. 2006). The amphipathic helices of the ENTH and N-BAR domains penetrate the lipid monolayers to a depth that is most
effective in generating local curvature (Gallop
et al. 2006); calculations show that with only
a few mole percent of these proteins, a curvature of a few tens of nanometers radius can be
generated (Campelo et al. 2008).
The scaffolding mechanism is exemplified
by N-BAR domains. These form bananashaped dimers, whose concave surface contains positive charges that interact with the
head groups of the phospholipids (Li et al.
2007, Peter et al. 2004, Pylypenko et al. 2007,
Weissenhorn 2005, Zhu et al. 2007). They can
form curvature radii of 11–14 nm. Another
type of BAR domain, the F-BARs, uses the
scaffolding mechanism to generate membrane
structures of lower curvature (∼30 nm radius)
(Henne et al. 2007, Shimada et al. 2007). They
can form oligomeric spirals of different pitch
through tip-to-tip and lateral interactions to
generate tubules of variable diameters (Frost
et al. 2008). The BAR domains are rigid, a prerequisite to impose their curvature onto the interacting bilayer. Interestingly, while all these
BAR domains generate positive curvature, yet
another type of BAR domains, the I-BARs, have
convex surfaces and induce negative curvature
in cellular protrusions (Saarikangas et al. 2009).
The fission of a vesicle from the plasma
membrane is mediated by the dynamin-1 protein (Kosaka & Ikeda 1983). Dynamin-1 is a
GTPase that also contains a lipid-interacting
PH domain and a proline-rich domain with
which it interacts with other curvatureinducing proteins, such as amphiphysin and
endophilin (Praefcke & McMahon 2004).
Dynamin-1 forms short spirals around the neck
of a budding vesicle that continuously form
and disassemble during the GTP hydrolysis cycle (Bashkirov et al. 2008, Pucadyil & Schmid
2008, Ramachandran & Schmid 2008). It was
suggested that upon GTP hydrolysis, the dynamin helix is released from the lipid surface but
transiently maintains a rigid frame around the
membrane neck, preventing it from swelling
(Bashkirov et al. 2008); the elastic stress accumulated within the membrane neck would then
relax by hemi-fission followed by fission of the
neck.
Most curvature-inducing proteins use a
combination of hydrophobic insertion (wedging) and scaffolding mechanisms. For example,
N-BAR proteins insert the N-terminal amphipathic helix into the lipid bilayer and use their
banana shape to scaffold the curved membrane.
Similarly, dynamin-1 inserts a loop of its PH
domain into the bilayer and forms a spiral scaffold around the neck of the budding vesicle
(Hinshaw & Schmid 1995, Ramachandran &
Schmid 2008, Takei et al. 1995).
Endocytosis appears to be facilitated by the
actin cytoskeleton. In fact, some curvaturegenerating proteins, such as the BAR proteins,
interact with actin (Itoh et al. 2005, Tsujita et al.
2006), suggesting that actin may facilitate membrane deformation or provide mechanical force
for vesicle fission (Engqvist-Goldstein & Drubin 2003). Taken together, the number of proteins involved in endocytosis is bewildering, and
it is still unclear how they cooperate to generate
curvature during vesicle formation and fission.
The same principles for generating
membrane curvature, hydrophobic insertion
(wedging) and scaffolding, apply to the budding
of vesicles from the ER and Golgi. However,
compared to the formation of an endocytic vesicle, only a small number of different proteins
is required. The small GTP binding proteins
Sar1 and Arf1 first use the hydrophobic insertion mechanism to generate local curvature
by inserting an N-terminal amphipathic helix
into the lipid bilayer (Beck et al. 2008, Bielli
et al. 2005, Krauss et al. 2008, Lee et al. 2005,
PX and PH domains:
lipid binding domains
often specific for
certain
phosphoinositides
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Lundmark et al. 2008). In the next stage of vesicle budding, coat proteins serve as scaffolds. In
the case of vesicles budding from the ER, the
COPII coat is assembled in two steps. First, the
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Sheets
Nuclear
envelope
Tubules
b
bowtie-shaped Sec23/24 complex forms the inner shell of the coat by binding to the membrane
through its concave surface (Bi et al. 2002).
In the next step, the Sec13/31 heterodimers
bind, generating the cage-like outer shell of the
COPII coat (Fath et al. 2007, Stagg et al. 2008).
In the case of vesicles budding from the Golgi,
Arf1 recruits the coatamer complex in a single
step to form the COPI coat (Donaldson et al.
1992, Zhao et al. 1997).
As will be discussed below, several of the
paradigms of curvature generation during vesicle budding apply to organelle morphogenesis.
However, some differences should be pointed
out. First, a vesicle is not a permanent structure; it generally fuses quickly with another
membrane, dissipating its high membrane curvature. By contrast, cellular organelles, though
dynamic, maintain their curvature over long
time periods. Second, all curvature-generating
proteins involved in vesicle formation are soluble proteins that associate only transiently with
the lipid bilayer, whereas at least some proteins
that determine organelle curvature are integral
membrane proteins. Finally, the budding of a
small vesicle only requires the generation of
local curvature, whereas organelles often need
mechanisms that shape large membrane areas.
GENERATION OF THE TUBULAR
ER NETWORK
Figure 4
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Morphology of the endoplasmic reticulum (ER). (a) The different subdomains
of the ER, the nuclear envelope, the peripheral sheets, and the tubular network,
are highlighted in a COS-7 cell expressing a general ER marker fused to green
fluorescent protein (GFP). (b) Tubule-forming proteins localize to the tubular
ER, as seen for a GFP fused version of Rtn4a (green) in a BSC1 cell. By
I E
contrast,
W the luminal ER protein calreticulin (red ) is also found in the nuclear
envelope and peripheral ER sheets.
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The ER is composed of sheet-like structures
and a polygonal array of tubules connected
by three-way junctions (Baumann & Walz
2001, Shibata et al. 2006, Voeltz et al. 2002)
(Figure 4a). ER sheets are relatively flat areas, often extending over several micrometers
with little membrane curvature. ER sheets include the nuclear envelope whose curvature is
negligible owing to the large size of the nucleus it surrounds. In addition, in higher eukaryotes ER sheets are found in the peripheral
ER (the ER outside the nuclear envelope), usually close to the nucleus. In contrast, ER tubules
are cylindrical units with high membrane curvature in cross-section. In Saccharomyces cerevisiae,
ER tubules have a diameter of about 30 nm and
are located close to the cell cortex. They are
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connected by a few tubules with the nuclear envelope. In higher eukaryotes, ER tubules have
a diameter of about 60 nm, and the peripheral
ER extends throughout the entire cytoplasm.
The whole ER is a continuous membrane system with a common luminal space. How the
morphologically distinct domains are set up is
still poorly understood, but significant progress
has been made in elucidating the mechanism by
which the tubular ER network is generated and
maintained.
In all eukaryotes the ER network is extremely dynamic, with tubules undergoing continuous fusion and fission reactions (Lee &
Chen 1988, Prinz et al. 2000). In mammalian
cells, ER tubules are often close to microtubules
and can be generated by the force exerted by either microtubule-associated molecular motors
or by polymerizing microtubule tips (Terasaki
et al. 1986, Waterman-Storer & Salmon 1998).
The proteins that mediate the interaction with
motors or microtubules have not yet been identified. In S. cerevisiae, ER tubules are generated along actin filaments, and some evidence
suggests that the myosin Myo4p is involved
(Estrada et al. 2003). Although the cytoskeleton is required for the generation and distribution of the ER network, ultimately the alignment of ER tubules with the cytoskeleton is not
perfect. In addition, the ER network does not
collapse upon depolymerization of actin filaments in yeast (Prinz et al. 2000), and it retracts
only with some delay upon depolymerization
of microtubules in mammalian cells (Terasaki
et al. 1986). Thus, mechanisms other then those
based on the cytoskeleton stabilize and maintain the tubules. In vitro, an ER network can
be generated by the fusion of small vesicles in
the absence of an intact cytoskeleton (Dreier
& Rapoport 2000), suggesting that these additional mechanisms are sufficient to generate
and stabilize ER tubules.
Proteins involved in ER tubule formation
were discovered by the use of an in vitro system, in which the formation of an ER network could be recapitulated, starting with small
ER-enriched vesicles from Xenopus laevis eggs
(Dreier & Rapoport 2000, Voeltz et al. 2006).
Network formation is inhibited by reagents that
modify cysteine residues in membrane proteins.
The target of these reagents was identified as
reticulon 4a (Rtn4a) and confirmed by the inhibitory effect of Rtn4a antibodies on network
formation (Voeltz et al. 2006). Rtn4a belongs
to the reticulon family, whose members exist
in most, if not all, eukaryotic cells (Oertle &
Schwab 2003). There are four reticulon genes
in mammals, each with different splice forms,
and two reticulon genes in S. cerevisiae (Rtn1p
and Rtn2p). The conserved region is the Cterminal reticulon homology domain of approximately 200 amino acids, which contains
two long hydrophobic segments that each sit in
the membrane as a hairpin and expose no significant part of themselves on the luminal side
of the membrane.
The deletion of RTN1 and RTN2 in S. cerevisiae does not result in ER morphology defects,
unless the cells are placed in a high osmolarity medium, indicating that additional proteins
are involved in ER tubule formation (Voeltz
et al. 2006). A candidate protein, DP1 (also
called REEP5), was found by pull-down experiments with Xenopus Rtn4a. Similarly, the yeast
homolog of DP1, Yop1p, was found to interact
with both Rtn1p and Rtn2p. The DP1/Yop1p
protein family is again ubiquitous and includes
six mammalian DP1 and REEP proteins. The
DP1/Yop1p proteins do not share any primary sequence homology with the reticulons,
but they contain a conserved domain of about
200 amino acids, which includes two long hydrophobic segments that each form a hairpin in
the membrane.
The deletion of RTN1, RTN2, and YOP1
in S. cerevisiae leads to a drastic loss of tubular
ER (Voeltz et al. 2006). Most of the peripheral ER converts into sheets, although some
tubules are still present. The depletion of three
reticulons in mammalian cells has similar effects
(Anderson & Hetzer 2008). These data suggest that the reticulons and DP1/Yop1p are major components that shape the tubular ER. Interestingly, the triple-deletion mutant in yeast
exhibits only a moderate growth defect, indicating that in yeast much of the tubular ER
Coatamer: a complex
of proteins that serves
as the building block
for COPI coated
vesicles
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is dispensable (Voeltz et al. 2006). However,
the simultaneous depletion of Rtn1 and Yop1
in Caenorhabditis elegans causes a 60% decrease
in embryonic viability (Audhya et al. 2007), suggesting that an intact tubular ER is important
in higher eukaryotes.
Consistent with their proposed role in
ER tubule formation, the reticulons and
DP1/Yop1p localize almost exclusively to the
tubular ER; they avoid the low-curvature areas of the nuclear envelope and the sheets of
the peripheral ER even when highly overexpressed (Voeltz et al. 2006) (Figure 4b). In
mammalian cells, the overexpression of some
isoforms of the reticulons leads to long, unbranched tubules and to the disappearance of
peripheral ER sheets. In addition, the overexpression of the reticulons or DP1 makes the ER
network resistant to the collapse that normally
follows the depolymerization of microtubules,
indicating that these proteins can stabilize ER
tubules (Shibata et al. 2008). In yeast, the overexpression of Rtn1p or Yop1p also leads to long
tubules (Voeltz et al. 2006).
The reticulons and DP1/Yop1p are not only
necessary, but also sufficient for tubule formation (Hu et al. 2008). Purified yeast Yop1p or
Rtn1p, reconstituted with pure lipids into proteoliposomes, deform the lipid bilayer into narrow tubules. The diameter of these structures
(15–17 nm) is smaller than that of normal ER
tubules, owing to a higher concentration of the
tubule-inducing proteins. When the reticulons
or DP1/Yop1p are overexpressed in vivo, the
ER tubules also become narrower, and luminal
proteins are displaced.
The reticulons and DP1/Yop1p might shape
ER tubules by utilizing two cooperating mechanisms, hydrophobic insertion (wedging) and
scaffolding (Hu et al. 2008, Shibata et al. 2008)
(Figure 5). The proposed hydrophobic insertion mechanism is based on the presence of the
two hydrophobic hairpins in the tubule-shaping
proteins. Although the structure of the hairpins
is unknown, they might displace the lipids preferentially in the upper part of the outer monolayer, causing local curvature in an analogous
way to the insertion of amphipathic helices. It is
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a
Hairpin structure of
reticulons and DP1/Yop1p
in membrane
b
Figure 5
Wedging and scaffolding by the reticulons and DP1/
Yop1p. (a) The conserved region of the reticulons
and DP1/Yop1p contains two long hydrophobic
segments that each form a hairpin in the membrane.
The double-hairpin structure might adopt a wedge
shape that curves the membrane. (b) The reticulons
and DP1/Yop1p also form homo- and heterooligomers that could serve as arc-like scaffolds
(orange) around a membrane tubule ( gray).
also conceivable that they span the outer monolayer and shallowly insert into the inner monolayer, close to the membrane mid surface, which
would bend the bilayer in a similar way. Finally,
they could simply form a cone shape across the
entire bilayer and bend the membrane by the
bilayer-coupling mechanism (Figure 5a), although this alone would likely be insufficient
to generate enough curvature.
A scaffolding mechanism for the reticulons
and DP1/Yop1p is suggested by the observation
that these proteins form homo- and heterooligomers and are relatively immobile in the
plane of the membrane (Shibata et al. 2008).
Mutants of yeast Rtn1p, which no longer localize exclusively to the tubular ER or are inactive
17:48
in inducing ER tubules, oligomerize less extensively and are more mobile. The conserved domains of the reticulons and DP1/Yop1p, which
contain the two hairpin-shaped membrane segments, are responsible for the localization of the
proteins and their oligomerization. One possibility is that the oligomers form arc-like scaffolds that would mold the bilayer into tubules
(Figure 5a). Calculations show that these arcs
or rings around the tubules would need to occupy only about 10% of the total membrane
surface to generate near perfect tubules (Hu
et al. 2008). Rtn1p and Yop1p are among the
most abundant membrane proteins in yeast and
could together occupy 10% of the total surface
of the peripheral ER. Assuming that the reticulons and DP1/Yop1p form arcs rather than rings
around the tubules could explain why other ER
membrane proteins can diffuse in the plane of
the membrane over long distances.
The oligomers formed by the reticulons and
DP1/Yop1p must be of relatively small size and
undergo continuous assembly and disassembly,
because otherwise all molecules would eventually localize to one site of the ER and no
molecules would be available for the rest of the
ER. The disassembly of the oligomers may indeed be an active process because the depletion
of ATP in yeast or mammalian cells decreases
the diffusional mobilities of the reticulons and
DP1/Yop1p in the plane of the membrane
(Shibata et al. 2008). The mechanism by which
the oligomers are disassembled remains to be
clarified.
The reticulons and DP1/Yop1p are not the
only proteins required for the formation of
the tubular ER network. Recent work indicates
that a class of membrane-bound, dynamin-like
proteins, which includes the atlastins in mammals and Sey1p in yeast, play an important
role (Hu et al. 2009). Although the atlastins
and Sey1p do not share sequence homology,
they belong to the same family of dynamin-like
GTPases and have the same membrane topology; that is, they possess at their C-termini two
hydrophobic membrane anchors with only 3–
4 amino acids between them on the luminal
side of the membrane. These GTPases localize
predominantly to the tubular ER and interact with the reticulons and DP1/Yop1p. Depletion of the atlastins or expression of dominantnegative versions leads to long ER tubules with
little branching. Antibodies to the atlastins inhibit ER network formation in vitro, and yeast
cells lacking Sey1p and either Rtn1p or Yop1p
have strong defects in the morphology of the
tubular ER. Together, these data indicate that
the atlastins and Sey1p function in ER network
formation, likely by affecting the generation of
tubule branches.
A possible mechanism by which the atlastins
and Sey1p function in ER network formation is
based on the analogy to dynamin-1 (Hu et al.
2009). Assuming a similar mechanism for the
atlastins and Sey1p, these proteins could be involved in the fission of ER tubules. This process
would be facilitated if the ER tubules were under tension by the cytoskeleton, either by direct
association or through molecular motors. Alternatively, the atlastins and Sey1p could function in fusion. In fact, they are actually more
similar to the dynamin-like mitofusion/Fzo1p
proteins, which are anchored in the outer mitochondrial membrane by two closely spaced hydrophobic segments and are implicated in the
fusion of the tubular mitochondria (see below).
Given that Sey1p is much less abundant than
Rtn1p or Yop1p in yeast, it seems likely that
the reticulons and DP1/Yop1p are the main
structural components that shape the tubules
of the ER, whereas the atlastins and Sey1p associate with them at discrete points along the
tubules. During GTP hydrolysis, the GTPases
would destabilize the lipid bilayer, which in
turn would lead to fission or fusion of the ER
tubules.
Interestingly, mutations in one atlastin isoform cause the most frequent early-onset
hereditary spastic paraplegia (HSP), a disease
affecting the long axons of corticospinal motor neurons (Salinas et al. 2008). Given the
role of the atlastins in ER network formation,
the disease may be caused by ER morphology
defects (Hu et al. 2009). It is plausible that
mutants of atlastin would affect most severely
the longest axons, in which the ER network
Hereditary spastic
paraplegia: a group of
neuronal disorders
marked by progressive
spasticity and weakness
of the lower limbs
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needs to extend a long distance from the cell
body to the axon terminus. Two other proteins,
spastin and REEP1, mutant alleles of which
are known to frequently cause HSP, may also
affect ER network formation. A long isoform
of the microtubule-severing adenosine triphosphatase (ATPase) spastin is membrane-bound
and interacts with the atlastins and the reticulons (Evans et al. 2006, Mannan et al. 2006,
Sanderson et al. 2006); it could affect microtubule dynamics and thus ER tubule formation.
REEP1, a member of the DP1/Yop1p family of
tubule-forming proteins could affect the shape
of ER tubules directly. These three proteins collectively account for well over 50% of all HSP
cases, suggesting that ER morphology defects
may be a predominant mechanism underlying
these neuropathogenic disorders.
In summary, the generation of the tubular
ER network appears to involve several principles: (a) the cytoskeleton-based pulling of
tubules, (b) the shaping of tubules by the reticulons and DP1/Yop1p, which might utilize both
hydrophobic insertion (wedging) and scaffolding mechanisms, and (c) the formation of an
interconnected network, which involves the atlastin and Sey1p GTPases. These principles are
still rather speculative, but they are consistent
with how membranes are shaped and remodeled in other systems. Clearly, the mechanism
of ER network formation requires further studies, particularly in systems composed of purified components. Structural studies are needed,
but will be challenging, given that most of the
players are integral membrane proteins that
form heterogeneous oligomers. The regulation
of the components is another unresolved issue.
For example, there are major changes of ER
morphology during mitosis in higher eukaryotes, and it is therefore possible that the components involved in ER network formation are
modified, perhaps by phosphorylation. Finally,
it is possible that at least some of the components involved in network formation have other
functions as well. For example, some of the
reticulon isoforms have long segments in the
cytosol, which presumably interact with other
proteins. It will be interesting to see whether
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these proteins link the tubule-forming proteins
to other cellular processes.
GENERATION OF MEMBRANE
SHEETS IN THE ER
Peripheral ER sheets have little overall curvature along the plane of the two membranes.
The abundance of sheets varies among different
cell types, but is particularly prominent in cells
that are highly specialized in secretion. Here,
the sheets are generally stacked on top of each
other, and both the cytosolic and luminal spacings are constant (∼50–100 nm). Because the
luminal width of sheets is about the same as
the diameter of ER tubules, the edges of the
sheets must have a similarly high curvature as
the tubules.
The mechanisms used to form peripheral
ER sheets are largely unknown. However, one
possibility is that the same proteins that stabilize ER tubules, the reticulons and DP1/Yop1p,
stabilize the high curvature of the sheet edges
by forming arc-like scaffolds along the curved
membrane (Figure 6a). Scaffolding the edges
could generate even large areas of sheets, although the width between the membranes
would likely fluctuate. This mechanism may explain why the observed luminal width of ER
sheets is similar to the diameter of ER tubules,
and why the reticulons are upregulated when
ER sheets proliferate (Benyamini et al. 2009).
Indeed, endogenous and overexpressed forms
of the reticulons, as well as the atlastins, localize to the edges of ER sheets (Hu et al 2009
and Y. Shibata, unpublished data). The sheetedge localization would also allow for tubules
to readily emerge from or fuse into the edges
of peripheral ER sheets.
An alternative or additional mechanism by
which peripheral ER sheets could be formed
is intraluminal bridges (Figure 6b). Such
bridges have been observed by deep-etch electron microscopy and could prevent excessive fluctuations of the luminal width (Senda
& Yoshinaga-Hirabayashi 1998). A candidate
bridging protein may be CLIMP-63, an abundant integral membrane protein with a large
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b
c
Figure 6
Three possible mechanisms to generate the sheets of
the peripheral endoplasmic reticulum (ER). The
cross-section of an ER membrane sheet is shown in
gray. (a) The tubule-shaping proteins (the reticulons
and DP1/Yop1p) could form arc-like scaffolds
(orange) around the edges of the sheet and stabilize
their high curvature. (b) Transmembrane proteins
(blue) that are localized in both membranes could
interact through their luminal domains to keep the
two membranes flat and at a fixed distance from one
another. (c) Transmembrane proteins ( yellow) could
form a flat scaffold on the cytosolic side of the ER
membrane.
coiled-coil luminal domain that can form 90nm rod-like oligomers (Klopfenstein et al.
2001). At endogenous levels, this protein localizes to peripheral sheets and, when overexpressed, it induces the proliferation of sheets
with a constant luminal width (Y. Shibata, unpublished data). However, depletion of this protein by RNAi does not noticeably affect the
abundance of peripheral sheets (unpublished
data), suggesting that other functionally equivalent proteins may exist.
Peripheral ER sheets could also be generated by proteins that form a flat scaffold
on the cytoplasmic side of the ER membrane
(Figure 6c). A candidate scaffolding protein is
p180, an abundant integral membrane protein
with a large cytosolic coiled-coil domain. At endogenous levels, this protein also localizes predominantly to ER sheets (Y. Shibata, unpublished data). Overexpression of p180 in yeast or
in tissue culture cells leads to the proliferation
of ER sheets, whereas depletion of this protein
in a monocyte cell line prevents ER proliferation during differentiation (Becker et al. 1999,
Benyamini et al. 2009). p180 may also be involved in stacking sheets by bridging the cytosolic surfaces of ER membranes. The functions proposed for p180 are analogous to those
of the golgins, which are large coiled-coil proteins that are thought to flatten and link the
stacks of the Golgi apparatus (Short et al. 2005).
Because CLIMP-63 and p180 are found only in
vertebrates, it is possible that additional players
are involved in sheet formation.
Interestingly, several observations suggest a
tug-of-war between peripheral ER sheets and
tubules. The overexpression of Rtn4a in mammalian cells leads to the disappearance of ER
sheets (Voeltz et al. 2006), whereas the overexpression of CLIMP-63 reduces the abundance
of ER tubules (Y. Shibata, unpublished data).
The reduction or absence of the tubule-forming
proteins in mammalian or yeast cells converts
the tubules into sheets (Anderson & Hetzer
2008, Voeltz et al. 2006). These data indicate
that the relative abundance of ER sheets and
tubules is dictated by the expression levels of
sheet- and tubule-shaping proteins.
The other sheet-like ER domain is the nuclear envelope. The sheets formed by the inner
and outer nuclear membranes are again separated by a distance of ∼50 nm, which means
that at the sites where they are connected, at the
nuclear pores, the membrane has a high curvature (Antonin & Mattaj 2005). The nuclear
envelope might be shaped in large part by a
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luminal bridging mechanism involving the
SUN and nesprin proteins (Worman &
Gundersen 2006). The SUN proteins are integral membrane proteins that localize to the
inner nuclear membrane through their interaction with the nuclear lamins, whereas the nesprins span the outer nuclear membrane. These
proteins interact with each other in the nuclear
envelope lumen through nesprin’s conserved
KASH domain (Padmakumar et al. 2005). Depletion of either the SUNs or the nesprins results in a nuclear envelope with large bulges
(Crisp et al. 2006). Homologs of both the SUNs
and nesprins also exist in yeast and have been
shown to affect nuclear envelope morphology
(King et al. 2008). However, these proteins appear to be involved in other processes as well,
such as nuclear positioning and spindle pole
body duplication.
The high curvature of the nuclear membranes around the nuclear pores is likely stabilized by a nuclear pore protein subcomplex
(Nup84 complex). This complex contains the
Sec13 protein, which is also a component of the
COPII coat required for vesicle budding from
the ER. In addition, it contains a nuclear porespecific Sec13 homolog, Seh1, as well as five
other proteins with structural motifs that are
also found in vesicle coats (Devos et al. 2004).
In one model, these proteins form rods along
the curved nuclear pore membrane vertical to
the plane of the envelope (Debler et al. 2008,
Hsia et al. 2007). Alternatively, they could form
a membrane-associated lattice, similar to that
seen in the COPII coats (Brohawn et al. 2008).
In either case, they would serve as a scaffold
that stabilizes the high curvature. Additionally,
several nuclear pore proteins contain amphipathic sequence motifs, which at least in the
case of a component of the Nup84 complex
can insert into the lipid bilayer and stabilize local curvature (Drin et al. 2007). The nuclear
pores could contribute to maintain the flat nuclear envelope, similar to how the reticulons
and DP1/Yop1p might generate peripheral ER
sheets. However, the number of nuclear pores
is variable in different cell types (Hetzer et al.
2005), and nuclear envelopes without pores can
IM: inner
mitochondrial
membrane
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be formed in Xenopus egg extracts (Goldberg
et al. 1997, Hetzer et al. 2005). Recent results suggest that reticulons and Yop1p are involved in early stages of nuclear pore assembly
(Dawson et al. 2009), perhaps by transiently stabilizing high curvature at the site of pore insertion. However, these proteins must eventually
leave the nuclear envelope and are not essential
for nuclear pore insertion.
SHAPING MITOCHONDRIA
Several aspects discussed for the morphogenesis
of the ER apply to mitochondria as well. However, mitochondria are more complex than the
ER in that they have two membranes, the outer
mitochondrial membrane (OM) and the inner mitochondrial membrane (IM) (Figure 7a).
The IM separates the innermost compartment,
the matrix, from the intermembrane space; it
folds and undulates to form deep invaginations,
called cristae. Although mitochondrial shape
can vary greatly among different species and
cell types and change under varying physiological conditions, these architectural features are
conserved in all eukaryotes.
Like the ER, mitochondria form an interconnected tubular network (Figure 7b) that
continuously fuses and divides, and they use
the cytoskeleton to move and distribute within
the cell. In budding yeast and plants, this
movement occurs mostly along actin filaments
via myosin motors (Boldogh & Pon 2007,
Frederick & Shaw 2007). In other eukaryotes,
including fission yeast, mitochondria move on
microtubules using kinesin or dynein motors.
Depolymerization of actin or microtubules
inhibits mitochondrial movement. While prolonged perturbation leads to clustered and/or
coalesced mitochondria, acute cytoskeletal
depolymerization has little effect (for review,
see Griparic & van der Bliek 2001), indicating
that mitochondrial morphology may not be as
highly dependent on the cytoskeleton as the
ER. In higher eukaryotes, the interaction between mitochondria and the kinesin motor may
be mediated by the large GTPase Miro, a protein that is anchored to the OM via a C-terminal
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a
Outer
membrane
Inner
membrane
b
c
Outer
membrane
Cristae
Inner
membrane
Figure 7
The morphology of mitochondria. (a) Thin-section electron micrograph of a mitochondrion from a bat
pancreatic cell shows the different membrane structures of the outer mitochondrial membrane (OM), inner
nuclear membrane (IM), and cristae. (Adapted from Fawcett 1981.) (b) Mitochondria form an interconnected
tubular network, as visualized by Mitotracker staining in a C2C12 cell (Griparic & van der Bliek 2001).
(c) A reconstructed three-dimensional view, using electron microscopic tomography, of the cristae of a
mitochondrion isolated from rat liver (Mannella 2000). Arrows denote the tubular cristae junctions.
transmembrane segment, as well as the cytosolic Milton protein (Glater et al. 2006, Guo
et al. 2005, Wang & Schwarz 2009). In budding
yeast, the Miro-related protein Gem1p also
plays a role in mitochondrial morphology
(Frederick et al. 2004), but its precise function
is unclear because mitochondrial movement is
actin-based; Gem1p does not associate with the
myosin Myo2p, and functions independently of
known myosin adaptors (Frederick et al. 2008).
The overall morphology of the mitochondrial network depends on balanced rates of fission and fusion. Perturbing the fusion machinery leads to more fragmented mitochondria,
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whereas defects in the fission apparatus lead to
a netlike, interconnected morphology (for review, see Detmer & Chan 2007, Hoppins et al.
2007, Okamoto & Shaw 2005). Interestingly,
perturbing both processes restores normal morphology in some cells. Both fission and fusion
events involve dynamin-like proteins.
For mitochondrial fission, the soluble
dynamin-related protein Drp1/Dnm1p is first
recruited to certain sites of the mitochondrial
OM by the soluble adaptor Cav4/Mdv1p and
the membrane protein Fis1. In vitro experiments and the analogy to classical dynamin-1
suggest that these proteins form oligomeric
spirals around the mitochondria (Ingerman
et al. 2005); fission of both the OM and IM
would then occur through some nucleotidedependent conformational change of these
oligomers. Fusion of the OM is mediated by the
dynamin-like protein mitofusin/Fzo1p (fuzzy
onion). The protein is anchored in the OM
via two transmembrane segments that form
a hairpin within the membrane, a topology
that is similar to that of the atlastins and
Sey1p; the GTPase domain faces the cytosol.
Mitofusin/Fzo1p proteins interact in trans with
each other via a C-terminal coiled-coil domain,
which could tether together opposing OMs
(Koshiba et al. 2004). However, because the interacting proteins form an antiparallel coiledcoil, the association would be expected to keep
the membranes at some distance, rather than
force them into close apposition for fusion as
the zippering of SNARE helices does. It appears
more likely that mitofusin/Fzo1p, like other
dynamin-like GTPases, forms spirals around
membrane tubules; the highly curved free tips
of the tubules would be fusogenic (Praefcke &
McMahon 2004). A mitofusin-related, bacterial protein indeed forms spirals around lipid
tubes (Low & Lowe 2006). Interestingly, the
X-ray structure of this protein indicates that
in the nucleotide-free and GDP-bound states
the hydrophobic segments are buried (Low &
Lowe 2006). Although the bacterial dynamins’
function is unknown, they might offer a simplified system to study the mechanism of the
mitofusin/Fzo1p.
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Fusion of the IM is mediated by another
dynamin-like protein, OPA1/Mgm1p, which
associates with the IM and exposes its GTPase
domain to the intermembrane space. This
protein is proteolytically processed; in yeast,
both the shorter, soluble form and the longer,
membrane-bound form are necessary for IM
fusion (Herlan et al. 2003), whereas in mammals, only the membrane-bound version is required (Ishihara et al. 2006). Mechanistic details of how these proteins mediate fusion are
unclear, but perhaps they also create fusogenic
areas by promoting high curvature. In yeast, IM
and OM fusion is coupled by a physical link between Mgm1p and Fzo1p through the adaptor
protein Ugo1p (Sesaki & Jensen 2004).
How the overall tubular structure of mitochondria is generated is not well understood.
Whereas mitochondrial tubules have a typical
diameter of 0.5–1 um and are therefore less
curved than ER tubules (30–60 nm), their overall cylindrical shape suggests that some active
mechanism is required to generate them. Are
dynamin-mediated fusion and fission and cytoskeletal tethering sufficient for shaping mitochondrial tubules? Most likely not, because
disruption of the fusion and fission machinery or acute depolymerization of the cytoskeleton does not discernibly affect the diameter of
the tubules. These results indicate that some
other structural components are involved in directly shaping mitochondrial tubules. In yeast, a
number of proteins have been identified whose
absence or mutation cause tubulation defects
(for review, see Okamoto & Shaw 2005); the
mitochondria are condensed into spherical, giant structures with very large diameter. Some
of these proteins—(Mmm1p, Mdm10p, and
Mdm12p) were initially proposed to mediate
mitochondria-actin attachments by recruiting
the Arp2/3 complex (Boldogh et al. 2003).
However, these proteins have also been implicated in beta-barrel protein insertion into
the OM (Meisinger et al. 2004, 2007), suggesting that some unknown beta-barrel protein may be involved in the tubulation of
mitochondria. The possibility also exists that
more classical membrane-bending proteins that
17:48
employ hydrophobic insertion and scaffolding
mechanisms are involved in shaping mitochondrial tubules. For example, endophilin B1, a
BAR domain-containing protein, is involved
in maintaining mitochondrial morphology in
mammalian cells (Karbowski et al. 2004), although it has been implicated in fission. In addition, still unidentified proteins may exist, similar to the reticulons/DP1/Yop1p involved in ER
tubule formation.
How the cristae are shaped is another interesting issue. These membrane structures originate at narrow tubular openings of the IM
called cristae junctions (Figure 7c). The cristae
themselves are tubular, sheet-like, or even crystalloid (Figure 7c). They can be interconnected by three-way junctions and can span the
entire width of a mitochondrion (for review,
see Frey & Mannella 2000, Mannella 2006).
Whereas cristae adapt their shapes depending
on varying physiological conditions, the shape
of cristae junction tubules remains constant (a
cross-sectional diameter of ∼28 nm), suggesting that a tight protein scaffold maintains this
structure.
At least two independent mechanisms
that involve wedging and/or scaffolding have
emerged that contribute to the formation of
the high curvature areas of cristae. The first
involves the ATP synthase machinery, which
is an abundant component of cristae membranes. Mutant yeast cells lacking the e and g
subunits of the ATP synthase have mitochondria with onion-like sheets of IM (Paumard
et al. 2002). These subunits are nonessential
for the enzymatic activity, but are required for
the dimerization and oligomerization of the
ATP synthase. Structural data indicate that the
ATP synthase dimer has a wedge shape, where
the dimer complex takes up more space on
the matrix side of the membrane (MinauroSanmiguel et al. 2005). Additionally, electron
microscopy shows that ATP synthase dimers
form 1-um oligomeric ribbons around the highcurvature tips of the cristae (Strauss et al. 2008).
Thus, like the reticulons and DP1/Yop1p, the
ATPase complex may cause membrane curvature by both wedging and scaffolding. Recent
results suggest the e and g subunits of the yeast
ATP synthase cooperate with Fcj1p, a homolog
of the mammalian mitofilin protein, to form
cristae junctions (Rabl et al. 2009). The overexpression of Fcj1p increases the number of
cristae junctions and its deletion results in the
loss of junctions. Fcj1p may stabilize the negative curvature at branch points of the IM and
cristae.
The second mechanism involves OPA1/
Mgm1p, the IM dynamin-like proteins implicated in fusion, and the prohibitins. Mutation or depletion of OPA1/Mgm1p or of
the prohibitins leads to more fragmented
mitochondria with aberrantly large, bulging
cristae, which sometimes appear disconnected
(Meeusen et al. 2006, Merkwirth et al. 2008).
Prohibitins themselves oligomerize into large,
ring-shaped scaffold-like structures (Tatsuta
et al. 2005), but seem to indirectly affect cristae
structure by influencing the proteolytic processing of OPA1 (Merkwirth et al. 2008); expression of the OPA1 long isoform containing
the transmembrane segment rescues the cristae
morphology defect in cells lacking prohibitin.
How exactly these proteins affect cristae morphology remains to be elucidated.
Lipid rafts:
cholesterol-enriched
microdomains in cell
membranes
GENERATING THE CURVATURE
OF CAVEOLAE
Caveolae can be considered to be distinct, relatively stable organelles. They are flask-like
invaginations located on the surface of many
vertebrate cells (Figure 8); the largest diameter of the bulb is 60–80 nm (for review, see
Parton et al. 2006, Parton & Simons 2007, Stan
2005). Although discovered more than half a
century ago, the function of these structures is
not entirely clear; they have been implicated
in lipid uptake and regulation, signal transduction, endocytosis, and virus entry. Caveolae are specialized forms of lipid rafts, enriched in cholesterol and sphingolipids (Simons
& Toomre 2000). The major structural component of caveolae is the integral membrane protein caveolin (Rothberg et al. 1992). In mammals there are three caveolin genes. Caveolin-1
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100 nm
b
Figure 8
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The morphology of caveaolae. (a) The flask-like invaginations of caveolae in an
endothelial cell are visualized by thin-section electron microscopy (Stan 2002).
(b) Caveolin forms filaments on the surface of caveola, as viewed from the cytoplasm
I E using rapid-freeze, deep-etch electron microscopy in a human fibroblast
W lower panels show individual caveolae. Image courtesy of J. Heuser.
cell. The
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is the main member of the family and is expressed in many cell types. Its deletion in mice
leads to the complete absence of caveolae (Drab
et al. 2001).
Caveolin may use both wedging and scaffolding to shape caveolae, similar to the mechanisms by which the reticulons and DP1/Yop1p
are thought to shape ER tubules. Caveolin is
extremely abundant, with about 145 molecules
per typical caveola (Pelkmans & Zerial 2005).
The protein contains an amphipathic helix (α2a
helix) that is inserted into the bilayer (Le Lan
et al. 2006, Parton et al. 2006). This helix
also binds cholesterol and could cause the raftlike lipid composition of caveolae (Epand et al.
2005). Calculations show that the curvature of
caveolae could be generated by the bilayer coupling mechanism if the amphipathic helix was
inserted into the outer leaflet of the bilayer and
each helix sequestered 13 cholesterol molecules
(Parton et al. 2006). In addition, curvature
might be generated by caveolin’s membrane anchor. The protein contains a hydrophobic segment of 33 residues that forms a hairpin in
the membrane, with no residue on the external side of the membrane (Glenney & Soppet
1992). This is supported by the fact that caveolin can move into lipid bodies (Fujimoto et al.
2001, Ostermeyer et al. 2001, Pol et al. 2001),
in which the protein must sit in a lipid monolayer that surrounds an entirely hydrophobic
core. The hairpin structure could cause membrane bending in a similar way as discussed for
the tubule-shaping ER proteins.
The scaffolding mechanism is supported
by electron microscopy studies, which indicate
that caveolae are coated by filaments of caveolin
(Rothberg et al. 1992) (Figure 8b). In sucrose
gradient centrifugation, detergent-solubilized
caveolin migrates at approximately 350 kDa
(Sargiacomo et al. 1995). Caveolin appears to
oligomerize into 10-nm particles (about 10–
15 molecules), which then interact with each
other to form filaments. The oligomerization
of caveolin requires several regions of the protein (Das et al. 1999, Schlegel & Lisanti 2000,
Song et al. 1997), but the exact mechanism of
assembly remains unclear.
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Whether caveolin is sufficient to form caveolae is unknown. Recent studies have identified a conserved cytoplasmic protein, cavin (also
known as PTRF), as a putative, additional coat
protein (Hill et al. 2008, Liu & Pilch 2008).
The cellular level of cavin correlates with that
of caveolin, cavin interacts with caveolin in mature caveolae, and the deletion of cavin results in the loss of caveolae (Hill et al. 2008,
Liu et al. 2008). What precisely cavin is doing
and how it cooperates with caveolin remains to
be elucidated. In some endothelial cell types a
specialized structure is seen at the neck of caveolae (stomatal diaphragm), whose major component is the PV1 protein (Stan 2005). Several
molecules of this single-spanning membrane
protein may interact through their extracellular
domains to bridge the neck.
CONCLUSIONS
The regulation of membrane curvature is
now recognized as a major factor that determines the shape of membrane-bound organelles. It appears that high-curvature membranes are mainly formed by proteins that
utilize hydrophobic insertion (wedging) and
scaffolding mechanisms, similar to how curvature is generated in budding vesicles. These
curvature-generating mechanisms cooperate
with cytoskeleton-based force generation. In
addition, membrane remodeling by fusion and
fission often involves GTPases of the dynamin
family. These principles of membrane shaping and remodeling appear to be quite general, although we have focused only on a few
organelles. However, in all cases the mechanistic details are still unclear. It is also generally
unknown how flat membrane sheets are generated. Although this does not necessarily require
active mechanisms, it is likely that proteins
help keep lipid bilayers equidistant and stacked
on top of each other. Finally, a major question is how membrane morphology is linked to
function. For example, are ER sheets and ER
tubules functionally distinct? It has been proposed that ER sheets correspond to rough ER,
where membrane-bound polysomes synthesize
secretory and membrane proteins; ER tubules
may correspond to a protected membrane area
where the high membrane curvature precludes
the binding of large polysomes, allowing other
proteins to gain access to the membrane surface
to carry out processes such as lipid synthesis
and Ca2+ signaling (Shibata et al. 2006). Again,
these proposals are speculative and require experimental testing. Although our discussion of
the shaping of organelles demonstrates how
little we know, the recent advances in the
field give confidence that rapid progress can
be made on this fundamental problem in cell
biology.
Lipid bodies:
Particles consisting of
a lipid monolayer
surrounding a
hydrophobic interior
made of cholesterol
esters and triglycerides
SUMMARY POINTS
1. Membrane-bound organelles have characteristic shapes that require mechanisms that
regulate membrane curvature.
2. High membrane curvature in organelles is generated by two major mechanisms that are
similar to those involved in forming small intracellular vesicles: hydrophobic insertion
(wedging) and scaffolding. In addition, cytoskeleton-based force generation plays an
important role in shaping organelles.
3. Fusion and fission is a major mechanism of membrane remodeling. It appears to be
generally catalyzed by proteins of the dynamin family of GTPases.
4. The tubules of the ER are shaped by the reticulons and DP1/Yop1, which might
use hydrophobic insertion (wedging) and scaffolding mechanisms for membrane
deformation. The dynamin-like proteins atlastin and Sey1p appear to be required for
network formation.
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5. Similar principles of membrane shaping and remodeling may apply to the morphogenesis
of mitochondria and caveolae.
FUTURE ISSUES
1. What are the actual structures of the reticulons and DP1/Yop1p? How deeply do the
hydrophobic segments insert into the lipid bilayer? Do they form arc-like scaffolds around
tubules?
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2. Are the reticulons and DP1/Yop1p regulated? Do they interact with other proteins that
link them to additional cellular functions?
3. Are the atlastins and Sey1p directly involved in fusion or fission of ER tubules?
4. How are ER tubules associated with molecular motors or the tips of growing microtubule
or actin filaments?
5. How are the sheets of the peripheral ER and the nuclear envelope generated and
maintained?
6. What is the advantage for a cell to have different ER morphologies? Do sheets correspond
to sites where membrane-bound polysomes are located, and how are these membrane
areas kept distinct?
7. How are the tubules of mitochondria generated? How are the cristae shaped? How do
the dynamin-like proteins cause fusion and fission, and how is their activity coupled to
cause coordinated changes of the inner and outer membranes?
8. How exactly do caveolin and other proteins cause the formation of flask-like membrane
structures? Can the formation of caveolae be reproduced with pure proteins and lipids?
DISCLOSURE STATEMENT
The authors are not aware of any biases that might be perceived as affecting the objectivity of this
review.
ACKNOWLEDGMENTS
We thank Janet Iwasa for help with the figures and Adrian Salic for critical reading of the
manuscript. Y.S. is supported by the American Heart Association. T.A.R. is a Howard Hughes
Medical Institute Investigator.
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