Content uploaded by William J. Tyler
Author content
All content in this area was uploaded by William J. Tyler on Feb 09, 2015
Content may be subject to copyright.
Cells are continuously subjected to
mechanical forces that influence cell division,
gene expression, cell migration, morpho-
genesis, cell adhesion, fluid homeostasis, ion
channel gating and vesicular transport1–3.
The rapidly growing field of mechanobiology
is focused on investigating the influence of
mechanical forces on cellular and molecular
processes. The primary motivations of mech-
anobiological investigations are to uncover
the mechanisms that enable cells to sense,
transduce and respond to mechanical stimuli,
as well as to characterize the mechanical
properties of molecules andcells2.
Numerous mechanical events are known
to occur in neurons. In axons, for instance,
action potentials are accompanied by propa-
gating membrane deformations (volumetric
changes)4,5. Mechanical impulses have also
been recorded at axon terminals during
action potential firing and vesicle fusion6.
Dendritic spines ‘twitch’7 and experience rapid
actin-mediated contractions in response to
synaptic activity8. The extent to which these
cellular-mechanical dynamics influence brain
function, however, remains a mystery. This
gap in our knowledge probably exists because
neuroscientists have not traditionally consid-
ered the roles of classical mechanics in brain
function. The intent of this article is therefore
to challenge our current models of neuronal
physiology and plasticity, which do not at pre-
sent account for the cellular mechanics that
affect neurons. I provide a brief overview of
the principles by which phospholipid mem-
branes, the cytoskeleton, extracellular matrix
(ECM) proteins, cell adhesion molecules and
membrane proteins endow the brain with
mechanical properties (FIG.1). I also discuss
hypotheses pertaining to how interactions
among some of the mechanical features
of the brain underlie various aspects of
synaptic signalling, neuronal plasticity and
traumatic injury. In addition, I describe
mechanobiological methods and tools that
are readily applicable to modern neurobiol-
ogy to further encourage the investigation of
cellular mechanics in neuroscience. Overall,
the literature reviewed in this article indi-
cates a need to expand our consideration
of the forces that underlie the mechanical
(physical) plasticity of the brain and their
consequences for neuronal signalling.
Mechanical properties of the brain
The Young’s modulus or elastic modulus (E)
of a material describes its resistance (or ten-
dency) to deform in response to mechanical
stress. Data from studies characterizing the
macroscopic physical characteristics of the
brain indicate that it is a viscoelastic material
and is one of the softest tissues in the body9–12.
Many values describing the elastic properties
of the brain have been estimated by convert-
ing the shear modulus (G) of the brain to E,
where the brain is assumed to have a Poisson’s
ratio (ν) of 0.5 and E = 2G(1 + ν). Rodent and
human brains have been described as having
an estimated E ranging from 0.1 to 16 kPa
(1 nN/μm2 = 1,000 Pa)9–12. Other tissues in the
body have higher elastic moduli and are more
rigid than brain tissue. For example, bone,
which is recognizably stiff, has an E of about
15–30 GPa, whereas less rigid connective
tissues and arteries have E ≈ 0.1–1 MPa and
even softer muscle has E ≈ 10–100kPa13.
Elastography uses magnetic resonance or
ultrasound approaches to estimate the stiff-
ness of tissues by imaging their responses
to sound (shear) waves propagated through
thebody. Magnetic resonance elastography
(MRE) is useful for characterizing and map-
ping the nonlinear viscoelastic properties of
the intact human brain11,14. MRE has shown
that the stiffness of brain regions varies sub-
stantially in normal humans11,14,15 and that
these mechanical properties change with
age16 and disease state17,18. Understanding the
molecular and cellular properties of neurons
that underlie these mechanical changes and
how they give rise to functional outcomes
should serve as focal points for mechanobio-
logical studies of the brain. Owing to the inte-
grated structural and mechanical properties of
cells and tissues, forces affecting one of their
components can in turn produce tension and
strain in others3. Below, I discuss the plasma
membrane, cytoskeleton, ECM, cell adhesion
proteins and ion channels with respect to how
they generate or are functionally affected by
mechanical forces in neurons (FIG.1a).
The plasma membrane. The phospholipid
bilayers of plasma membranes give rise to
many of the viscoelastic properties of the
brain. Plasma membranes are dynamic and
experience structural changes across broad
time and length scales ranging from nano-
scopic (nanosecond and nanometre) to
microscopic and mesoscopic (microsecond
and millimetre)19,20. Phospholipids have lat-
eral diffusion constants in bilayers on the
order of 10−12 m2/s21, theyundergo
trans-gauche isomerization every 10–20
nanoseconds, and rotate or wobble on
nanosecond timescales20. Plasma mem-
branes respond to force in a time-varying
manner as nonlinear functions of strain,
meaning that they are viscoelastic or
non-Newtonian fluids19. Here, phospho-
lipid bilayers reflect a Maxwell material
that shows frequency-dependent changes
in tension and viscosity with viscoelastic
relaxation times on the order of tens of
microseconds19.
The deformation of plasma membranes
in response to a force can be described by
their compression (KC), area expansion (KA)
and bending (KB) moduli (FIG.1b). In each
case, a larger modulus indicates greater
resistance to a deformation force, whereas a
smaller modulus indicates lower resistance.
Membrane deformations can affect the activ-
ity of ion channels on millisecond timescales
relevant to neuronal activity (FIG.2; see ‘Ion
channels’ and BOX1). The basic viscoelastic
properties of plasma membranes can be
OPINION
The mechanobiology of
brain function
William J.Tyler
Abstract | All cells are influenced by mechanical forces. In the brain,
force-generating and load-bearing proteins twist, turn, ratchet, flex,
compress, expand and bend to mediate neuronal signalling and plasticity.
Although the functions of mechanosensitive proteins have been thoroughly
described in classical sensory systems, the effects of endogenous mechanical
energy on cellular function in the brain have received less attention, and many
working models in neuroscience do not currently integrate principles of
cellular mechanics. An understanding of cellular-mechanical concepts is
essential to allow the integration of mechanobiology into ongoing studies of
brain structure and function.
PERSPECTIVES
NATURE REVIEWS
|
NEUROSCIENCE VOLUME 13
|
DECEMBER 2012
|
867
© 2012 Macmillan Publishers Limited. All rights reserved
Nucleus
Nature Reviews | Neuroscience
Plasma membrane
Cadherin
Extracellular matrix
Integrin
Lipid ra
Osmolarity
Microtubule
Actin filament
Neurofilament
Motor
protein
Spectrin
Channel
a
bAdjacent cell
KBKC
Fadhesion
Finteract KA
Fstall
Actin filament
Fmatch
Protein inclusion
Cargo
Fpush
Cellular anchor
Neurofilament
Fpull
Microtubule
Fbuckle
Motor protein
Adhesion molecules
Spectrin
Figure 1 | Mechanical forces are generated and transduced in neurons.
a | The cell body of a neuron, showing cellular and molecular components
that transduce or sense micromechanical forces. The components illustrated,
such as the plasma membrane, ion channels, actin filaments, microtubules,
neurofilaments, motor proteins, spectrins, integrins, extracellular matrix,
lipid rafts and osmolarity, have key roles in neuronal and glial function.
b | A neuron or glial cell showing several properties of plasma membranes,
including the bending (KB), compression (KC) and area expansion (KA) moduli.
Stalling (Fstall) and buckling (Fbuckle) forces act on actin filaments, and microtu-
bules and neurofilaments interact and exert pushing (Fpush) and pulling (Fpull)
forces. Adhesion forces (Fadhesion) generated by cell adhesion molecules, such
as integrins and cadherins, couple cells together. In addition, interaction
(Finteract) and hydrophobic matching (Fmatch) forces are generated by the
inclusion of a protein, such as an ion channel, in the plasma membrane.
PERSPECTIVES
868
|
DECEMBER 2012
|
VOLUME 13 www.nature.com/reviews/neuro
© 2012 Macmillan Publishers Limited. All rights reserved
approximately summarized as KB < KA < KC
22
(meaning that it is most sensitive to
bending orces and least sensitive to com-
pression forces). The neuronal plasma
membrane’s extreme sensitivity to bending
deformation mediates its ability to exocy-
tose and recapture vesicles, as well as to
respond to protrusive and repulsive forces
experienced during growth and motility.
The intrinsic viscoelastic properties of
neuronal membranes are further governed
by cytoskeletal elements, which provide
structural tension within acell.
Actin and spectrins. Protoplasm was first
described as having both viscous and elastic
properties through observations that com-
bined optical microscopy and force-gener-
ation using magnetic fields to manipulate
protoplasm-embedded particles23. The
pioneering observations made by William
Seifriz (1924) and others ultimately led to
the realization that cells possess some type
of cytoskeleton. Today, we know that actin
fibres form part of this cytoskeleton, which
shows dynamic structural plasticity24 and
functions as a three-dimensional array of
force transducers25. The polymerization
and depolymerization of actin monomers
(G-actin) into actin polymers (F-actin) gen-
erates mechanical forces that are important
for many cellular processes, such as gener-
ating cell membrane propulsion and pro-
trusion, counteracting plasma membrane
tension and deformation changes during
endo- and exocytosis, and acting as molecu-
lar tension sensors to regulate numerous
aspects of intracellular homeostasis. The
energy required for actin to generate force
arises from a chemical potential differ-
ence between G-actin and F-actin. Actin
polymerization, and the chemical potential
difference (Gibb’s free energy) required for it,
occurs when the amount of G-actin reaches
a critical concentration at which the free
energy of monomeric actin exceeds the free
energy of F-actin26,27.
When polymerizing actin filaments
approach a biological load, such as a
plasma membrane, they generate push-
ing forces (Fpush), and thermal fluctua-
tions enable the continued incorporation
of G-actin monomers into F-actin. This
actin elongation is thought to resemble
a ‘Brownian ratchet’28, as random thermal
fluctuations enable a gear-like churning
of F-actin polymerization29. The elonga-
tion of F-actin will continue to occur
until the counteracting load forces stall
polymerization at a thermodynamic limit
commonly referred to as the stalling force
(Fstall ≈ 1 pN)30,31. Besides stalling, actin can
buckle under other forces (Fbuckle), allowing
it to continue elongating along a bound-
ary30,31 (FIG.1b). F-actin fibre bundles can
generate forces of several kPa (nN/μm2)
by contacting surface loads from differ-
ent angles and continuously undergoing
branch formation and elongation31,32. Actin
motor proteins (known as myosins), GTP-
binding proteins (such as RAS, RAC, RHO
and CDC42) and a host of other proteins
can influence forces generated by actin
through various mechanisms33,34.
Actin-generated forces regulate
axonal growth cone dynamics35. Growth
cones have a low elastic modulus
(E = 106 ± 21 Pa; 1 Pa = 1 pN/μm2) and can
generate internal stress on the order of
30 Pa36. As actin polymerizes at the lead-
ing edge of growing axons, growth cones
begin to form focal adhesions with ECM
proteins to navigate their environment.
Growth cones are weak (that is, they are
not capable of generating high mechanical
stress) and soft (that is, they are not rigid
or stiff) force generators, which renders
them particularly sensitive to the mechan-
ical properties of their environment36. The
dynamics of retrograde flow of F-actin
can change abruptly to accelerate the
growth of embryonic chick forebrain filo-
podia when they encounter a substrate
stiffness of about 1 kPa37. These proper-
ties might encourage mechanically tuned
synapse formation (see ‘Patterned synapse
formation’). On the postsynaptic side of
synapses, actin is an established regulator
of dendritic spine formation and plastic-
ity38. However, quantitative descriptions
of how actin spatially and temporally dis-
tributes mechanical forces in spines while
working towards such outcomes are lack-
ing. Neuroscience can begin to bridge these
gaps by adopting experimental methods
used in the study of mechanobiology (see
‘Tools for studying mechanobiology’).
Dendritic spine plasticity is also
affected by spectrins, which are cytoskel-
etal proteins forming spring-like tetram-
ers that interact with actin and thereby
Glossary
Axon blebbing
The appearance of pathologically swollen regions (blebs)
along an axon, which are thought to result from the local
breakdown of cytoskeleton proteins in response to
mechanical or oxidative stress.
Brownian ratchet
A perpetual motion machine constructed of a paddle wheel,
ratchet and pawl that is useful for describing how work (or
lack thereof) can be generated by random thermal
fluctuations.
Catastrophic depolymerization
This term refers to the dynamic instability of microtubules;
catastrophic depolymerization occurs when they rapidly
switch from a growing to a shrinking state.
Elastic modulus
A numerical value describing the stiffness of a material or
the tendency of a material to be deformed in response to
force. The stiffer a material is, the higher its elastic modulus
will be.
Elastomeric micropost substrate
A microfabricated substrate consisting of an array of tiny
posts made of soft polymers that bend or become deflected
to report traction forces generated by cells growing on them.
Elastomers
Materials such as rubber (usually polymers) with elastic or
viscoelastic properties that enable them to return to their
original state after deformation.
Gibb’s free energy
Usually denoted G, it is the amount of energy available for
work in a closed system at equilibrium under a constant
temperature and pressure.
Magnetic resonance elastography
(MRE). A medical imaging technique used to measure the
stiffness of tissue by introducing shear waves in tissue and
quantifying their wavelengths as they propagate through the
tissue using MRI.
Maxwell material
A material with both viscous and elastic properties; see
viscoelastic material.
Mitotic spindle
A subcellular structure observable during the
metaphase of cell division, when microtubules attach to
kinetochores and begin to generate the forces required
to pull chromosomes apart for subsequent
incorporation into daughter cells.
Poisson’s ratio
The ratio of transverse strain to axial strain in the direction
of the stretching force.
Traction force microscopy
An optical method used to estimate the traction forces
generated by growing cells as reported by the displacement
of fluorescent microbeads embedded in a polyacrylamide
hydrogel substrate.
Trans-gauche isomerization
A process by which fatty acids (acyl chains) experience
changes in their conformational state and develop kinks in
their molecular structure.
Viscoelastic material
Often referred to as a non-Newtonian material, this is a
material that has both viscous and elastic properties and
that experiences strain as a nonlinear function of time when
stress is applied to it.
Viscoelastic relaxation time
The nonlinear recovery time for a viscoelastic material
to return to its original state after experiencing a
deformation force or stress.
PERSPECTIVES
NATURE REVIEWS
|
NEUROSCIENCE VOLUME 13
|
DECEMBER 2012
|
869
© 2012 Macmillan Publishers Limited. All rights reserved
Nature Reviews | Neuroscience
Membrane patch clamp
Single channel
currents baseline
10 ms
Negative pressure
–20 mmHg
–80 mmHg
a
Popen
Pressure
1
0 –120
0
(mmHg)
b
Rest
Ion channel Expansion KA
Increased
conductance
Compression Bending
Increased
conductance
KB
KC
Bending
Increased
conductance
KB
Tension
help to maintain the integrity, stability
and elasticity of the plasma membrane
(FIG.1a). The βIII isoform of spectrin is
crucial for the formation of Purkinje cell
dendrites and spines39, whereas βI spectrin
regulates CA1 hippocampal spine motility
and AMPA receptor current amplitudes40.
It is unknown, however, to what extent
the mechanical consequences mediated by
spectrins contribute to these functional
outcomes. Mechanical coupling forces that
join proteins and other cellular compart-
ments to one another can be mediated by
spectrin and have been shown to influence
presynaptic excitability. For example, βIV
spectrin stabilizes voltage-gated sodium
channels in the axon initial segment and
at nodes of Ranvier41. The activity of
KCNQ2 potassium channels and their
clustering at nodes of Ranvier also depend
on mechanical forces generated by intact
βIV spectrin42. Other mechanical effects
on the activity of ion channels, which may
in part be mediated by the interaction
of spectrin with plasma membranes and
proteins, are discussed below (see ‘Ion
channels’).
Microtubules and neurofilaments.
Microtubules can exert molecular forces, as
observed by their effects on chromosome
separation during mitotic spindle forma-
tion. Microtubules generate forces in cells
through the polymerization of αβ-tubulin
dimers or by catastrophic depolymerization
events that result from dynamic microtubule
instability24,43. In addition, microtubules can
store elastic energy from frequent bend-
ing and can experience breaking, which
has been proposed to mediate mechano-
chemical signalling in cells44. Microtubules
generate pushing forces (Fpush) or pulling
forces (Fpull), thereby providing structural
support to membranes and proteins or
transporting molecular cargo (FIG.1b). The
maximum force a single microtubule can
generate occurs when Fstall ≈ 5 pN45. This
Fstall of microtubules is also consistent with
the stall force of the microtubule motor
protein kinesin46,47. Kinesins form dimers
that bind to microtubule subunits, where
they undergo a rotational conformational
change upon ATP hydrolysis that facili-
tates their movement along microtubules
to transport cargo or assist in microtubule
elongation. Reinforcement from interme-
diate filaments, actomyosin networks and
microtubule-associated proteins in cells is
thought to enable microtubules to handle
large compression loads and withstand
buckling forces of more than 100 pN48.
Besides their classical cargo transport and
structural support actions, microtubules
have recently been shown to have previ-
ously unrealized functions in the regula-
tion of dendritic spine morphology and
synaptic plasticity49,50. The extent to which
microtubule-generated forces coordinate
with actin-generated forces to mediate
changes in spine stability and plasticity is
not yet known, although several possibili-
ties exist (see ‘Signalling between dendritic
spines’).
Neurofilaments represent the most
abundant cytoskeletal protein in myeli-
nated axons and can dynamically interact
with microtubules and other cytoskeletal
elements51,52. Several observations indi-
cate that neurofilaments can modulate
mechanical forces in axons. Structurally,
neurofilaments are long filamentous
fibres that have side-arms giving rise to a
‘whisker-like’ or bottle-brush appearance53
(FIG.1b). The side-arms of neurofilaments
are thought to provide structural support
in axons and thereby to protect them from
compression loads54. Similarly, intermedi-
ate filaments of astrocytes are also thought
to exert a protective role against mechani-
cal forces in the brain, as well as to mediate
responses to injuries caused by mechani-
cal stress55. The radial growth of axons is
mediated by neurofilaments56, and this
process might rely partly on the long-range
repulsive forces that neurofilaments gener-
ate between each other57. Several models
exist for how neurofilaments interact with
one another and with other cytoskeletal
elements52. In some models, neurofila-
ments form crossbridges with one another,
and under certain conditions invitro they
can form polymeric gels with E >100 Pa58.
Collectively, neurofilaments and interme-
diate filaments serve structural and pro-
tective support roles in neurons and glia,
respectively, by functioning as abundant
protein elastomers.
Figure 2 | Ion channel activity is sensitive to membrane mechanics. a | A general approach to the
study of how mechanical forces affect ion channel activity. A modified pressure-clamp experiment
is shown in which the activity of single ion channels is recorded using conventional inside-out or
outside-out membrane patch clamp methods (left). When negative pressure (suction) is applied to
the membrane, an increase in mechanosensitive channel activity can be recorded (middle). The prob-
ability of channel opening (Popen) can be represented as a function of pressure applied to the mem-
brane (right). b | An ion channel in a plasma membrane under resting conditions (top left). Membrane
expansion, compression, bending and tension can elicit changes in the conformational structure of
the ion channel to modulate its opening and membrane conductance.
PERSPECTIVES
870
|
DECEMBER 2012
|
VOLUME 13 www.nature.com/reviews/neuro
© 2012 Macmillan Publishers Limited. All rights reserved
ECM and cell adhesion molecules. By
connecting the plasma membrane and
cytoskeleton of one cell to another, cell
adhesion molecules and ECM proteins
act as a dynamic polymer matrix to sup-
port the function of cellular networks and
tissues. Besides providing a mechanical
support framework for neurons and glia,
ECM molecules such as reelin have well-
established roles in the induction, expres-
sion and maintenance of synaptic plasticity
that involve facilitating the turnover of
neurotransmitter receptors, modulating
dendritic spine morphogenesis and regu-
lating ion channel activity59. Several other
ECM proteins, such as collagens, laminins
and fibronectins, also provide mechani-
cal support networks that are required for
normal CNS growth, development and
plasticity. Highlighting the integral role of
the ECM in brain function, its dysregu-
lation has been implicated in a host of
brain disorders ranging from neurological
diseases such as epilepsy60 to neuropsychi-
atric diseases such as schizophrenia61. Cell
adhesion molecules enable force transduc-
tion and sensing to occur among neurons,
glia and theECM.
One family of cell adhesion mol-
ecules, the integrins, are transmembrane
receptors that link the cytoskeleton to
ECM proteins (for example, fibronectin)
through interactions with actin. Single
integrin molecules can form adhesions
with fibronectin that have a bond break-
ing force (the force required to dissociate
an adhesion) of about 20 pN62. In neurons,
integrins regulate synapse formation and
maturation63 and provide molecular sup-
port for a host of ECM signalling mecha-
nisms that influence synaptic homeostasis
and plasticity59. Another family of trans-
membrane cell adhesion molecules, the
cadherins, regulate cellular structure
and stability by forming adhesion sites
between cells that express other cadherins
and cell adhesion molecules. Neuronal
cadherin (NCad) binds to its partners
with binding forces of about 40 pN64.
Through the actions of these adhesion
forces, NCad mediates some aspects of
dendritic spine stability65 and is thought to
have other roles in synapse formation and
neuronal plasticity66. Besides their direct
coupling actions, both integrins and cad-
herins can transduce forces in cells. Such
actions might underlie previously unreal-
ized mechanical signalling mechanisms
between presynaptic and postsynaptic
compartments (see ‘Physical coupling at
synapses’).
Ion channels. Pressure, tension, stretch and
stress at a plasma membrane can activate a
broad range of mechanosensitive channels
(MSCs) in the CNS, as well as in sensory
systems1,67. The gating mechanisms that
underlie MSC activity involve a number of
complex membrane deformations (BOX1)
and membrane–protein interactions1,68,69
(FIG.2). Besides the bulk effects of pressure
and tension on plasma membranes, inter-
molecular mechanical forces are generated
when the hydrophobic regions of a protein
and a lipid try to constrain themselves to
each other’s physical lengths (hydrophobic
matching; Fmatch; FIG.1b). These and other
interaction forces (Finteract) arising from pro-
tein inclusion in membranes can also modu-
late ion channel activity1,68,69 (FIG.1b).
Many polymodal channels from diverse
families, including the transient recep-
tor potential (TRP) channels, two-pore
domain potassium (K2P) channels and
calcium-activated potassium (BK) chan-
nels, are modulated by membrane defor-
mations67,69,70 (FIG.2). Recent advances in
our knowledge of ion channel biophysics
indicate that the classic voltage-sensing
mechanisms of many voltage-gated chan-
nels (VGCs), for example voltage-gated
sodium (NaV), potassium (KV), and
calcium (CaV) channels, are sensitive to
mechanical fluctuations in plasma mem-
branes68–70. Examples of different types of
channels that are expressed in the brain
and have mechanosensitive properties are
listed in TABLE1.
More evidence for the influence of
membrane mechanics on ion channel
activity comes from thermodynamic
investigations into the mechanisms of
action of some anaesthetics. Ketamine and
isoflurane are thought to act by increas-
ing the lateral pressure profile of lipids,
which alters channel activity71,72. Even
small membrane deformations with length
scales of a few angstroms are sufficient to
affect channel behaviour1,68. Accepting that
micromechanical forces influence channel
gating raises important issues for neurosci-
ence — the predominant one being that our
conventional understanding of neuronal
excitability does not account for cellular-
mechanical consequences. How does the
realization that many ion channels (includ-
ing voltage-gated ones) are mechanosensi-
tive affect our comprehension of neuronal
activity and plasticity? Confronting this
problem seems to represent a particularly
difficult challenge, especially as structural
changes (tension and stress) at the plasma
membrane of neurons occur continually in
the brain. For example, inserting ion chan-
nels into a postsynaptic membrane or mod-
ulating the rate of vesicle exocytosis would
almost certainly lead to dynamic changes
in synaptic membrane tension and stress.
Addressing the consequences of these
mechanical changes for ion channel activ-
ity and synaptic signalling will increase our
overall understanding of the mechanisms
that underlie brain function.
Consequences of mechanical forces
Physical coupling at synapses. Synapses
comprise discrete compartments coupled
by the cytoskeleton, cell adhesion mol-
ecules and ECM proteins, each of which
exerts adhesion forces (Fadhesion) on synaptic
Box 1 | Losing mechanical energy in the brain?
During time-lapse optical imaging of live brain circuits, field stimulation almost always evokes
micrometre-scale movement artefacts near stimulating electrodes. Depending on the stimulus
parameters and other factors, these movements stem from the effects of electrical fields on
polarized viscoelastic cells. Owing to the interfacial polarization of cell membranes, a dipole is
generated when cells are subjected to an electrical field. Thus, voltage pulses acting at viscoelastic
membrane interfaces can generate elastic strain with a magnitude that depends on the electrical
field strength and the effective polarization of cells in the field2,115. The area expansion moduli (KA)
of erythrocytes, Chinese hamster ovary (CHO) cells and other cells have been shown to vary, with
cells elongating by up to 10% in response to voltage pulses116,117.
Even during action potential firing, axons undergo mechanical changes in which slight
volumetric changes propagate throughout the axonal membrane4,5. Thus, the motion artefacts
arising from field stimulation seem to be a physicochemical property of membrane responses to
electrical activity and probably cannot be avoided. Given our growing awareness that many ion
channels are mechanosensitive, the consequences of plasma membrane deformations in response
to electrical fields are important when considering the interpretations of large numbers of
experiments on synaptic plasticity. It is not clear how one can fully comprehend mechanisms of
synaptic plasticity without considering the mechanical forces influenced by the delivery of
electrical or chemical stimulation paradigms. Mechanical energy cannot be simply lost in the brain.
As it seems difficult for neuroscience to escape the influence of mechanobiology on brain function,
we should benefit by expanding our study of it.
PERSPECTIVES
NATURE REVIEWS
|
NEUROSCIENCE VOLUME 13
|
DECEMBER 2012
|
871
© 2012 Macmillan Publishers Limited. All rights reserved
elements. Presynaptic and postsynaptic
compartments can change their struc-
tures73 and physiological strengths74 in a
correlated manner. However, we do not
know whether mechanical forces trans-
duced by one of these compartments can
trigger functional changes in the other.
Do dendritic spines transmit mechanical
forces to presynaptic compartments to
influence neurotransmitter release or recy-
cling? Several observations with respect to
synaptic coupling, as well as the influence
of presynaptic membrane tension on neu-
rotransmitter vesicle dynamics, support
this possibility.
Depending on the cellular-mechanical
matching of a synapse (the degree of equal-
ity in the stiffness or elasticity between
connected presynaptic and postsynaptic
compartments), it seems likely that a spine
can exert retraction forces (Fretract), twitching
forces (Ftwitch) or extension forces (Fextend) on
its presynaptic partner to trigger functional
changes in presynaptic membrane tension
or bending75 (FIG.3a). If a spine generates a
retraction force that is greater than the bend-
ing modulus of the presynaptic membrane,
but less than the bond-breaking forces of
synaptic adhesion molecules, then tension in
the presynaptic membrane should be influ-
enced by the spine’s retraction. At hippocam-
pal synapses, the motility of a spine is often
coupled to the motility of its presynaptic
bouton partner, and expansion of one com-
partment leads to contraction of the other76.
Presynaptic membrane tension can influence
synaptic vesicle organization75 and modulate
the rate of synaptic vesicle exocytosis in an
integrin-mediated manner77. In fact, since
the seminal descriptions of miniature end
plate potentials provided by Fatt and Katz78,
it has been recognized that mechanical forces
can modulate synaptic transmission78. These
and other observations suggest that mechan-
ical signals can be functionally transmitted
in the brain from one synaptic compartment
to the other through mechanisms mediated
by interactions between the cytoskeleton and
cell adhesion molecules (FIG.3a). Evidence of
similar adhesion-mediated changes between
presynaptic and postsynaptic partners has
been observed at neuromuscular junctions
during muscle growth79. Mechanobiological
studies in neuroscience should aim to
determine to what degree anterograde and
retrograde mechanical signalling at synapses
participates in development, information
transfer and plasticity.
Signalling between dendritic spines.
Signalling among neighbouring dendritic
spines involves the diffusion of several
GTPase molecules80. For instance, RHOA
diffuses out of stimulated spines and lat-
erally about 5 μm through dendrites to
affect the plasticity of nearby spines81. The
fundamental mechanisms that underlie
the downstream signalling consequences
remain unclear. One possibility is that, in
response to synaptic activity, local GTPase-
induced changes in microtubule and
F-actin dynamics enable mechanical signals
to be transmitted among small networks of
cytoskeleton-coupled spines (FIG.3b). For
example, microtubules can behave as elas-
tic rods and exert forces over distances of
about 10 μm45, and the average inter-spine
distance is only about 1 μm. Furthermore,
force–velocity measures of microtubules
(on the order of 1 μm per minute)45 are
consistent with the distance of spatially
segregated spines affected by GTPase activ-
ity, which can persist for around 30min-
utes after spine activation81. These time
and length scales support the hypothesis
that changes in GTPase activity modulate
mechanical forces transmitted by cytoskel-
etal elements as a means of inter-spine
signalling (FIG.3b). Moreover, recent obser-
vations have shown that microtubules can
invade spines and regulate their morphol-
ogy in an activity-dependent manner49,50.
On the basis of the microtubule properties
previously discussed (see ‘Microtubules
and neurofilaments’), it should be expected
that microtubule spine invasion will be
accompanied by local changes in micro-
tubule pushing (Fpush) and pulling (Fpull)
forces. Whether Fpull or Fpush transduced by
cytoskeletal elements can trigger mechani-
cal changes in spatially segregated spines as
a means of inter-spine signalling is not yet
known, but the evidence seems to justify
investigations into this possibility.
Patterned synapse formation. To ensure
optimal growth, cellular processes must
sense the mechanical properties of their
environment while making necessary adjust-
ments in the traction forces they generate.
Although differential modulation of growth
cone mechanical properties and ECM stiff-
ness has been shown to regulate synapse
formation, the intricacies of the mechanical
interplay between these elements are not
completely understood12. The cellular layers
of the rodent hippocampus possess markedly
different rigidities (CA1 stratum pyramidale,
0.14 kPa; CA1 stratum radiatum, 0.20 kPa;
CA3 stratum pyramidale, 0.23 kPa; and CA3
stratum radiatum, 0.31 kPa)9. When cultured
on substrates with rigidities ranging from 0.5
to 7.5 kPa, hippocampal axons grow faster on
softer substrates82. Similarly, neurons from
the embryonic spinal cord develop a fivefold
higher neurite branch density when grown
on soft substrates (0.05 kPa) compared to
more rigid ones (0.55 kPa)83. Interestingly,
the axons of dorsal root ganglion neurons
grow significantly more quickly when they
are mechanically stretched84. The initiation
of growth, and the growth rate of embryonic
chicken forebrain85 and sensory neurons86,
Table 1 | Examples of channels shown to be activated by mechanical forces
Ion type Channel/receptor Refs
K+Shaker (KV1) 118
TRAAK (K2P4.1) 119,120
TREK1 (K2P2.1) 121
HCN2 122
Ca2+-activated K+ (BK) 123
Na+NaV1.5 124,125
Ca2+ N-type 126,127
Mixed cationic NMDA receptor 128,129
TRPC1 130
TRPV1 131
TRPM3 132
ATI TREK1 (Δ1–56 K2P2.1) 133
Cl−CFTR 134
ATI TREK1, alternative translational initiation isoform of the TWIK-related potassium channel; BK, big
potassium channel; CFTR, cystic fibrosis transmembrane conductance regulator channel; HCN, hyperpolariza-
tion-activated cyclic nucleotide-gated channel; K2P
, two-pore domain potassium channel; KV
, voltage-gated
potassium channel; NaV
, voltage-gated sodium channel; TRAAK, TWIK-related arachidonic acid-stimulated
PERSPECTIVES
872
|
DECEMBER 2012
|
VOLUME 13 www.nature.com/reviews/neuro
© 2012 Macmillan Publishers Limited. All rights reserved
Nature Reviews | Neuroscience
Axon
terminal
Fadhesion
(coupling)
Dendritic
spine
Cell adhesion
molecules
F
contract
F
expand
Ftwitch
Actin
Active spine Inactive spine
Signalling
RHOA
Ca2+
Actin
Microtubule
a
F
push
Fpull
b
Layer 1
Layer 2
Layer 3
HighLow
Axon A
Axon B
Mechanically matched
Mechanically matched
Axon growth rate
Growth cone traction forceLow High
c
can also be modulated by mechanically
applied tension. Whether axons of corti-
cal or hippocampal neurons respond to
mechanical stretching in the same manner
is not known. This is a particularly curious
issue as traction force microscopy has shown
that dorsal root ganglion neuron growth
cones can generate significantly greater trac-
tion forces than can hippocampal neurons
(~537 pN versus 71 pN)87. Expressing differ-
ential growth rates as a function of substrate
stiffness or growth cone traction force might
represent a process for mechanically gener-
ating patterned synapse formation (FIG.3c).
Molecular mediators of traction force
generation cooperate with force-sensing
mechanisms to optimize axonal growth
dynamics37. Myosin IIB mediates the gen-
eration of growth cone traction forces88;
the filopodia of superior cervical ganglion
neurons from myosin IIB knockout mice
generate significantly less traction force than
filopodia from wild-type mice (~660 pN ver-
sus 970 pN)88. Both F-actin37 and integrins13
can report substrate rigidity to growing cel-
lular processes, which can in turn optimize
their own mechanical properties to govern
growth dynamics within their environment.
This type of closed-loop feedback system
might be functionally relevant in enabling
the growth rates of axons to keep up with the
hypertrophy of their organism’s growing body
by responding to mechanical stress cues89.
Collectively, the dynamic cellular-mechan-
ical matching principles described above
probably provide mechanisms for tuning
synapse formation during developmental
and adult plasticity (FIG.3c). Characterizing
changes in growth cone traction force across
different anatomical regions, levels of activ-
ity, and stages of development should further
reveal how cellular-mechanical matching
Figure 3 | Functional implications of mechanical force transduction
between synaptic compartments. a | A dendritic spine is coupled to an
axon terminal by cell adhesion molecules, which exert adhesion forces
(Fadhesion) on the synapse. Actin-mediated changes in the spine’s structure
are represented as expansion (Fexpand), contraction (Fcontract) and twitching
(Ftwitch) forces. Such forces generated in either the spine or the axon ter-
minal are hypothesized to produce structural and functional changes in
the opposite compartment. These actions might stem from the transla-
tion of forces from one compartment to the other through cytoskeleton
and cell adhesion molecule synaptic coupling mechanisms. b | Signalling
among neighbouring dendritic spines could involve pushing (Fpush) and
pulling (Fpull) forces exerted by F-actin and microtubules. RHOA diffusion
from an active spine might elicit local changes in forces generated by
cytoskeletal elements. As microtubules can exert mechanical forces
across relatively large distances, they might influence physical contrac-
tion or twitching at neighbouring inactive spines, which could in turn
influence membrane tension and ion channel activity. c | The growth
dynamics of two axons in three different tissue layers, each with different
substrate rigidities. Blue, low stiffness; red, high stiffness. Cellular-
mechanical matching principles (blue growth cone on blue substrate and
red growth cone on red substrate) regulate the ability of growth cones to
generate traction forces and grow.
PERSPECTIVES
NATURE REVIEWS
|
NEUROSCIENCE VOLUME 13
|
DECEMBER 2012
|
873
© 2012 Macmillan Publishers Limited. All rights reserved
principles influence synapse formation. This
seems to be a particularly important issue
because the mechanical properties of the
brain change during development10. In addi-
tion, cellular-mechanical matching might
be important for signalling at gap junctions,
neurovascular junctions and other cell
adhesion sites at which cells with different
mechanical properties physically interact
with one another.
Traumatic injuries. At one end of the spec-
trum, endogenous micromechanical energy
is important for normal brain function, as
discussed above. At the opposite end of the
spectrum, however, substantially greater
mechanical forces acting on the brain can
result in loss of consciousness, irreversible
cognitive dysfunction, progressive neuro-
degeneration and even death90,91. The del-
eterious consequences of concussions and
traumatic brain injury (TBI) have recently
been of central interest to the neuroscience
community. However, we know little about
the cellular-mechanical consequences of
head trauma and how these injuries trigger
a host of deleterious molecular signalling
pathways, which in turn give rise to the
clinical manifestations of TBI92. Consistent
with the cellular-mechanical features of the
brain discussed above, the evidence sug-
gests that traumatic injuries act at least on
protein ion channels, the cytoskeleton and
the plasma membrane.
Diffuse axonal injury resulting in pro-
gressive neurodegeneration stems from
whiplash-like injuries in which the mechani-
cal activation of tetrodotoxin-sensitive
sodium channels ultimately leads to calcium-
mediated excitotoxicity93. Whether a similar
phenomenon is associated with head trauma
resulting in concussion or TBI is not known,
but it warrants investigation. Integrin-
mediated activation of RHO could also be
an important contributor to diffuse axonal
injury after mild TBI94. At the level of the
plasma membrane, mechanoporation result-
ing in increased membrane conductance
occurs in response to TBI in rats95. Similarly,
TBI-induced mechanoporation of the plasma
membrane can trigger axon blebbing and focal
microtubule disruption96. Whether mecha-
noporation perse is a primary consequence
of TBI has not been clearly established. It
seems likely that disruptions to the functions
of the ECM, cytoskeletal elements and cell
membranes after head injury are responsible
for the deleterious consequences. For exam-
ple, intermediate filaments of astrocytes have
been shown to protect against mechanical
injuries to the brain and spinal cord and
to mediate injury responses in the CNS55.
Further, the breakdown and remodelling of
ECM proteins have been implicated in mod-
ulating injury responses to brain trauma97.
How can an expansion of studies investi-
gating the mechanical properties of the CNS
be used to better inform us about the pre-
vention and treatment of TBIs? By gaining
a fundamental grasp on the cytomechanical
features of brain circuits, neuroscience will
be in a better position to address the chal-
lenges associated with TBI. At present, we
do not fully comprehend the extent to which
mechanical energy regulates endogenous
brain function. Thus it remains difficult to
understand how extreme impact forces can
disrupt the natural mechanical properties of
the CNS. These facts alone demonstrate the
need for an increased emphasis on studying
the basic mechanobiology of brain function.
Tools for studying mechanobiology
The mechanical properties of cells and their
networks can be studied using a variety of
methods2,3,98. Modified patch-clamp methods
referred to as pressure-clamp techniques
can be used to study the activity of single
ion channels in response to membrane
deformations99 (FIG.2a). Numerous fluores-
cence microscopy methods have also been
developed for the study of mechanobiol-
ogy. Fluorescence correlation spectroscopy
(FCS) is an optical method in which small
fluctuations in fluorescence are monitored.
Time-correlated single-photon counting
(TCSPC) is another optical method in which
fluorescence decay times can be measured.
Both FCS and TCSPC have been useful for
characterizing mechanical forces in cells
because they depend on physical interac-
tions between photons and their environ-
ment100. Optical methods relying on Förster
resonance energy transfer (FRET) principles
(when an excited fluorophore emits photons
that can excite a neighbouring fluorescent
molecule with a different absorption/emis-
sion spectrum) are useful because they ena-
ble the visualization of molecular mechanical
actions101. A particularly noteworthy FRET
approach involves uniquely designed strain-
sensitive FRET sensors, which can be used
to actively monitor the stress exerted on
ECM and cytoskeletal proteins102.
Atomic force microscopy (AFM)
involves the use of a cantilever probe that
forms a bond with a cell or molecule of
interest. The cantilever can then apply
micromechanical forces to the cell or mol-
ecule, and measure forces generated by it
(FIG.4a). AFM has proven to be a key tool
for the study of molecular and cellular
mechanics103. AFM experiments have
revealed differences in the elasticity of the
rodent brain as a function of anatomical
region9 and have been useful in describing
the force–velocity relationships of grow-
ing actin networks in cells32. Additional
reproducible and quantitative approaches
to mechanobiology have been provided
by trapping methods involving magnetic
particles or optical tweezers, in which
magnetic fields or lasers are used to trap,
control and monitor forces between objects
adhering to cells or molecules2,23 (FIG.4b,c).
Magnetic traps have recently been used to
show that integrin-mediated RHO signal-
ling might underlie some of the conse-
quences of diffuse axonal injury and mild
traumatic brain injury94.
Advances in microengineering have led
to a recent explosion in the development of
mechanobiological research devices2. These
devices include microelectromechanical sys-
tems (MEMS) acting as microactuator arrays
that can generate and measure electrical and
micromechanical changes in cell cultures98;
elastomeric micropost substrates that can
monitor traction forces generated by growing
cells104 (FIG.4d); integrated strain arrays com-
posed of deformable membranes upon which
cell cultures grow while being subjected
to strain105 (FIG.4e); and microfluidic chips
that can apply shear forces by flowing fluids
through channels integrated with cell culture
substrates106 (FIG.4f). Microfluidic chambers
have been useful in applying shear stresses to
axons in order to model and study traumatic
injuries and might have other applications in
the study of brain mechanobiology as their
use becomes more widespread107. Elastomeric
post assemblies have been successfully imple-
mented in studying mechanotransduction
processes in sensory nerves during whole-cell
patch clamp recordings108 but have not yet
been used to study the mechanobiological
properties of central neurons. As engineering
and neuroscience continue their interdis-
ciplinary integration with one another, it is
anticipated that microengineering platforms
that have already proven to be useful for the
study of mechanobiology will foster investiga-
tions of brain function and dysfunction at a
cellular-mechanicallevel.
Ultrasound is a potentially valuable tool
to study the mechanobiology of intact brain
circuits. It can be transmitted and focused
through tissues, including skull bone, to
image the brain or to influence its physiologi-
cal activity through mechanical actions109.
The mechanical bioeffects of transcranial
ultrasound can be used noninvasively to
stimulate brain circuits110. In an imaging
PERSPECTIVES
874
|
DECEMBER 2012
|
VOLUME 13 www.nature.com/reviews/neuro
© 2012 Macmillan Publishers Limited. All rights reserved
Nature Reviews | Neuroscience
a b
c d
ef
g
AFM probe
Force
Cell
Optical trap
Silica bead
Magnetic
bead Magnetic
field
Focal
adhesions Elastomeric
microposts
Integrated
strain array Shear flow
Microfluidic
chamber
MRI
Passive acoustic driver
Sound waves
Brain (tissue)
Shear waves
-
+
Strain
mode, ultrasound can be used for functional
tissue pulsatility imaging, which moni-
tors the displacement of brain tissue due to
cerebrovascular blood flow111. From a diag-
nostic perspective, it has been consistently
shown that many patients who will develop
Parkinson’s disease show an ultrasound
hyperechogenecity in the substantia nigra
before developing motor impairments112. As
an ultrasound echo occurs during imaging
where there are differences in the mechanical
properties of tissues, the pronounced echo
in patients before the onset of Parkinson’s
disease symptoms seems to indicate that the
early stages of the disease might be associated
with changes in the stiffness or elasticity of
the substantia nigra. Ongoing developments
in exploiting the mechanical interactions
between acoustic fields and the brain are
beginning to explore previously unrealized
applications of ultrasound in diagnosing and
treating brain diseases.
MRE is conducted by generating a strain
elastogram from magnetic resonance imaging
data of tissue responses to externally applied
quasi-static compression or harmonic shear
waves14,113 (FIG.4g). These elastograms are used
in clinical diagnostics, as many diseases are
associated with changes in cellular elastic-
ity113. Recently, MRE has been used to map
and characterize the viscoelastic properties
of the normal, aged and diseased human
brain11,14–18. MRE has also been useful in
characterizing TBI in rodent models114. The
recent increase in the number and breadth
of observations from neuroimaging studies
using MRE indicates a strong and growing
interest in determining how the mechanical
properties of the brain relate to its function
and dysfunction.
As expected, there is a fair degree of vari-
ability in the mechanical quantities obtained
when implementing the methods described
throughout this Perspective. This variability
arises from differences in the accuracy and
sensitivity of any given method, as well as
from the diverse experimental conditions
used. Future mechanobiological studies in
neuroscience should focus on using methods
appropriate for a given question. Further,
the limitations of the mechanobiological
approaches implemented should lead to
caution in drawing conclusions about the
functional implications of findings.
Conclusions and perspectives
The brain is a mechanically sensitive organ,
the properties of which enable endogenous
forces to regulate many aspects of neuronal
function. The influence of mechanical
energy on the brains of living organisms is
omnipresent. For instance, cerebrovascular
blood flow accompanying every heartbeat
in humans generates forces that can dis-
place brain tissue by tens of micrometres111.
Nanoscopic changes in plasma membrane
stress and tension can influence ion channel
activity67, synaptic vesicle clustering75, neuro-
transmitter release77 and axonal growth cone
dynamics35. Therefore, we should not disre-
gard the physical mechanics of the nervous
systems westudy.
Over the past several decades, neuro-
science has been dominated by electro-
physiological, biochemical, molecular
and genetic studies of brain function.
Consequently, the mechanical forces that
influence neuronal processes remain
largely unexplored. Using synaptic plastic-
ity as an example, I would like to illustrate
why mechanobiological investigations
need to be expanded in neuroscience.
Briefly, our modern understanding of
Figure 4 | Experimental approaches useful to the study of mechanobiology in neuroscience.
a | An atomic force microscope (AFM) uses a cantilever-style AFM probe to apply or monitor mechani-
cal forces in cells. b | One example of an optical trap method is shown, in which a pair of lasers form a
weakly focused beam to control the movement of a silica bead attached to a cell or molecule. The cell
or molecule can be pushed or pulled by moving the laser beams, and the resulting forces can be meas -
ured. c | Other trapping methods use magnetic particles bound to cells or molecules. The cell and its
associated magnetic probes are subjected to weak electromagnetic fields to apply external forces
while measuring and monitoring mechanical responses. d | Elastomeric micropost assemblies make it
possible to measure traction forces generated by growing cells. Such methods rely on focal adhesion
sites being formed between a cell and micropost assemblies. Growing cells elicit mechanical deflec-
tions of the microposts from which traction forces can be derived. e | Deformable membranes can be
microfabricated on culture substrates in integrated strain arrays, which allow researchers to apply or
measure strain in cell cultures under a variety of experimental conditions. f | Microfluidic chambers
can be used to apply shear forces, which are generated when fluids are forced to flow across the cells
and through chambered assemblies. g | The image illustrates the basic approach to conducting mag-
netic resonance elastography. Shear waves are introduced into the brain (usually through small head
movements translated using a piston or bite bar) and monitored using MRI to estimate the shear and
elastic moduli of the brain.
PERSPECTIVES
NATURE REVIEWS
|
NEUROSCIENCE VOLUME 13
|
DECEMBER 2012
|
875
© 2012 Macmillan Publishers Limited. All rights reserved
synaptic plasticity is that it involves finely
tuned, calcium-mediated signalling
mechanisms that give rise to potentiated
or depressed synapses. Calcium signalling
itself is known to influence the activity of
several motor proteins, including myosin,
to affect cytoskeletal polymerization and
depolymerization dynamics, and to medi-
ate interactions between cell adhesion mol-
ecules and ECM proteins — all of which
regulate mechanical forces at synapses.
Furthermore, integrins have been shown to
be necessary for the functional maturation
of hippocampal synapses63. Therefore it
appears that plasticity also depends, albeit
to a mostly uncharacterized degree, on
changes in membrane tension and synaptic
adhesion. Knowledge gained from stud-
ies of mechanical forces in neurons will
probably not disprove our current working
models; rather, it should enable us to refine
and expand uponthem.
In conclusion, the amalgamation of
neuroscience and mechanobiology (neu-
romechanobiology) will provide greater
insight into how endogenous mechanical
forces physically collide with conventional
signal transduction pathways to govern
neuronal development and plasticity. I
anticipate that neuromechanobiology will
reveal basic brain functions that require the
regulated contraction, expansion, pushing,
pulling and relaxing of neurons, glia and
their molecular and cellular components. At
a minimum, neuromechanobiology studies
will improve our understanding of the pri-
mary sequela associated with concussion,
traumatic brain injury and disease states
that affect neuronal viscoelasticity.
William J.Tyler is at The Virginia Tech Carilion Research
Institute, School of Biomedical Engineering and
Sciences, Virginia Tech, Roanoke, Virginia 24016, USA.
e-mail: wtyler@vt.edu
doi:10.1038/nrn3383
1. Hamill, O.P. & Martinac, B. Molecular basis of
mechanotransduction in living cells. Physiol. Rev. 81 ,
685–740 (2001).
2. Kim, D.H., Wong, P.K., Park, J., Levchenko, A. &
Sun, Y. Microengineered platforms for cell
mechanobiology. Annu. Rev. Biomed. Eng. 11,
203–233 (2009).
3. Eyckmans, J., Boudou, T., Yu, X. & Chen, C.S.
A hitchhiker’s guide to mechanobiology. Dev. Cell 21,
35–47 (2011).
This paper provides a comprehensive review of the
field of mechanobiology.
4. Hill, D.K. The volume change resulting from
stimulation of a giant nerve fibre. J.Physiol. 111 ,
304–327 (1950).
5. Tasaki, I., Kusano, K. & Byrne, P.M. Rapid mechanical
and thermal changes in the garfish olfactory nerve
associated with a propagated impulse. Biophys. J. 55,
1033–1040 (1989).
This study describes the propagation of a
mechanical wave along axons during action
potential firing.
6. Kim, G.H., Kosterin, P., Obaid, A.L. & Salzberg, B.M.
A mechanical spike accompanies the action potential
in Mammalian nerve terminals. Biophys. J. 92,
3122–3129 (2007).
7. Crick, F. Do dendritic spines twitch? Trends Neurosci.
5, 44–46 (1982).
8. Star, E.N., Kwiatkowski, D.J. & Murthy, V.N. Rapid
turnover of actin in dendritic spines and its regulation
by activity. Nature Neurosci. 5, 239–246 (2002).
9. Elkin, B.S., Azeloglu, E.U., Costa, K.D. &
Morrison, B. Mechanical heterogeneity of the rat
hippocampus measured by atomic force microscope
indentation. J.Neurotrauma 24, 812–822 (2007).
10. Gefen, A., Gefen, N., Zhu, Q., Raghupathi, R. &
Margulies, S.S. Age-dependent changes in material
properties of the brain and braincase of the rat.
J.Neurotrauma 20, 1163–1177 (2003).
11. Kruse, S.A. etal. Magnetic resonance elastography of
the brain. Neuroimage 39, 231–237 (2008).
12. Moore, S.W. & Sheetz, M.P. Biophysics of substrate
interaction: influence on neural motility,
differentiation, and repair. Dev. Neurobiol. 71,
1090–1101 (2011).
13. Moore, S.W., Roca-Cusachs, P. & Sheetz, M.P.
Stretchy proteins on stretchy substrates: the
important elements of integrin-mediated rigidity
sensing. Dev. Cell 19, 194–206 (2010).
14. McCracken, P.J., Manduca, A., Felmlee, J. &
Ehman, R.L. Mechanical transient-based magnetic
resonance elastography. Magn. Reson. Med. 53,
628–639 (2005).
15. Zhang, J., Green, M.A., Sinkus, R. & Bilston, L.E.
Viscoelastic properties of human cerebellum using
magnetic resonance elastography. J.Biomech. 44,
1909–1913 (2011).
16. Sack, I., Streitberger, K.J., Krefting, D., Paul, F. &
Braun, J. The influence of physiological aging and
atrophy on brain viscoelastic properties in humans.
PLoS ONE 6, e23451 (2011).
This paper uses MRE to characterize changes in the
rigidity of the human brain as a function of age.
17. Wuerfel, J. etal. MR-elastography reveals degradation
of tissue integrity in multiple sclerosis. Neuroimage
49, 2520–2525 (2010).
18. Murphy, M.C. etal. Decreased brain stiffness in
Alzheimer’s disease determined by magnetic
resonance elastography. J.Magn. Reson. Imaging 34,
494–498 (2011).
19. Crawford, G.E. & Earnshaw, J.C. Viscoelastic relaxation
of bilayer lipid membranes. Frequency-dependent
tension and membrane viscosity. Biophys. J. 52,
87–94 (1987).
20. Pastor, R.W. & Feller, S.E. Time scales of lipid
dynamics and molecular dynamics. Biol. Membr. 1,
4–29 (1996).
21. Almeida, P.F.F. & Vaz, W.L.C. in Structure and
Dynamics of Membranes: From Cells to Vesicles Vol. 1
(eds Lipowsky, R. & Sackmann, E.) 305–357 (Elsevier,
1995).
22. Evans, E.A. & Hochmuth, R.M. in Current Topics in
Membranes and Transport Vol. 10 (eds Bronner, F. &
Kleinzeller, A.) 1–64 (Academic Press, 1978).
23. Seifriz, W. An elastic value of protoplasm, with further
observations on the viscosity of protoplasm. J.Exp.
Biol. 2, 1–11 (1924).
24. Kueh, H.Y. & Mitchison, T.J. Structural plasticity in
actin and tubulin polymer dynamics. Science 325,
960–963 (2009).
25. Galkin, V.E., Orlova, A. & Egelman, E.H. Actin
filaments as tension sensors. Curr. Biol. 22,
R96–R101 (2012).
26. Hill, T.L. & Kirschner, M.W. Bioenergetics and kinetics
of microtubule and actin filament assembly-
disassembly. Int. Rev. Cytol. 78, 1–125 (1982).
27. Theriot, J.A. The polymerization motor. Traffic 1,
19–28 (2000).
28. Feynman, R.P., Leighton, R.B. & Sands, M. in The
Feynman Lectures on Physics Vol. 1, Ch. 46, 46–49
(Addison-Wessley, 1963).
29. Peskin, C.S., Odell, G.M. & Oster, G.F. Cellular
motions and thermal fluctuations: the Brownian
ratchet. Biophys. J. 65, 316–324 (1993).
30. Hill, T.L. & Kirschner, M.W. Subunit treadmilling of
microtubules or actin in the presence of cellular
barriers: possible conversion of chemical free energy
into mechanical work. Proc. Natl Acad. Sci. USA 79,
490–494 (1982).
31. Footer, M.J., Kerssemakers, J.W., Theriot, J.A. &
Dogterom, M. Direct measurement of force generation
by actin filament polymerization using an optical trap.
Proc. Natl Acad. Sci. USA 104, 2181–2186 (2007).
Using optical trapping methods, this paper
quantifies the force generated by small bundles of
F‑actin.
32. Parekh, S.H., Chaudhuri, O., Theriot, J.A. & Fletcher,
D.A. Loading history determines the velocity of actin-
network growth. Nature Cell Biol. 7, 1219–1223
(2005).
33. Kosztin, I., Bruinsma, R., O’Lague, P. & Schulten, K.
Mechanical force generation by G proteins. Proc. Natl
Acad. Sci. USA 99, 3575–3580 (2002).
34. Schliwa, M. & Woehlke, G. Molecular motors. Nature
422, 759–765 (2003).
35. Smith, S.J. Neuronal cytomechanics: the actin-based
motility of growth cones. Science 242, 708–715
(1988).
36. Betz, T., Koch, D., Lu, Y.B., Franze, K. & Kas, J.A.
Growth cones as soft and weak force generators. Proc.
Natl Acad. Sci. USA 108, 13420–13425 (2011).
37. Chan, C.E. & Odde, D.J. Traction dynamics of
filopodia on compliant substrates. Science 322,
1687–1691 (2008).
This paper provides a quantitative description of a
‘motor‑clutch’ model in which retrograde F‑actin flow
can differentially generate traction forces in growth
cones of neurons.
38. Matus, A. Actin-based plasticity in dendritic spines.
Science 290, 754–758 (2000).
39. Gao, Y. etal. β-III spectrin is critical for development of
Purkinje cell dendritic tree and spine morphogenesis.
J.Neurosci. 31, 16581–16590 (2011).
40. Nestor, M.W., Cai, X., Stone, M.R., Bloch, R.J. &
Thompson, S.M. The actin binding domain of
βI-spectrin regulates the morphological and functional
dynamics of dendritic spines. PLoS ONE 6, e16197
(2011).
41. Komada, M. & Soriano, P. βIV-spectrin regulates
sodium channel clustering through ankyrin-G at axon
initial segments and nodes of Ranvier. J.Cell Biol.
156, 337–348 (2002).
42. Devaux, J.J. The C-terminal domain of ssIV-spectrin is
crucial for KCNQ2 aggregation and excitability at
nodes of Ranvier. J.Physiol. 588, 4719–4730 (2010).
43. Desai, A. & Mitchison, T.J. Microtubule
polymerization dynamics. Annu. Rev. Cell Dev. Biol.
13, 83–117 (1997).
44. Odde, D.J., Ma, L., Briggs, A.H., DeMarco, A. &
Kirschner, M.W. Microtubule bending and breaking in
living fibroblast cells. J.Cell Sci. 112 , 3283–3288
(1999).
This paper describes the elastic energy stored in
microtubules as they are bent.
45. Dogterom, M. & Yurke, B. Measurement of the force-
velocity relation for growing microtubules. Science
278, 856–860 (1997).
This article provides a quantitative description of
the mechanical forces generated by growing
microtubules.
46. Svoboda, K. & Block, S.M. Force and velocity
measured for single kinesin molecules. Cell 77,
773–784 (1994).
47. Visscher, K., Schnitzer, M.J. & Block, S.M. Single
kinesin molecules studied with a molecular force
clamp. Nature 400, 184–189 (1999).
48. Brangwynne, C.P. etal. Microtubules can bear
enhanced compressive loads in living cells because of
lateral reinforcement. J.Cell Biol. 173, 733–741
(2006).
49. Hu, X., Viesselmann, C., Nam, S., Merriam, E. &
Dent, E.W. Activity-dependent dynamic microtubule
invasion of dendritic spines. J.Neurosci. 28,
13094–13105 (2008).
This article provides evidence that microtubules
can affect hippocampal and cortical dendritic
spines in an activity‑dependent manner.
50. Jaworski, J. etal. Dynamic microtubules regulate
dendritic spine morphology and synaptic plasticity.
Neuron 61, 85–100 (2009).
51. Lee, M.K. & Cleveland, D.W. Neuronal intermediate
filaments. Annu. Rev. Neurosci. 19, 187–217 (1996).
52. Mukhopadhyay, R., Kumar, S. & Hoh, J.H. Molecular
mechanisms for organizing the neuronal cytoskeleton.
Bioessays 26, 1017–1025 (2004).
53. Geisler, N. & Weber, K. Self-assembly in vitro of the
68,000 molecular weight component of the
mammalian neurofilament triplet proteins into
intermediate-sized filaments. J.Mol. Biol. 151,
565–571 (1981).
54. Brown, H.G. & Hoh, J.H. Entropic exclusion by
neurofilament sidearms: a mechanism for maintaining
interfilament spacing. Biochemistry 36,
15035–15040 (1997).
PERSPECTIVES
876
|
DECEMBER 2012
|
VOLUME 13 www.nature.com/reviews/neuro
© 2012 Macmillan Publishers Limited. All rights reserved
55. Pekny, M. & Lane, E.B. Intermediate filaments and
stress. Exp. Cell Res. 313, 2244–2254 (2007).
56. Cleveland, D.W. etal. Involvement of neurofilaments
in the radial growth of axons. J.Cell Sci. Suppl. 15,
85–95 (1991).
57. Kumar, S., Yin, X., Trapp, B.D., Paulaitis, M.E. &
Hoh, J.H. Role of long-range repulsive forces in
organizing axonal neurofilament distributions:
evidence from mice deficient in myelin-associated
glycoprotein. J.Neurosci. Res. 68, 681–690
(2002).
58. Leterrier, J.F., Kas, J., Hartwig, J., Vegners, R. &
Janmey, P.A. Mechanical effects of neurofilament
cross-bridges. Modulation by phosphorylation, lipids,
and interactions with F-actin. J.Biol. Chem. 271,
15687–15694 (1996).
59. Dityatev, A., Schachner, M. & Sonderegger, P. The dual
role of the extracellular matrix in synaptic plasticity
and homeostasis. Nature Rev. Neurosci. 11 , 735–746
(2010).
60. Dityatev, A. & Fellin, T. Extracellular matrix in
plasticity and epileptogenesis. Neuron Glia Biol. 4,
235–247 (2008).
61. Pantazopoulos, H., Woo, T.U., Lim, M.P., Lange, N. &
Berretta, S. Extracellular matrix-glial abnormalities in
the amygdala and entorhinal cortex of subjects
diagnosed with schizophrenia. Arch. Gen. Psychiatry
67, 155–166 (2010).
62. Thoumine, O., Kocian, P., Kottelat, A. & Meister, J.J.
Short-term binding of fibroblasts to fibronectin: optical
tweezers experiments and probabilistic analysis. Eur.
Biophys. J. 29, 398–408 (2000).
63. Chavis, P. & Westbrook, G. Integrins mediate
functional pre- and postsynaptic maturation at a
hippocampal synapse. Nature 411 , 317–321 (2001).
64. Baumgartner, W., Golenhofen, N., Grundhofer, N.,
Wiegand, J. & Drenckhahn, D. Ca2+ dependency of
N-cadherin function probed by laser tweezer and
atomic force microscopy. J.Neurosci. 23,
11008–11014 (2003).
65. Mendez, P., De Roo, M., Poglia, L., Klauser, P. &
Muller, D. N-cadherin mediates plasticity-induced
long-term spine stabilization. J.Cell Biol. 189,
589–600 (2010).
66. Arikkath, J. & Reichardt, L.F. Cadherins and catenins
at synapses: roles in synaptogenesis and synaptic
plasticity. Trends Neurosci. 31 , 487–494 (2008).
67. Arnadottir, J. & Chalfie, M. Eukaryotic
mechanosensitive channels. Annu. Rev. Biophys. 39,
111–137 (2010).
68. Reeves, D., Ursell, T., Sens, P., Kondev, J. & Phillips, R.
Membrane mechanics as a probe of ion-channel
gating mechanisms. Phys. Rev. E Stat. Nonlin. Soft
Matter Phys. 78, 041901 (2008).
69. Sukharev, S. & Corey, D.P. Mechanosensitive
channels: multiplicity of families and gating
paradigms. Sci. STKE 2004, re4 (2004).
70. Morris, C.E. Voltage-gated channel
mechanosensitivity: fact or friction? Front. Physiol. 2,
25 (2011).
71. Cantor, R.S. The lateral pressure profile in
membranes: a physical mechanism of general
anesthesia. Biochemistry 36, 2339–2344 (1997).
72. Jerabek, H., Pabst, G., Rappolt, M. & Stockner, T.
Membrane-mediated effect on ion channels induced
by the anesthetic drug ketamine. J.Am. Chem. Soc.
132, 7990–7997 (2010).
73. Schikorski, T. & Stevens, C.F. Quantitative
ultrastructural analysis of hippocampal excitatory
synapses. J.Neurosci. 17, 5858–5867 (1997).
74. Murthy, V.N., Schikorski, T., Stevens, C.F. & Zhu, Y.
Inactivity produces increases in neurotransmitter
release and synapse size. Neuron 32, 673–682
(2001).
75. Siechen, S., Yang, S., Chiba, A. & Saif, T. Mechanical
tension contributes to clustering of neurotransmitter
vesicles at presynaptic terminals. Proc. Natl Acad. Sci.
USA 106, 12611–12616 (2009).
This elegant study shows that mechanical tension
at neuromuscular junction synapses can accelerate
synaptic vesicle clustering in presynaptic terminals.
76. Umeda, T., Ebihara, T. & Okabe, S. Simultaneous
observation of stably associated presynaptic
varicosities and postsynaptic spines: morphological
alterations of CA3-CA1 synapses in hippocampal slice
cultures. Mol. Cell. Neurosci. 28, 264–274 (2005).
This article provides evidence that hippocampal
synapses are tightly mechanically coupled.
77. Chen, B.M. & Grinnell, A.D. Integrins and modulation
of transmitter release from motor nerve terminals by
stretch. Science 269, 1578–1580 (1995).
This paper shows that presynaptic membrane
stretch can increase synaptic vesicle exocytosis in
an integrin‑mediated fashion at neuromuscular
junctions.
78. Fatt, P. & Katz, B. Spontaneous subthreshold activity
at motor nerve endings. J.Physiol. 117 , 109–128
(1952).
79. Balice-Gordon, R.J. & Lichtman, J.W. In vivo
visualization of the growth of pre- and postsynaptic
elements of neuromuscular junctions in the mouse.
J.Neurosci. 10, 894–908 (1990).
80. Yasuda, R. & Murakoshi, H. The mechanisms
underlying the spatial spreading of signaling activity.
Curr. Opin. Neurobiol. 21, 313–321 (2011).
81. Murakoshi, H., Wang, H. & Yasuda, R. Local,
persistent activation of Rho GTPases during plasticity
of single dendritic spines. Nature 472, 100–104
(2011).
82. Kostic, A., Sap, J. & Sheetz, M.P. RPTPα is required
for rigidity-dependent inhibition of extension and
differentiation of hippocampal neurons. J.Cell Sci.
120, 3895–3904 (2007).
83. Flanagan, L.A., Ju, Y.E., Marg, B., Osterfield, M. &
Janmey, P.A. Neurite branching on deformable
substrates. Neuroreport 13, 2411–2415 (2002).
84. Pfister, B.J., Iwata, A., Meaney, D.F. & Smith, D.H.
Extreme stretch growth of integrated axons.
J.Neurosci. 24, 7978–7983 (2004).
85. Fass, J.N. & Odde, D.J. Tensile force-dependent
neurite elicitation via anti-β1 integrin antibody-coated
magnetic beads. Biophys. J. 85, 623–636 (2003).
86. Zheng, J. etal. Tensile regulation of axonal elongation
and initiation. J.Neurosci. 11, 1117–1125 (1991).
87. Koch, D., Rosoff, W.J., Jiang, J., Geller, H.M. &
Urbach, J.S. Strength in the periphery: growth cone
biomechanics and substrate rigidity response in
peripheral and central nervous system neurons.
Biophys. J. 102, 452–460 (2012).
88. Bridgman, P.C., Dave, S., Asnes, C.F., Tullio, A.N. &
Adelstein, R.S. Myosin IIB is required for growth cone
motility. J.Neurosci. 21, 6159–6169 (2001).
89. Bray, D. Axonal growth in response to experimentally
applied mechanical tension. Dev. Biol. 102, 379–389
(1984).
90. Hoge, C.W. etal. Mild traumatic brain injury in U.S.
soldiers returning from Iraq. N.Engl. J.Med. 358,
453–463 (2008).
91. Meaney, D.F. & Smith, D.H. Biomechanics of
concussion. Clin. Sports Med. 30, 19–31 (2011).
92. Farkas, O. & Povlishock, J.T. Cellular and subcellular
change evoked by diffuse traumatic brain injury: a
complex web of change extending far beyond focal
damage. Prog. Brain Res. 161, 43–59 (2007).
93. Wolf, J.A., Stys, P.K., Lusardi, T., Meaney, D. &
Smith, D.H. Traumatic axonal injury induces calcium
influx modulated by tetrodotoxin-sensitive sodium
channels. J.Neurosci. 21 , 1923–1930 (2001).
This study provides evidence that traumatic injuries
to the CNS can involve the mechanical activation of
voltage‑gated sodium channels.
94. Hemphill, M.A. etal. A possible role for integrin
signaling in diffuse axonal injury. PLoS ONE 6,
e22899 (2011).
95. Farkas, O., Lifshitz, J. & Povlishock, J.T.
Mechanoporation induced by diffuse traumatic brain
injury: an irreversible or reversible response to injury?
J.Neurosci. 26, 3130–3140 (2006).
96. Kilinc, D., Gallo, G. & Barbee, K.A. Mechanically-
induced membrane poration causes axonal beading
and localized cytoskeletal damage. Exp. Neurol. 21 2,
422–430 (2008).
97. Lo, E.H., Wang, X. & Cuzner, M.L. Extracellular
proteolysis in brain injury and inflammation: role for
plasminogen activators and matrix
metalloproteinases. J.Neurosci. Res. 69, 1–9
(2002).
98. Rajagopalan, J. & Saif, M.T.MEMS sensors and
microsystems for cell mechanobiology. J.Micromech.
Microeng. 21, 54002–54012 (2011).
99. McBride, D.W.Jr & Hamill, O.P. Pressure-clamp
technique for measurement of the relaxation kinetics
of mechanosensitive channels. Trends Neurosci. 16,
341–345 (1993).
100. Gullapalli, R.R., Tabouillot, T., Mathura, R.,
Dangaria, J.H. & Butler, P.J. Integrated multimodal
microscopy, time-resolved fluorescence, and optical-
trap rheometry: toward single molecule
mechanobiology. J.Biomed. Opt. 12, 014012
(2007).
101. Wang, Y. & Wang, N. FRET and mechanobiology.
Integr. Biol. 1, 565–573 (2009).
102. Meng, F., Suchyna, T.M., Lazakovitch, E.,
Gronostajski, R.M. & Sachs, F. Real time FRET based
detection of mechanical stress in cytoskeletal and
extracellular matrix proteins. Cell. Mol. Bioeng. 4,
148–159 (2011).
103. Azeloglu, E.U. & Costa, K.D. Atomic force microscopy
in mechanobiology: measuring microelastic
heterogeneity of living cells. Methods Mol. Biol. 736,
303–329 (2011).
104. Yang, M.T., Fu, J., Wang, Y.K., Desai, R.A. & Chen,
C.S. Assaying stem cell mechanobiology on
microfabricated elastomeric substrates with
geometrically modulated rigidity. Nature Protoc. 6,
187–213 (2011).
105. Simmons, C.S. etal. Integrated strain array for
cellular mechanobiology studies. J.Micromech.
Microeng. 21, 54016–54025 (2011).
106. Taylor, A.M., Dieterich, D.C., Ito, H.T., Kim, S.A. &
Schuman, E.M. Microfluidic local perfusion chambers
for the visualization and manipulation of synapses.
Neuron 66, 57–68 (2010).
107. Taylor, A.M. etal. A microfluidic culture platform for
CNS axonal injury, regeneration and transport. Nature
Methods 2, 599–605 (2005).
108. Cheng, C.M. etal. Probing localized neural
mechanotransduction through surface-modified
elastomeric matrices and electrophysiology. Nature
Protoc. 5, 714–724 (2010).
109. Dalecki, D. Mechanical bioeffects of ultrasound. Annu.
Rev. Biomed. Eng. 6, 229–248 (2004).
110. Tufail, Y., Yoshihiro, A., Pati, S., Tauchmann, M.L. &
Tyler, W.J. Ultrasonic neuromodulation by brain
stimulation with transcranial ultrasound. Nature
Protoc. 6, 1453–1470 (2011).
111. Kucewicz, J.C. etal. Functional tissue pulsatility
imaging of the brain during visual stimulation.
Ultrasound Med. Biol. 33, 681–690 (2007).
112. Berg, D. Hyperechogenicity of the substantia nigra:
pitfalls in assessment and specificity for Parkinson’s
disease. J.Neural Transm. 11 8 , 453–461 (2011).
113. Mariappan, Y.K., Glaser, K.J. & Ehman, R.L.
Magnetic resonance elastography: a review. Clin. Anat.
23, 497–511 (2010).
114. Boulet, T., Kelso, M.L. & Othman, S.F. Microscopic
magnetic resonance elastography of traumatic brain
injury model. J.Neurosci. Methods 201, 296–306
(2011).
115. Engelhardt, H., Gaub, H. & Sackmann, E. Viscoelastic
properties of erythrocyte membranes in high-
frequency electric fields. Nature 307, 378–380
(1984).
116. Katnik, C. & Waugh, R. Electric fields induce reversible
changes in the surface to volume ratio of micropipette-
aspirated erythrocytes. Biophys. J. 57, 865–875
(1990).
117. MacQueen, L.A., Buschmann, M.D. &
Wertheimer, M.R. Mechanical properties of
mammalian cells in suspension measured by electro-
deformation. J.Micromech. Microeng. 20, 065007
(2010).
118. Tabarean, I.V. & Morris, C.E. Membrane stretch
accelerates activation and slow inactivation in Shaker
channels with S3-S4 linker deletions. Biophys. J. 82,
2982–2994 (2002).
This paper describes the effects of membrane
tension on the activation and inactivation of a
prototypical voltage‑gated channel, the Shaker
potassium channel.
119. Maingret, F., Fosset, M., Lesage, F., Lazdunski, M. &
Honore, E. TRAAK is a mammalian neuronal
mechano-gated K+ channel. J.Biol. Chem. 274,
1381–1387 (1999).
120. Brohawn, S.G., del Marmol, J. & MacKinnon, R.
Crystal structure of the human K2P TRAAK, a lipid-
and mechano-sensitive K+ ion channel. Science 335,
436–441 (2012).
121. Maingret, F., Patel, A.J., Lesage, F., Lazdunski, M. &
Honore, E. Mechano- or acid stimulation, two
interactive modes of activation of the TREK-1
potassium channel. J.Biol. Chem. 274,
26691–26696 (1999).
122. Lin, W., Laitko, U., Juranka, P.F. & Morris, C.E. Dual
stretch responses of mHCN2 pacemaker channels:
accelerated activation, accelerated deactivation.
Biophys. J. 92, 1559–1572 (2007).
123. Niu, X., Qian, X. & Magleby, K.L. Linker-gating ring
complex as passive spring and Ca2+-dependent
machine for a voltage- and Ca2+-activated potassium
channel. Neuron 42, 745–756 (2004).
124. Morris, C.E. & Juranka, P.F. Nav channel
mechanosensitivity: activation and inactivation
PERSPECTIVES
NATURE REVIEWS
|
NEUROSCIENCE VOLUME 13
|
DECEMBER 2012
|
877
© 2012 Macmillan Publishers Limited. All rights reserved
accelerate reversibly with stretch. Biophys. J. 93,
822–833 (2007).
125. Beyder, A. etal. Mechanosensitivity of Nav1.5, a
voltage-sensitive sodium channel. J.Physiol. 588,
4969–4985 (2010).
126. Calabrese, B., Tabarean, I.V., Juranka, P. & Morris,
C.E. Mechanosensitivity of N-type calcium channel
currents. Biophys. J. 83, 2560–2574 (2002).
127. Etzion, Y. & Grossman, Y. Pressure-induced depression of
synaptic transmission in the cerebellar parallel fibre
synapse involves suppression of presynaptic N-type Ca2+
channels. Eur. J.Neurosci. 12, 4007–4016 (2000).
128. Paoletti, P. & Ascher, P. Mechanosensitivity of NMDA
receptors in cultured mouse central neurons. Neuron
13, 645–655 (1994).
The modulation of NMDA receptor activity by
mechanical pressure applied to neuronal membranes
is described in this article.
134. Zhang, W.K. etal. Mechanosensitive gating of CFTR.
Nature Cell Biol. 12, 507–512 (2010).
Acknowledgements
W.J.T is supported by funds from a US Department of
Defense grant from the US Army Research, Development,
and Engineering Command (RDECOM W911NF-09-0431), a
Defense Advanced Research Projects Agency Young Faculty
Award (DARPA N66001-10-1-4032) and a McKnight
Technological Innovation in Neuroscience Award.
Competing interests statement
The author declares no competing financial interests.
129. Singh, P. etal. N-methyl-d-aspartate receptor
mechanosensitivity is governed by C terminus of
NR2B subunit. J.Biol. Chem. 287, 4348–4359
(2012).
130. Maroto, R. etal. TRPC1 forms the stretch-activated
cation channel in vertebrate cells. Nature Cell Biol. 7,
179–185 (2005).
131. Liedtke, W. Handbook of Experimental Pharmacology
Vol. 179 (eds Flockerzi, V. & Nilius, B.) 473–487
(Springer, 2007).
132. Grimm, C., Kraft, R., Sauerbruch, S., Schultz, G. &
Harteneck, C. Molecular and functional characterization
of the melastatin-related cation channel TRPM3.
J.Biol. Chem. 278, 21493–21501 (2003).
133. Thomas, D., Plant, L.D., Wilkens, C.M., McCrossan,
Z.A. & Goldstein, S.A. Alternative translation initiation
in rat brain yields K2P2.1 potassium channels
permeable to sodium. Neuron 58, 859–870 (2008).
FURTHER INFORMATION
William J. Tyler’s homepage: http://www.tylerlab.com
ALL LINKS ARE ACTIVE IN THE ONLINE PDF
PERSPECTIVES
878
|
DECEMBER 2012
|
VOLUME 13 www.nature.com/reviews/neuro
© 2012 Macmillan Publishers Limited. All rights reserved
