Molecular Medicine Program
Ιατρικη Σχολη Πανεπιστημιου Κρητης
Diomedes E. Logothetis
Membrane Structure and Proteins
November 14, 2007
The first part of the lecture will consider the membrane environment that ion channel
proteins reside examining the lipid composition and forces that contribute to the
thermodynamic stability of the lipid bilayer. A brief general introduction to membrane
proteins and methods used to study them is included.
The second part of the lecture will present ion channels that are gated by intracellular
molecules (e.g. ATP, G proteins, cyclic nucleotides) as well as ion channels that are
gated by extracellular molecules (e.g. the neurotransmitters acetylcholine and
1. Know the major classes of natural lipids, their structural characteristics and the properties of
head groups and hydrocarbon chains
2. Understand the thermodynamic basis of lipid and detergent assembly: the hydrophobic
3. Understand the physical connection between the shape of lipid molecules and that of the
aggregates they form: critical packing parameter
4. Understand the connection between the configuration of acyl chains, their packing and the
properties of the bilayer
5. Know the connection between bilayer structure and dynamics and translational diffusion of
lipids and proteins. Experimental approach to the measurement of diffusion of membrane
components: Fluorescence Recovery After Photobleaching
6. Understand the characteristics of the fluid phase bilayer as shown by diffraction experiments
and computer simulations of lipid dynamics
7. Modes of protein-membrane interaction.
8. Prediction of membrane protein structure and topology: Hydropathy analysis and the
9. Lipid-modifications of proteins: hydrophobicity and membrane-binding affinity.
10. Modulation of reversible protein-membrane binding: the myristoyl switch.
11. Know the mechanism by which the metabolic state of the cell is coupled to membrane
electrical events, such as those leading to secretion of insulin.
12. Know the mechanism of activation of G protein-gated K channels, as an example of a
membrane-delimited pathway of regulating the activity of intracellular ligand-gated ion
13. Modulation of Ion channels by soluble second messengers
14. Sensory transduction: Know the role of CNG channels in phototransduction. Understand
how the balance of CNG and K channels gives rise to the quot;dark currentquot;, which is inhibited
during a light flash.
15. Know the subunit composition of nicotinic ACh channels and general topology of the α
16. Know the general activation mechanism for NMDA and non-NMDA channels and role in
• Alberts B., Johnson A., Lewis J., Raff M., Roberts K., Walter P. Molecular Biology of the
Cell. Fourth Edition, Garland Science. pp. 583-614 and pp. 631-657.
• R. B. Gennis. Biomembranes: Molecular Structure and Function, Springer-Verlag, 1989
• Dr. Stephen H. White’s lab, at University of California at Irvine, maintains an interesting
Web site you should visit: http://blanco.biomol.uci.edu
Biological membranes are formed by two layers, or leaflets, of lipids. The outer
surfaces of this bilayer are hydrophilic and exposed to water, whereas the interior of the
bilayer is shielded from water and forms the hydrophobic core of the membrane. The
main function of biological membranes is to act as permeability barriers, defining the
inside and outside of the cell and delimiting functionally different compartments. This
function alone, however, would not require the existence of the huge diversity of lipidic
constituents found in nature. If the only function of membranes were to act as
permeability barriers, it is reasonable to believe that just a few lipid species would be
Indeed, it has become evident that lipids perform a variety of functions in the
physiology of the cell, some of which will be briefly discussed below.
Lipids: chemical structures and classification
Two features common to all lipids are a polar portion, or head group, which is
exposed to the aqueous medium, and a non-polar portion, which is buried within the
interior of the bilayer.
The major classes of lipids are:
• Glycerophospholipids are the most abundant lipids, in which glycerol forms the
common backbone of the molecule, with one of its hydroxyls linked to a phosphate
via a phosphoester bond. The most common nomenclature
used to describe the chemical structure of these lipids is the
stereospecific numbering (sn) system.
If the glycerol is drawn in a Fisher projection with the middle
hydroxyl on the left, the three carbons are numbered as
shown in the figure, and indicated as sn-1 for C(1), etc.
Glycerophospholipids include several subclasses:
- 1,2-diacylphosphoglycerides or phospholipids, in which two of the glycerol
hydroxyls (sn-1 and sn-2) are linked via ester bonds to fatty acid chains and the
phosphate is at the sn-3 position. These are the most abundant lipids in
eukaryotic and prokaryotic cells, excluding Archaebacteria, in which the
phosphate is at the sn-1 position and the chains are linked to the glycerol via
ether rather than ester bonds.
- Lysophospholipids are phospholipids from which one of the acyl chains is
- Plasmalogens are phosphoglycerides in which one of the hydrocarbon chains is
linked to glycerol via a vinyl-ether bond. Plasmalogens with an ethanolamine
head group are abundant in myelin and in cardiac sarcoplasmic reticulum.
- Cardiolipids are formed by the linkage of two phospholipids via phosphoester
bonds to the two outer hydroxyls of a glycerol moiety, hence the alternative name
They are found in significant
amounts in the inner membrane
of mitochondria and in
chloroplasts, but are rare in
• Glycoglycerolipids are lipids in which the sn-3 position of glycerol is linked to a
carbohydrate via a glycosidic bond, rather than forming a phosphoester bond.
Monogalactosyldiacylglycerol has been named “the most abundant lipid in nature”,
since it constitutes half of the thylakoid membrane in plant chloroplasts (the R
groups in the figure represent the hydrocarbon chains of fatty acids). However, they
are rare in animals.
• Phosphosphingolipids, represented by sphingomyelin in the figure on p. 3, contain a
backbone formed by sphingosine, rather than by glycerol. The details of the core
structure of sphingolipids are illustrated in the diagram of the ganglioside GM1.
Sphingosine is an
amino alcohol in
which the sn-2 OH
is substituted by an
amino group and a
is attached via a
vinyl bond to the sn-1 carbon, leaving the sn-1 hydroxyl unreacted. A fatty acyl
chain is linked to the sn-2 amino group via an amide bond, forming ceramide. As in
glycerophospholipids, a phosphate bearing the head group is attached to the sn-3
position via a phosphoester bond. The amide and the hydroxyl group give these
lipids the ability to form intermolecular hydrogen bonds, which may be significant in
establishing interactions with proteins or in the formation of specialized membrane
regions, or microdomains.
• Glycosphingolipids are sphingolipids in which the sn-1 hydroxyl of ceramide is linked
to a carbohydrate via a glycosidic bond. The carbohydrates, which constitute the
head groups of these lipids, vary from a single sugar to very complex polymers.
Monogalactosyl ceramide is the most abundant component of the myelin sheath in
nerve. Gangliosides contain oligosaccharides with one or more molecules of anionic
sialic acid, while neutral oligosaccharides are contained in globosides.
Glycosphingolipids are usually minor components of the outer leaflet of plasma
membranes of animal cells, but they are significant in epithelial cells. The blood
group antigens consist of glycosphingolipids on the surface of erythrocytes.
• Sterols, represented by cholesterol in the figure above, are structurally separate
from the previous lipid classes. The membrane-embedded portion of the molecule
consists of a stiff ring structure and a short side chain, while the surface exposed
portion is limited to a single OH group. The stiffness of the ring structure leads to
changes in the dynamics and packing of the surrounding lipids. Cholesterol is found
in animal cells, where it can contribute as much as 30% of the mass of lipid
membranes, whereas plants contain other sterols.
In addition to the carbohydrates already mentioned, a variety of polar head
groups, differing in size, charge and chemical properties, are found in lipids. The
structures of the most common head groups are shown in the context of a typical
phospholipid, where the phosphate group is common to all head groups.
An unmodified phosphate represents the smallest head group, such as in phosphatidic
acid, which carries a double negative charge at neutral pH. Addition of choline or
ethanolamine, both positively charged at physiological pH, result in formation of the
neutral head groups of phosphatidylcholine (PC) and phosphatidylethanolamine (PE),
two of the most common lipids. Additional negatively charged head groups are formed
by modification of the phosphate with the zwitterionic amino acid serine, to give
phosphatidylserine (PS), or with the uncharged glycerol or inositol moieties, yielding
phosphatidylglycerol (PG) and phosphatidylinositol (PI), respectively. The individual
charges contributed by the various lipid head groups are the major determinant of the
overall electrostatic properties of biological membranes. Although only a minor
constituent of cell membranes, phosphatidylinositol is the precursor of many important
signal mediators generated by phosphorylation of several hydroxyls on the inositol by
An even greater variety exists in the types of hydrocarbon chains that are
attached to lipids. The table below lists some of the more abundant acyl chains,
including their common names and a useful condensed nomenclature.
The first number indicates the number of carbons, or methylene units, in the
chain, whereas the number on the right of the colon indicates its degree of unsaturation,
i.e. the number of C=C bonds in the chain. For unsaturated chains, the isomeric form of
the double bond, whether cis or trans, may also be given, as well as the number of the
first carbon at which the double bond is located, sometimes preceded by a greek letter
∆. Acyl chains with even numbers of carbons are more abundant than those with odd
numbers of carbons. The most common lengths are C16, C18 and C20, while the most
common unsaturated chains are 18:1, 18:2, 18:3 and 20:4. The double bonds are
usually in the cis configuration and in multiple unsaturated chains they are not
conjugated, i.e. they are separated by at least two C-C bonds. A large fraction of
phospholipids have one saturated and one unsaturated chain, the latter usually linked to
the sn-2 position of glycerol in animal cells.
Thus, the number of possible combinations of head groups and acyl chains gives
rise to a huge variety of lipid structures. As mentioned above, membranes within an
organism do not contain uniform mixtures of these various lipids. Not only are cells in
certain tissues enriched in specific types of lipids, but also different cellular organelles
contain membranes formed by unique mixtures of lipids. Mitochondria, for example, are
rich in cardiolipin and, indeed, the activity of cytochrome c oxidase, the component of
the respiratory electron transfer chain responsible for the final step of oxygen reduction,
is dependent on the presence of this lipid.
Such fine-tuning of lipid composition extends even further, as demonstrated in
the plasma membrane of human erythrocytes, in which the inner leaflet contains a
different lipid mixture than the outer leaflet.
Thus, with respect to neutral
phospholipids, the extracellular leaflet is
enriched in phosphatidylcholine and
sphingomyelin, both bearing a
phosphorylcholine head group, whereas
phosphatidylethanolamine is localized
preferentially in the inner leaflet. Most
striking, though, is the absolute exclusion from the outer leaflet of the anionic
phospholipid phosphatidylserine. Such asymmetric distribution of amino-phospholipids
is believed to be maintained by ATP-requiring enzymes, called translocases, which
catalyze the transbilayer transport of these lipids. The physiological function of the
surface expression of phosphatidylserine is related to the clearance of aged red cells.
After release into the bloodstream, a red cell remains in circulation for 120 days on
average. During this time, oxidative stress progressively deteriorates the biochemical
machinery of the cell, including the enzymatic activities responsible for the segregation
of this lipid. Eventually, phosphatidylserine appears on the outer surface of the
erythrocyte, and this signal is recognized by phagocytic cells within the spleen, which
clear the aged cell from circulation.
Platelets constitute another system in which the asymmetric phosphatidylserine
distribution is physiologically significant. In this case, the anionic lipid, which is normally
absent from the surface, becomes expressed on the outer membrane leaflet upon
activation of platelets at a site of injury. Together with calcium, phosphatidylserine is an
essential activator of blood coagulation factors.
Under physiological conditions, as well as in culture, cells may undergo
apoptosis, a controlled process of suicide. Thus, during development of the immune
system, self-recognizing thymocytes are eliminated in order to prevent autoimmune
reactions. One of the signatures of an apoptotic cell, in addition to DNA degradation, is
the expression of phosphatidylserine on the extracellular surface. This observation has
been used to develop an apoptosis test, which makes use of fluorescence-labeled
recombinant annexin, a normally cytoplasmic protein that binds specifically to
phosphatidylserine via calcium bridges.
Phospholipases and lipids as precursors of second messengers
Another essential function of lipids is as precursors of many second messengers
that participate in a variety of signaling pathways. Each of these molecules is
generated by cleavage of a lipid precursor at a specific bond. For this purpose, several
classes of enzymes, named phospholipases, exist.
Phospholipase A2 enzymes hydrolyze the ester bond
between glycerol and the sn-2 acyl chain of
phospholipids, thus generating a lysophospholipid
and a free fatty acid. Among the released fatty acids
is arachidonic acid, which is oxidized to other active
metabolites, such as prostaglandins, which are
involved in inflammation and other patho-
physiological processes. Analogous enzymes, called phospholipase A1, hydrolyze the
ester bond at the sn-1 position, but they are not yet well characterized.
Phospholipase C catalyzes the hydrolysis of the phosphoester bond, releasing a
soluble phosphorylated head group and a diacylglycerol. Of widespread significance is
the cleavage of phosphatidylinositol(4,5)-bisphosphate (PIP2), which generates two
second messengers: the water-soluble inositol(1,4,5) trisphosphate (IP3), which causes
the release of calcium from the endoplasmic reticulum by binding to the IP 3 receptor
found in the membrane of this organelle, and the membrane-bound 1-stearyl-2-
arachidonyl diacylglycerol, which is responsible for activation of protein kinase C and
enhanced phosphorylation of downstream signaling proteins.
Phospholipase D instead generates phosphatidic acid and a free head group.
Phosphatidic acid is becoming recognized as a second messenger, for example as an
activator of the NADPH oxidase responsible for the generation of reactive oxygen
products in neutrophils activated upon binding to immunoglobulin-coated bacteria.
Hydrophobicity and thermodynamics of lipid assembly
In this section, we will try to answer semi-quantitatively the following questions:
Which intermolecular forces are involved in the assembly of lipid membranes? What is
the thermodynamic basis for the formation and stability of the bilayer?
Three basic forces contribute to the stability of lipid aggregates:
The van der Waals force, which is a short-range electrostatic interaction
between instantaneous dipoles on adjacent molecules. This interaction develops as a
result of the formation of a dipolar charge in one molecule, due to fluctuations of its
electronic-nuclear distribution, and the instantaneous induction of a dipole of opposite
orientation on an adjacent molecule. This dipole-induced-dipole interaction stabilizes
the overall interaction between the two molecules and is proportional to the
intermolecular contact surface. These van der Waals interactions occur among all
types of molecules, between head groups at the water-bilayer interface as well as
between hydrocarbon chains in the interior of the membrane.
The electrostatic force is also responsible for the stronger ionic interaction
between charged groups and for the formation of hydrogen bonds between hydrogen-
bond donors, such as NH and OH, and acceptors, such as CO, on adjacent
sphingolipids. Hydrogen bonding is particularly extensive between water molecules
and is the origin of many of its solvent properties, such as its dielectric constant.
Attractive intermolecular ionic interactions form between the head groups of zwitterionic
lipids, such as phosphaditylcholine, whereby the negatively charged phosphate on one
head group interacts with the positively charged ammonium ion of the choline on a
nearby head group.
However, the greatest contribution to the stability of the membrane bilayer comes
from the hydrophobic force, or hydrophobic effect. Its physical origin can be
understood as follows. A hydrophobic molecule, such as a hydrocarbon chain, placed
in water must be surrounded by water molecules interacting with it. However, whereas
water-water interactions are stabilized by intermolecular hydrogen bonds, these
favorable polar interactions cannot be established between water and a molecule that
does not have hydrogen-bond donor or acceptor groups. Therefore, although bound
water molecules still maintain most of their hydrogen bonds with free water molecules,
some favorable electrostatic interactions are lost when water binds to the hydrocarbon
chain. This loss of hydrogen bonds represents an energetic, or enthalpic, penalty for
the system. However, the entropic cost of immobilizing water molecules on the surface
of the hydrocarbon chain is much more significant than this enthalpic penalty. The
water molecules bound to the hydrophobic surface lose the rotational and translational
degrees of freedom they had in pure liquid water. Because the free energy of the
system, ∆G, is given by
∆G = ∆H − T ∆S
where ∆H and ∆S represent the contributions of enthalpy and entropy, respectively, the
reduction in the ∆S term due to immobilization of water on the hydrophobic surface
leads to an increase in the ∆G of the system. For analogous reasons, placing water
molecules within the hydrophobic core of the membrane is also very unfavorable.
However, if separate hydrocarbon chains in water aggregate and pack tightly, the water
molecules immobilized on their surfaces can be released back into the bulk, thus
regaining their lost degrees of freedom. This leads to an overall increase in the ∆S term
and a reduction in the ∆G of the system.
Thus, the thermodynamic basis for the stability of the lipid bilayer lies in the
entropy difference between a system consisting of isolated hydrocarbon chains coated
with immobilized water molecules and a system in which aggregation of hydrocarbon
chains and formation of a hydrophobic bilayer phase lead to release of solvation water
back to the bulk aqueous phase. Additional stabilization arises from the enthalpic
contributions of van der Waals interactions between lipid chains as well as ionic and
H-bonding interactions between lipid head groups.
The hydrophobicity of a molecule can be determined quantitatively by measuring
its distribution, or partition, in a solvent system composed of water and an immiscible
hydrocarbon phase, such as hexane. The equilibrium constant, or partition coefficient,
K of the solute in this system is defined as:
[ X] H O
[ X] HC
where the [X] values represent the solute concentration in mole fraction units in water
and in the hydrocarbon phase, respectively. The partition coefficient K is related to the
standard state free energy of transfer, ∆G0trans, of the solute from water to the
∆G 0 = µ 0 2O − µ 0 = − RT ln K
trans H HC
where the µ0 values represent the standard chemical potentials of the solute in water
and in the hydrocarbon phase, respectively, R is the gas constant (1.987 cal °K -1 mol-1)
and T is the absolute temperature (°K).
It turns out that the hydrophobicity, as measured by ∆G0trans, is proportional to the
surface of contact between the hydrophobic solute and water, which determines the
number of water molecules that would be constrained. The proportionality constant for
transfer of an alkane chain from water into a hydrocarbon phase is found to be
∆G0trans ≈ −25 cal / Å2. Based on surface area, each additional methylene (CH2) group
contributes ~ −800 cal/mol to the hydrophobicity of the chain. At 25°C, this contribution
increases the partition coefficient K of the alkane chain by a factor of ~4 in favor of the
hydrocarbon solvent. The partition coefficient of water in hexadecane (C16) indicates
that the water concentration within the membrane is in the millimolar range,
corresponding to about 1 water molecule per 1,000 phospholipid.
Detergents and micelle formation
A short-chain alkane, at very low concentration, can be dissolved in pure water.
However, as more alkane is added, a concentration is reached at which a separate
phase forms, into which any additional alkane partitions. This critical concentration is
called the solubility limit.
Detergents and lipids, however, are amphiphilic molecules. For example, sodium
dodecyl sulfate (SDS), a detergent commonly used as a denaturant for protein gel
electrophoresis, has a 12-carbon long chain terminated by a charged sulfate group. Up
to a critical concentration of ≈ 1-2 mM (in 0.1M Na+), SDS dissolves in monomeric form.
Upon reaching the critical concentration, however, it forms a new phase composed of
Each SDS micelle contains ≈100 monomers,
spherical aggregates called micelles.
whose hydrophilic head groups delimit the water-exposed surface while their methylene
chains form the “oily” interior. Additional SDS increases the concentration of micelles,
while the concentration of monomers in solution remains constant. The critical
concentration in this case is called the critical micellar concentration (CMC). Since the
CMC is a measure of hydrophobicity, its value is a function of the chemical structure of
the amphiphile. For example, detergents with longer chains have lower CMCs, as
shown by decylmaltoside (10-C long, CMC=1.5mM) versus dodecylmaltoside (12-C
Biological phospholipids, with two long methylene chains, have extremely low
CMCs, below 10−10 M. For this reason, exchange of lipids between membranes is
facilitated by water-soluble cytosolic proteins, called phospholipid transfer proteins,
which bind lipid monomers and carry out the one-for-one exchange.
Lipid structure, lipid shape and lipid aggregate shape
The high-resolution structures of several lipids have been determined by x-ray
diffraction, using lipid crystal containing very little hydration water. A few examples are
given in the figure below.
A few noticeable features of these structures are:
• Within the crystals, the lipid are found in a lamellar arrangement, with the hydrophilic
and hydrophobic groups organized in stacked bilayers
• The acyl chains are fully extended, or in the all-trans configuration, with the
exception of the first 2 methylenes in the sn-2 chains of PC and PE, which are
oriented parallel to the crystallographic bilayer plane
• The tilt of the acyl chains away from the normal to the bilayer surface increases from
PE, in which they are virtually perpendicular, to PC , where the angle is ≈12°, to the
cerebroside, where the angle is 41°.
• The glycerol is oriented perpendicular to the bilayer plane, whereas the choline and
ethanolamine head groups are almost parallel to the plane. In fact, the amino group
of ethanolamine interacts with the unesterified oxygens of an adjacent molecule
A closer examination of the structural parameters of a PC molecule allows us to
understand the reason for the differences in packing and acyl chain orientations among
the various lipids.
The volume occupied by
the molecule can be divided
into two parts: a polar region,
which includes the head group
and the glycerol, and a
hydrophobic region, comprising
the two acyl chains, except the
first two carbons of the sn-2
chain. Each region also defines
an area given by its projection onto the plane of the membrane: the head group cross-
section area, S, and the acyl chain area, 2Σ, where Σ is the cross-section area of each
acyl chain. For saturated chains in the all-trans configuration, Σ=19Å2. The relationship
between these two areas determines the degree of tilt of the acyl chains in the crystals,
as well as the shape of the aggregates formed by the lipid in aqueous solution. If
2Σ < S, the chains will tilt to maximize the intermolecular van der Waals contacts and
accommodate the larger head group. This occurs in PC, where choline occupies an
area S≈50Å2. On the other hand, in PE S≈39Å2, so that 2Σ ≈ S and the chains can
attain optimal packing without tilting.
These qualitative considerations can be formulated in a more quantitative, albeit
empirical, formalism that allows us to predict the shape of the aggregate that each lipid
forms in solution. This formulation is based on the concept of the critical packing
parameter, which is defined as v / lSo, where v is the volume of the hydrocarbon
portion of the molecule, l is the maximum length of the acyl chain, and So is the optimal
surface area occupied by the molecule at the interface between the aggregate and
water. So is determined by the balance of repulsive and attractive interactions between
head groups, and is sensitive to solution conditions, such as ionic strength, divalent
cations and pH. The ratio v/l is analogous to the cross-sectional area of the
hydrocarbon portion (≈2Σ). The molecular shapes of amphiphilic molecules
corresponding to each value of the critical packing parameter are illustrated in the figure
below, together with the shape of the aggregate they form in solution.
Phospholipids, in general, have a good match between the areas of the two
molecular regions, and therefore they tend to naturally assemble in planar membranes,
forming a so-called bilayer phase. Under certain conditions, however, some of them
tend to form non-bilayer structures, consisting of tubular assemblies in which the head
groups face the interior lumen filled with water while the acyl chains, oriented outwards,
contact the chains from nearby cylinders. This macroscopic structure is called an
inverted hexagonal phase, HII, because the hydrophobic cylinders pack in a hexagonal
pattern. Lysophospholipids and most detergents, in which the single hydrocarbon chain
has a much smaller cross-sectional area than their head groups, form spherical
assemblies and give rise to a micellar phase. Both of these lipid phases are thought to
form locally at sites of membrane fusion.
X–ray diffraction structures of lipid crystals show the bilayers as planar and well-
ordered structures, with all-trans acyl chains. Similarly, lower resolution TEM images
convey the impression that biological membranes are flat slabs, in which the lipid head
groups form smooth, planar surfaces. However, fluid-phase lipid bilayers are very
different from these pictures. The ‘structure’ of fluid bilayers has been determined by x-
ray and neutron diffraction methods using multilamellar stacks of bilayers, which present
a periodic order in the direction perpendicular to the membrane plane. This one-
dimensional order allows the measurement of the distribution of matter along the bilayer
normal. Thus, the ‘structure’ of a fluid bilayer represents the time-averaged spatial
distribution of structural groups of
the lipid (carbonyls, phosphates,
double bonds, etc.) projected onto
the axis perpendicular to the
bilayer plane. These distributions
give the probability of finding a
particular structural group at a
specific location along the axis,
which represents the distance
from the center of the hydrophobic
core of the bilayer.
The structure of a fluid
bilayer of dioleoylphosphatidyl-
choline (DOPC, di-C18:1 cis-∆9) is shown in panel (b). The gaussian peaks give an
accurate representation of the true thermal motion of the molecules, which is a
fundamental property of fluid bilayers that plays a critical role in the interaction of
peptides and proteins with lipid membranes.
Several features of the fluid DOPC bilayer are important. First, the great amount
of thermal disorder is revealed by the widths of the probability density peaks, as
illustrated by the ≈10Å widths of the PO4 in the head groups and of the C=C acyl chain
group. The width of the C=C group is a vivid graphical representation of the effect of
the random configurational rearrangements occurring within the acyl chains, which
originate from thermally activated trans-gauche bond isomerizations. Second, the
overall thermal thickness of the interfacial region, defined by the distribution of the water
of hydration, is ≈30 Å, equal to that of the hydrocarbon core of the bilayer. As illustrated
by the end view diagram in panel (b) of the figure above, a peptide in an α-helical
conformation has a cross-sectional diameter of ~10 Å and can be easily accommodated
within the 15 Å thickness of the interface. Third, the physico-chemical environment in
the interface region is highly heterogeneous, presenting a steep but not abrupt change
from the solvation properties of water to those of the membrane hydrocarbon core.
Many membrane-supported reactions and protein-protein or lipid-protein interactions
occur in this heterogeneous environment. As illustrated in panel (a) of the figure, a
transmembrane α-helical peptide of twenty amino acids can be completely
accommodated within the hydrophobic core of the bilayer, whereas longer peptides will
have their ends protruding into the interfacial region. This imposes constraints on the
allowed amino acid sequences of the peptide, as discussed later.
These conclusions drawn from the two-dimensional representation of the fluid
bilayer structure are fully confirmed by the three-dimensional pictures of bilayers
obtained by computer simulations of the molecular dynamics of lipid bilayers. A
snapshot of the transversal cross section of the bilayer extracted from one of these
simulations is shown in the figure below.
The acyl chains, represented by the gray lines, are very disordered, each with
several kinks introduced by gauche bond configurations and only short all-trans
segments. The head group atoms, shown in red, are distributed over a wide range of
depths, and the surface delimiting the head-group region from the hydrocarbon region is
very rough. In agreement with the neutron diffraction data, water molecules,
represented by the blue oxygen atoms and white hydrogen atoms, are found deep in
the interfacial region, indeed as far as the boundary of the hydrophobic core.
Despite the disorder and the high degree of conformational fluctuations, the acyl
chains maintain tight packing and good van der Waals contacts, in agreement with the
minor increase in specific volume at Tm. Intrachain motions and translational diffusion of
individual molecules occur via thermally induced structural and packing fluctuations of
the surrounding molecules. This dense and crowded hydrophobic environment is the
origin of the high performance of thin biological membranes as permeability barriers, as
well as of the high cooperativity of lipid phase transitions.
Based on these experimental and theoretical results, the common illustrations,
which depict bilayers as two smooth surfaces separating polar and apolar regions, must
be considered misleading.
Membrane Proteins: Structure and Interactions
Prediction of protein topology
Identification of protein transmembrane domains – Sequence hydropathy
The difficulties of membrane protein crystallization and structure determination,
on the one hand, and the abundance of sequence information, on the other, have led to
efforts to develop theoretical methods for the prediction of the location of potential
transmembrane domains. Perhaps the most widely used of such methods is one based
on sequence hydropathy analysis.
The first step in such analysis is to evaluate the degree of hydrophilicity or
hydrophobicity of the protein along its amino acid sequence. Because of the
hydrophobic environment in the bilayer core, a membrane-spanning protein segment is
expected to contain a preponderance of apolar amino acids. The transmembrane
sequence should also be folded, either as an α helix or as a β strand in a β barrel, so
that peptide H-bonds are satisfied intramolecularly rather than by bringing bound water
into the hydrophobic bilayer core. Thus, the method attempts to find the location and
number of transmembrane segments based on the relative hydrophilic and hydrophobic
properties of contiguous stretches of the amino acid sequence.
For this purpose, hydropathy scales have been devised to rank the relative
hydrophilicity and hydrophobicity of the 20 amino acids. One of these scales, proposed
by Kyte and Doolittle in 1982 (J. Mol. Biol. 157: 105-132), is still in widespread use.
This scale was defined using the water-vapor partition coefficient or standard
free energy of transfer, ∆G0transfer, of the amino acid side chains as well as their degree of
surface exposure in proteins of known crystallographic structure. The hydropathy
indexes are normalized between 4.5 and −4.5, with positive values indicating that free
energy is required to transfer the side chain to water and the amino acid is considered
hydrophobic. Conversely, negative values indicate that free energy is released upon
transferring the side chain into water and the amino acid is hydrophilic.
Given an amino acid sequence, the residue letter code is substituted with its
corresponding hydropathy index to give the sequence hydropathy profile. A window of
odd length, usually 7–13 residues, is scanned along the sequence. The hydropathy
index values within the window are summed, and the average hydropathy value of the
segment is computed by dividing the sum by the size of the window, as shown below.
E I T W I V G M V I Y L L M M G A
i= 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17
|−−−−−− sliding window −−−−−−|
Hi= -3.5 4.5 -0.7 -0.9 4.5 4.2 -0.4 1.9 4.2 4.5 -1.3 3.8 3.8 1.9 1.9 -0.4 1.8
Using a window size w = 7, the first value of the sequence hydropathy profile, Dl,
corresponds to position 4. The general recursive formula for the computation of Dl is
i =l + k
k = ( w − 1) 2
Dl = i =l − k
D4 = (-3.5+4.5-0.7-0.9+4.5+4.2-0.4) / 7 = 1.1
D5 = (4.5-0.7-0.9+4.5+4.2-0.4+1.9) / 7 = 1.87
Finally, a threshold value for the segmental hydropathy must be selected to
decide when a stretch of amino acids may be considered hydrophobic enough to be a
potential transmembrane protein sequence. For the Kyte-Doolittle scale, this threshold
value is usually taken to be 1.0-1.25.
Since the thickness of the bilayer hydrophobic core is ~30Å and each amino acid
contributes 1.5Å to the length of the α helix, a potential transmembrane segment in this
conformation is expected to be at least 20-residue long. Thus, a putative α-helical
transmembrane segment should be identifiable by a consecutive stretch of ~20 values
of Dl above the hydropathy threshold value.
As an example, the figure below shows the plot of the hydropathy profile of
glycophorin A, as it appeared in the original Kyte and Doolittle paper.
A sliding window of 7 residues was used to compute each value of hydropathy
index. However, the authors reported the sum of the indexes for the residues within the
sliding window rather than the average value, as it is now common practice (i.e. the
hydropathy index in the figure is 7×Dl). The single α-helical transmembrane segment is
easily identified in the region of residues 73-95.
The “positive inside” rule
Once the putative transmembrane segments have been identified, their
orientation in the membrane must be chosen. Statistical analysis of single- and multi-
spanning proteins indicated that their topological determinants may reside in the polar
and loop regions flanking the transmembrane segments. In particular, the distribution of
positively charged residues, arginine and lysine, was well correlated with the topology.
In a sample of bacterial inner membrane proteins, the frequency of Arg + Lys residues
was 4-fold higher in the cytoplasmic than in the periplasmic membrane flanking regions.
A similar bias, though not as strong, has been found to apply to proteins from higher
organisms as well.
Thus, the “positive inside” rule, first proposed by G. von Heijne, is thought to be a
fundamental determinant of the topology of most integral membrane proteins. Clusters
of positively charged amino acids found near one end of a predicted transmembrane α
helix, identify it as the cytoplasmic end.
Intrinsic proteins are anchored to membranes of eukaryotic cell not only via
transmembrane domains, but also via covalently attached hydrocarbon chains. Four
classes of lipid-anchored proteins are distinguished by the type of chain and linkage.
1. Glycosylphosphatidylinositol (GPI)-anchored proteins are a heterogeneous
family of proteins found on the exoplasmic surface of cell membranes. The GPI
anchor is formed by a phosphatidylinositol linked, by N-acetylglucosamine, to a
polymannose chain and a phosphoethanolamine. The linkage between protein and
anchor is always located at the carboxyl-terminal amino acid. The GPI anchor is
always added post-translationally after proteolytic removal of a peptide, 17-31
residues long, from the C-terminus of a protein precursor. GPI-anchored proteins
include hydrolytic enzymes, (alkaline phosphatase, acetylcholine esterase, 5’-
nucleotidase), adhesion proteins (neural cell adhesion molecule or N-CAM),
receptors (FcγRIIIB, a neutrophil receptor for Fc region of immunoglobulin G).
GPI-anchored proteins can be released from the surface by treatment with
exogenous phosphatidylinositol-specific phospholipase C.
2. N-Myristoylated proteins are found anchored to the cytoplasmic face of the plasma
membrane and to the membranes of other organelles. The 14-carbon chain is
attached co-translationally via a stable amide linkage to an N-terminal glycine
residue. Examples of N-myristoylated proteins are the protein tyrosine kinase p60src
and the catalytic subunit of cAMP-dependent protein kinase, also known as PKA.
3. S-prenylated proteins are anchored to the cytoplasmic plasma membrane surface
by a 15-carbon farnesyl or a 20-carbon geranylgeranyl unsaturated chain. The thio-
ether linkage of the anchor to a cysteine residue in the protein is catalyzed by a
farnesyl-transferase. The cysteine is initially the fourth residue from the C-terminus
of the protein. However, after the chain is attached, the 3 C-terminal residues are
cleaved by a protease and the carboxyl group of the cysteine is methylated.
4. S-palmitoylated proteins are anchored to the plasma membrane by a 16-carbon
saturated palmitic chain via a thio-ester linkage. This modification may occur by
spontaneous reaction of palmitoyl-coenzyme A with a protein already associated
with the membrane or may be catalyzed by a membrane-associated palmitoyl-
transferase. S-palmitoylation can take place on exposed cysteines anywhere in the
protein. Contrary to thio-ether bonds, a thio-ester linkage is labile and is hydrolyzed
under mild basic conditions. Proteins known to be palmitoylated include p21ras, the
glycoprotein hemagglutinin of influenza virus, the mammalian transferrin receptor
and the visual receptor rhodopsin, which is a seven-helix transmembrane protein. In
many cases, doubly acylated proteins are generated first by co-translational
N-myristoylation and, after interacting with the membrane, by S-palmitoylation.
The protein affinity for the membrane conferred by these lipid anchors has been
estimated by studies of the equilibrium binding of lipid-modified fluorescent peptides.
The results are summarized in the table below.
On the right column, the values of the effective dissociation constant, Kdeff, are
listed. The binding energy of fatty acids and acylated peptides to phospholipid bilayers
increases linearly with the number of carbons in the chain. The slope of 0.8 kcal mol-1
per -CH2 group is equal to that found for the partitioning of the neutral form of a fatty
acid from water into a bulk alkane phase. The values of Kdeff for myristoyl and palmitoyl
anchors reflect the increase in ∆G0transfer upon addition of two methylenes. Thus, the
binding energy of acylated peptides and proteins to membranes is due to the classical
hydrophobic effect. The membrane affinities of whole proteins are estimated to be
about 10-fold lower than those for short peptides, in other words their Kdeff values are
about tenfold higher. Thus, a myristoylated protein is expected to have a Kdeff ≈0.8mM
while the concentration of lipids in a cell is approximately millimolar. Since the lipid
concentration is comparable to Kdeff, simple myristoylation provides barely enough
hydrophobic energy to attach a protein to a phospholipid bilayer. Other factors, such as
electrostatic interactions between amino acid side chains and phospholipid head
groups, may help in partitioning these proteins to the membrane. These relatively weak
membrane-protein affinities create the potential for dynamic and reversible membrane
localization of these proteins, which may be targets of metabolic modifications leading
to changes in their binding affinity and effective concentration at the membrane.
Modulation of reversible protein-membrane interactions: The myristoyl-
An example of an electrostatic charge-induced reversible membrane association
is given by the myristoylated protein MARCKS, an acronym for myristoyl alanine-rich C-
kinase substrate. MARCKS binds to Ca2+-calmodulin and actin and is thought to
integrate protein kinase C (PKC) and Ca2+-calmodulin signals that affect interactions of
actin with the cytoskeleton and membranes. Binding of calmodulin requires Ca2+ and is
prevented by PKC phosphorylation of MARCKS, whereas binding and crosslinking of
actin filaments by MARCKS is blocked by phosphorylation and by Ca2+-calmodulin.
MARCKS is a rod-shaped protein with at least two domains: an N-terminal
myristoylation domain and a basic effector domain that contains the PKC
phosphorylation sites as well as the calmodulin and actin-binding sites.
Three peptide fragments, corresponding to the basic domain of MARCKS, were
used to characterize the effect of changes in their electrostatic properties on membrane
MARCKS 151-175: KKKKKRFSKKSFKLSGFSFKKNKK
Tetra-Asp MARCKS: KKKKKRFDKKDFKLDGFDFKKNKK
The MARCKS peptide is strongly cationic, because of all the lysine residues,
whereas the positive charges in the Asp-substituted and the tetra-phosphorylated
peptides are progressively neutralized by the added negative groups. Binding of the
peptides was measured as a function of the fraction of phosphatidylserine in the bilayer.
As shown in the figure below, addition of negative charges to the peptide diminishes its
affinity for the membrane. Thus, a higher density of anionic head groups is required to
reach 50% binding to the bilayer. Phosphorylation is particularly effective.
Thus, in the presence of bilayers containing 10-20% acidic lipids, MARCKS
phosphorylation leads to almost complete desorption of the peptide from the membrane.
This effect was demonstrated in a kinetic experiment by using mixed bilayers
containing phosphatidylserine and
fluorescence-labeled tracer lipids. Under the
experimental conditions, the fluorescence
intensity was proportional to the degree of
peptide binding to the membrane. Addition of
MARCKS 151-175 peptide produced an
increase in the fluorescence of the labeled
lipids. Upon addition of PKC in the presence
of ATP, serine phosphorylation caused the
immediate desorption of the peptide from the
membrane, which was completed after ~1 min, as judged by the decrease of the
fluorescence signal back to the level measured in the absence of peptide.
The binding of full-length MARCKS to membranes requires the contribution of
both the hydrophobic interactions of the myristoyl anchor with the bilayer core and the
electrostatic interactions between the basic domain and anionic lipid head groups.
Phosphorylation by PKC reduces the electrostatic binding energy and leads to
desorption of MARCKS, until dephosphorylation by protein phosphatases restores the
full interaction energy. Thus, the myristoyl anchor facilitates the initial transfer of the
protein to the membrane, which then increases the chance that the basic domain will
associate electrostatically with the anionic lipids. This mechanism for reversible protein-
membrane binding is known as the myristoyl-electrostatic switch.
Membrane anchoring of peripheral proteins via non-covalently bound lipids
Protein targeting to specific cellular compartments in response to external stimuli
is a fundamental component of signal transduction mechanisms in eukaryotic cells.
Such localization can be achieved by means of protein-protein interaction domains,
such as src-homology 2 (SH2) and src-homology 3 (SH3) domains, which recognize
specific phosphotyrosine motifs and proline-rich sequences, respectively, in the protein
binding partners (e.g. activated receptors).
Alternatively, localization can be carried out by protein domains that bind
specifically to lipids embedded in cell membranes. The best known members of this
class of domains are the protein kinase C (PKC) homology-1 (C1) and PKC homology–
2 (C2) domains, the FYVE domain, and the pleckstrin homology (PH) domain. Some
C1, C2, and PH domains interact with proteins in addition to or instead of lipids.
Fluorescence microscopy and fusion proteins derived from green fluorescent
protein (GFP) and several of these domains have allowed detailed studies of the
kinetics of spatial redistribution (e.g. from cytosol to membranes or vice versa) during
A model of the membrane-docked structure of a representative domain of each
type is shown in the figure below.
Panel A: structure of the complex between the C1 domain of PKCδ and phorbol ester is
shown with the model of a myristoyl chain. Panel B: structure of the Ca2+-bound C2
domain of cPLA2 interacting with the membrane. Panel C: structure of the FYVE
domain of Vps27p with a model of phosphatidylinositol 3-phosphate (PI3P). Panel D:
structure of the PH domain of PLCδ1 complexed with Ins(1,4,5)P3 and with a model of
the two myristoyl chains. The secondary structure and molecular surface of each
domain are shown. The surface colors indicate the nature of the residues, green for
hydrophobic and blue for basic. Some specific and nonspecific contact residues are
also indicated. The two Zn2+ in the C1 and FYVE domains are shown as cyan circles,
while the two Ca2+ in the C2 domain are shown as blue circles. The domains are
positioned so that known membrane-interacting residues penetrate the membrane and
patches of basic residues are near the membrane surface. The membrane leaflet,
drawn to scale, is divided into an interfacial zone and a hydrophobic core, each ~15 Å
Originally discovered as a conserved region responsible for the activation of
PKCs by diacylglycerol or phorbol esters, C1 domains have been found in >200 other
proteins. The C1 domain is a compact motif of ~50 amino acid residues, containing two
small β sheets and short C-terminal α helix that are built around two 3-Cys-1-His Zn2+-
binding clusters, with the two ions integral to the overall structure. One entire end of the
C1 domain surrounding the diacylglycerol-binding groove is very hydrophobic.
Membrane binding of C1 domains occurs by strong synergism between the
stereospecific interaction of diacylglycerol with its binding site and the nonspecific
hydrophobic interaction between the membrane and the C1 domain surface surrounding
the binding site.
In most PKCs, C1 domains occur in pairs. C1 domains from PKCγ have been
observed to translocate from the cytosol to the plasma membrane within a few seconds
after addition of diacylglycerol. In the inactive cytosolic form of PKCγ, the diacylglycerol-
binding sites are obstructed. Opening of these sites may require binding of the enzyme
to the membrane via Ca2+-mediated C2 domain-phospholipids interactions.
Diacylglycerol binding to the C1 domain is also believed to lead to allosteric activation of
the enzyme by a conformational change that alters the interactions of the C1 domains
with the catalytic domain of the enzyme.
C2 domains consist of ~120 residues folded in a β sandwich structure related to
that of immunoglobulin. Originally discovered as a conserved motif in Ca2+-dependent
PKCs, ~600 C2 domains have now been found in >400 proteins involved not only in
signal transduction, but also in inflammation, synaptic vesicle trafficking and fusion, and
many other processes. The properties of C2 domains vary, with some of them binding
to phospholipid membranes in a Ca2+-dependent manner, while others bind
constitutively. Other C2 domains exhibit both Ca2+-dependent and -independent binding
to proteins rather that membranes. The structures of C2 domains from synaptotagmin,
PKC-β and -δ, and phospholipases A2 (cPLA2) and C-δ1 (PLCδ1) have been
determined. The Ca2+-binding sites are formed by three loops at one tip of the structure.
Most Ca2+-dependent C2 domains bind acidic phospholipids, but that of cPLA2 seems to
prefer neutral lipids, especially phosphatidylcholine (PC).
The subcellular localization of C2 domains correlates with their phospholipid
specificity. Thus, when the free Ca2+ concentration increases in response to a stimulus,
C2 domains from PKCα and PKCγ translocate to the plasma membrane, rich in the
acidic phosphatidylserine lipid, whereas cPLA2 translocates to the PC-rich nuclear
envelope and endoplasmic reticulum.
FYVE domains have been found in ~60 proteins and consist of 70-80 residues
containing 8 Cys or 7 Cys and 1 His that coordinate two Zn2+. FYVE domains are
specific for phosphatidylinositol-3-phosphate (PI3P), whose concentration in the cell
increases following activation of phosphatidylinositol 3-kinases (PI3-kinase). As
illustrated in the figure above, FYVE domains bind to PIP3-containing membranes so
that the tip of the N-terminal loop, which contains hydrophobic residues, penetrates into
Proteins containing FYVE domains localize to endosomal membranes containing
PI3P, and this localization is blocked by inhibition of PI3-kinase.
Found in >500 proteins, PH domains bind various phosphorylated phosphatidyl-
inositols (phosphoinositides) with different affinities and thus respond sensitively to the
activities of phosphatidylinositol kinases, phosphatases and phospholipases. Of
particular interest, signaling through PI3-kinases depends on PH domain-containing
effectors, in addition to those containing FYVE domains. Structures are known for PH
domain of several proteins, among which spectrin, PLCδ1, β-adrenergic receptor kinase
(βARK) and insulin receptor substrate 1 (IRS-1). The PH domain structure contains two
orthogonal antiparallel β sheets of three and four β strands, followed by a C-terminal α
helix. The β sheets fold into a barrel-like structure, one end of which is capped by the
α helix. The loops connecting the β strands are involved in ligand binding and vary
substantially in sequence and structure between PH domains.
Based on the binding to different phosphoinosite polyphosphates and inositol
polyphosphates, PH domains have been classified into four groups.
Group1 contains proteins such as Bruton’s tyrosine kinase (Btk), whose PH
domains bind phosphatidylinositol 3,4,5-trisphosphate, PI(3,4,5)P3 with high specificity.
Group 2 includes proteins such as PLCδ1 and βARK, whose PH domains have
high affinity for PI(4,5)P2 and PI(3,4,5)P3 in vitro. In vivo, preferential binding to
PI(4,5)P2 may occur as a consequence of the higher abundance of this lipid rather than
discrimination against the 3-phosphorylated PI.
Group 3 includes proteins such as Akt, also known as protein kinase B (PKB),
whose PH domains bind preferentially PI(3,4)P2 and PI(3,4,5)P3.
Group 4 is a heterogeneous group that includes proteins with relatively low
affinity for all ligands mentioned above. The PH domain of PLC-γ binds
3-phosphoinositides, including PI3P, while the PH domains of PLCβ1 and PLCβ2 bind
nonspecifically and with low affinity to neutral and acidic phospholipids.
In addition to binding to membrane-bound phosphoinositides, PH domains
display variable affinity for soluble inositol phosphates. For examples, the PH domain of
PLCδ1 binds to PI(4,5)P2 in vesicles with micromolar affinity and to the soluble
Ins(1,4,5)P3 with Kd = 210 nM. The higher affinity for the latter may be important in
product inhibition of the enzyme. Stimulation of PLC causes repartitioning of a fusion
protein consisting of green fluorescent protein and PLCδ1 PH domain from the plasma
membrane to the cytosol concomitant with the hydrolysis of PI(4,5)P2 in the membrane
and formation of soluble Ins(1,4,5)P3. PH domains that bind 3-phosphorylated
phosphoinositides, including those of Btk and Akt, have similarly been observed to
translocate to the plasma membrane upon activation of PI3-kinases.
Many of the interactions described for these membrane-targeting domains are of
relatively low affinity and thus their physiological importance may be questioned.
However, many important interactions appear to be weak “by design”, so that
membrane binding of certain proteins may require the simultaneous presence of more
than one ligand, the coincident activation of more than one signaling pathway. This is
exemplified by PKCγ, which contains both diacylglycerol-binding C1 and
Ca2+-phospholipid-binding C2 domains, and requires both the production of
diacylglycerol and an increase in the concentration of free Ca2+ for full activation.
Intracellular Ligand-Gated Channels
Regulation of ion channel activity can modulate many physiological processes, such as
electrical excitability, secretion, and salt transport across epithelia. Most channel proteins are
post translationally modified (e.g. through phosphorylation or through interactions with
intracellular signaling molecules) but certain channels depend on such interactions in order to
be gated open or closed. Here we will consider three examples of intracellular ligand-gated
channels: a K channels that is inhibited by the ligand ATP, a K channel that is activated by G
proteins and a non-selective cationic channel that is activated by cyclic nucleotides.
ATP-sensitive K channels:
This inwardly rectifying K channel is inhibited by cytosolic adenosine triphosphate
(ATP), thus coupling the metabolic state of the cell to membrane electrical events.
These channels are found in all types of muscle cells (skeletal, cardiac and smooth), in
neurons, in renal tubular cells and in the β cells of the pancreas, where their physiologic
role is best understood. In pancreatic β cells they regulate insulin secretion in response
to glucose. These channels are normally active at rest and β cells are thus kept at
negative resting potentials. After a meal, when glucose is metabolized and ATP is
produced (actually the channel senses the increased ATP/ADP ratio) the channel is
inhibited the cell depolarizes, fires action potentials, allowing entry of Ca and secretion
of insulin. These channels are associated with a member of the larger ATP-binding
cassette proteins, called the sulphonylurea receptor or SUR (other members of these
transmembrane proteins include: P-glycoprotein or multidrug resistance protein, the
cystic fibrosis transmembrane regulator or CFTR, etc.- see lecture 12) (Fig. 1).
Figure 1. The inward rectifier Kir6.2
combines with the sulfonylurea
receptor (SUR) to generate ATP-
sensitive K currents.
The association of the SUR and the K channel produces a functional channel that is
blocked by sulphonylureas, drugs that constitute the principal treatment for adult onset
diabetes. Similarly, the SUR association causes the K channel to be activated by SUR-
binding drugs called potassium “channel openers” (e.g. diazoxide, pinacidil), and by
nucleoside diphosphates (e.g. ADP) that bind SURs at the nucleotide binding folds
(NBFs – Fig. 1). It has been established that there is an inverse relationship between
ATP sensitivity and PIP2 levels and that the channel activity depends on the presence of
PIP2. However, the details of the mechanism by which ATP inhibition or SUR-channel
interactions relate to channel gating by PIP2 are unclear but are the subject of an active
area of research. Moreover, the physiological importance of regulation of PIP2 levels on
the activity of this K channel is not clear. It is long known for example that at glucose
concentrations below the threshold for stimulation of insulin secretion and electrical
activity, muscarinic stimulation that leads to PLC activation initiates electrical activity
and insulin release. Whether, this effect is mediated by reduction of PIP2 levels and
enhanced sensitivity to ATP inhibition of the K channels remains to be shown.
G protein-gated K channels:
Using the cell-attached mode of the patch clamp technique it can be shown that extracellular
application of ACh is effective in stimulating channel activity in an atrial patch, when perfused
through the pipette but not through the bath (Fig 2). In their isolated patch, external signaling
was limited to ACh in the patch pipette (since the membrane patch seems to be physically
isolated at the sites of the gigaseal between the glass electrode and the plasma membrane from
substances in the bath and from membrane molecules other than those within the patch)
whereas internally, soluble second messengers (e.g. GTP) do have access to the cytoplasmic
surface of the patch. This result has been used as evidence for the membrane delimited nature
of the action of ACh. Using the whole-cell mode of the patch-clamp technique, it has been
shown that the ACh effect proceeded via a pertussis toxin (PTX) sensitive G protein and that
non-hydrolyzable GTP analogs could bypass signaling through the receptor and cause
persistent stimulation of channel activity. Experiments with inside-out patches provided further
evidence for the membrane delimited nature of the ACh signaling and the involvement of G
proteins. Perfusion of inside-out patches with purified Gβγ subunits caused persistent stimulation
Figure 2. Unitary inward K currents measured on a rabbit atrial cell with an on-cell patch pipette containing isotonic KCl. The control trace is
before ACh additions. The second trace is after perfusion of 100 nM ACh in the bath, and the third trace is after washing ACh out of the bath
and perfusing 10 nM ACh into the pipette. Em = -90 mV.
of K current activity in a Mg-independent manner. The Gβγ activation of this K current provided
the first example of the effector function of the Gβγ complex in any system. It has been shown
that Gβγ binds directly to both the carboxy- and amino- cytoplasmic segments of the channel
protein. It is thought that Gβγ binding stabilizes channel-PIP2 interactions that serve to open the
channel gate. The details of this mechanism are an active area of research. The atrial (or
nodal) K+ channel that is activated by acetylcholine through muscarinic m2 type receptors has
served as the prototypical G protein-gated K channel. Pertussis toxin-sensitive,
neurotransmitter-activated inwardly rectifying K+ currents have also been reported in the central
nervous system and in other peripheral tissues such as the pancreas and the pituitary. Direct
activation of potassium (K+) channels by G proteins is involved in the rapid inhibition of
membrane excitability, such as in the slowing of heart rate by the vagus nerve or the
autoinhibitory release of dopamine by midbrain neurons. Thus these K channels couple G
protein-coupled receptor signaling to membrane excitability.
Figure 3. In the hypothesis of
drawn here for muscarinic
modulation of a K(ACh)
channel, only three
macromolecules are used in the
signaling cascade: receptor (M),
G protein (Gk), and channel.
They remain in the membrane
throughout. The activated G
protein (G*) interacts directly
Figure 4 shows the crystal structure of the cytoplasmic domains of one of the G protein
sensitive channels GIRK1. On this structure we have mapped mutations that affect
Figure 4. Crystal structure of
the cytosolic domains of GIRK1.
Mutation sites have been
mapped onto the structure
revealing that PIP2 and Gβγ
interacting sites of the channel
are in close proximity to the lipid
(Nishida and MacKinnon, 12/27
2002, Cell 111:958-965)
channel-Gβγ and channel-PIP2 interactions. Most of these mutations seem to come
together to a region that is in close proxility to the lipid bilayer.
Cyclic nucleotide-gated channels involved in phototransduction:
Cyclic nucleotide-gated channels (CNG) are composed of two subunits in a tetrameric
arrangement, two α subunits and two β subunits. The α subunits can produce
functional channels when expressed in heterologous expression systems. The β
subunits do not express by themselves, but when co-expressed with their
corresponding α subunit, they produce channels with altered permeation,
pharmacology, and/or cyclic nucleotide selectivity. The primary structure of each CNG
channel subunit is a six-membrane spanning segment protein resembling that of
voltage-gated K channels (we will examine those in the next lecture). CNG channels
are found in all sensory organs and possess a cyclic nucleotide-binding domain. This is
a highly conserved stretch of approximately 120 amino acids that is homologous to
similar domains of other proteins, including the cAMP- and cGMP-dependent protein
kinases and the catabolite-activating protein (CAP), a bacterial transcription factor.
Although the retinal and olfactory CNG channels exhibit a high degree of sequence
similarity (over 80% amino acid identity) in the putative binding region, the native
channels exhibit different cyclic nucleotide selectivities. For the retinal channel, cGMP
is a much more potent and effective agonist than cAMP. For the native olfactory
channel cAMP and cGMP have very similar effects.
Let us consider the role of retinal CNG channels in phototransduction. In the
Figure 5. The retina has five major classes of
neurons arranged into three nublear layers:
photoreceptors (rods and cones), bipolar cells,
horizontal cells, amacrine cells, and ganglion
cells. Photoreceptors, bipolar, and horizontal
cells make synaptic connections with each
other in the outer plexiform layer. The bipolar,
amacrine, and ganglion cells make contact in
the inner plexiform layer. Bipolar cells bridge
the two layers. Information flows vertically from
photoreceptors to bipolar cells to ganglion cells.
Information also flows laterally, mediated by
horizontal cells in the outer plexiform layer and
amacrine cells in the inner plexiform layer.
retina of vertebrates phototransduction is accomplished by sensory cells (the rods and
the cones), connected by interneurons (bipolar, horizontal and amacrine cells) to
ganglion cells that transmit signals to the optic nerve and the brain (Fig. 5). Rods and
cones have different sensitivities and respond to different frequencies of light. The
cones cells can further be subdivided into cells that preferentially sense different colors.
However, it is rods, from a variety of species, that have been the favored cell type for
studying visual transduction. Figure 6a shows the structure of a rod photoreceptor.
Figure 6a. A photoreceptor. The
drawing (left) is of an entire rod
photoreceptor. The micrograph
Figure 6b. The dark current (left). In the dark,
(right) shows only the outer segment current flows through sodium channels in the
of a salamander cone. outer segment of a rod photoreceptor. A pulse of
light (right) closes these channels, resulting in
hyperpolarization of the rod.
The cell has two parts. The rod outer segment is elongated and contains a stack
of flattened disks made from internal membranes. This is connected by a thin bridge to
the remainder of the cell, the inner segment that contains the nucleus, the mitochondria,
and the presynaptic terminal that synapses onto other neurons in the retina. It is the
outer segment that is the business end for visual transduction. Within the internal
membranous disks is found the light-sensitive protein rhodopsin. This is made up of an
opsin protein, bound to a light-sensitive molecule or chromophore termed retinal. The
later molecule may exist in a number of different forms, of which 11-cis-retinal and all-
trans-retinal are the two major isomers. On its own, neither opsin nor retinal absorbs
visible light. In combination, however, absorption of a photon of light causes and
isomerization of retinal from the 11-cis form to the all-trans form. The light-dependent
isomerization of retinal then causes a structural rearrangement of the protein.
Rhodopsin that has been activated in this way is termed meta-rhodopsin. For all of the
subsequent steps in visual transduction, it is useful to think of this molecule as
analogous to a receptor that has just bound its neurotransmitter. In fact, the structure of
the opsin protein is similar to G-protein coupled receptors. Not surprisingly, therefore,
the steps that follow the production of meta-rhodopsin involve the production of a
second messenger through the action of a G-protein. This G-protein is called
transducin. When meta-rhodopsin binds to transducin, GDP is replaced by GTP, and
the αT-subunit of transducin is liberated from its complex with the βγ subunit. The target
of the newly liberated αT is an enzyme in the membranous disks, a phosphodiesterase,
that cleaves the second messenger cGMP to 5 ' GMP. Even in the dark, the levels of
cGMP in the outer segments are maintained by a balance between its rate of synthesis
through guanylate cyclase and degradation by the phosphodiesterase. The action of
αT, formed after exposure to light, is to stimulate the phosphodiesterase, producing a
drop in the levels of cGMP. This drop occurs within about 100 ms of the onset of a light
flash, sufficiently fast to account for a visual response. In many respects,
photoreceptors are built backwards. When excited by light, they respond by dropping,
rather than raising, their concentration of the second messenger cGMP.
The dominant type of ion channel in the plasma membrane of the outer
segments is the CNG channel that allows sodium and calcium to enter the cell.
Because of the abundance of sodium ions in the extracellular fluid, the major ion that
enters the outer segments through these channels is sodium. We would expect a cell
with a preponderance of such CNG channels to have a very positive resting potential.
The effect of the rod CNG channels is, however, counterbalanced by potassium
channels. The interesting thing about these potassium channels is that they are found
in a very different part of the cell, the membrane of the inner segment that includes the
nucleus and synaptic terminal. Because there is good electrical continuity between the
inner and outer segments, the mean membrane potential is kept fairly negative as a
result of the open potassium channels. This spatial distribution of channels, however,
creates a circulating current, termed the dark current (Fig 6b), which flows in through
the outer segment CNG channels, through the bridge into the inner segment and out
through the potassium channels. The effect of shining light on a rod is to shut down
many of the CNG channels in the outer segment. This produces a marked decrease in
the dark current. As a result the potassium channels, which remain open in the inner
segment, hyperpolarize the cell toward EK, reducing the spontaneous release of
neurotransmitter from the synaptic terminal. The closure of the CNG channels can be
attributed directly to the drop in cGMP in the cytoplasm of the outer segment. The CNG
channels normally bind cGMP directly, and remain open only when cGMP is bound.
This can be demonstrated by making inside-out patch recording on membrane from the
outer segments. When cGMP is added to the cytoplasmic face of the patch a large
increase in conductance, attributable to the opening of the CNG channels can be
measured. One interesting feature of the CNG channels is that, under normal
conditions, the conductance of a single channel is extremely low. The reason for this is
that calcium and magnesium ions, which are normally present in physiological solutions,
partially block these channels. This block is relieved when calcium and magnesium are
omitted from the solutions, and individual openings of the channel are much larger.
This cascade of reactions that follows the formation of meta-rhodopsin produces
a very significant amplification of the signal generated by light. It has been estimated
that a single molecule of meta-rhodopsin, which is formed by the action of a single
photon of light, diffuses in the membrane and activates several hundred transducin
molecules before it is rendered inactive. The subsequent stimulation of the
phosphodiesterase by αT provides further amplification such that a single photon of light
can lead to the destruction of more than 100,000 molecules of cGMP.
The analogies between visual transduction and neurotransmitter action can be
taken further, when one considers how the response to a flash of light is terminated. A
protein called rhodopsin kinase phosphorylates meta-rhodopsin making it relatively
ineffective at activating transducin, and thus terminating the light response. After
phosphorylation, the all-trans-retinal dissociates from rhodopsin, leaving the opsin
protein, which must bind another 11-cis-retinal before it can again be activated by light.
Similarly, the β-adrenergic receptor (stimulated by isoproterenol or norepinephrine) is
phosphorylated by the β-adrenergic receptor kinase (βARK) to make it ineffective in
The diversity of neurotransmitters is extensive, but their receptors can be grouped into two
broad classes: ligand-gated ion channels and G protein-coupled receptors. In this section, we
describe two important receptors that are also ligand-gated ion channels. By far the most-
studied receptor is the muscle nicotinic acetylcholine receptor, the first ligand-gated ion channel
to be purified, cloned, and characterized at the molecular level. The structure and mechanism
of this receptor are understood in considerable detail, and it provides a paradigm for other
neurotransmitter-gated ion channels. When activated, these receptors induce rapid changes,
within a few milliseconds, in the permeability and potential of the postsynaptic membrane. In
contrast, the postsynaptic responses triggered by activation of G protein-coupled receptors
occur much more slowly, over seconds or minutes, because these receptors regulate opening
and closing of ion channels indirectly.
Opening of Acetylcholine-Gated Cation Channels Leads to Muscle Contraction
The nicotinic acetylcholine receptor, a ligand-gated cation channel, admits both K+ and Na+.
Although found in some neurons, this receptor is best known for its role in synapses between
motor neurons and skeletal muscle cells. Patch-clamping studies on isolated outside-out
patches of muscle plasma membranes have shown that acetylcholine causes opening of a
cation channel in the receptor capable of transmitting 15,000-30,000 Na+ or K+ ions a
Since the resting potential of the muscle plasma membrane is near Ek, the potassium
equilibrium potential, opening of acetylcholine receptor channels causes little increase in the
efflux of K+ ions; Na+ ions, on the other hand, flow into the muscle cell. The simultaneous
increase in permeability to Na+ and K+ ions produces a net depolarization to about –15mV from
the muscle resting potential of –85 to –90 mV. This depolarization of the muscle membrane
generates an action potential, which – like an action potential in a neuron – is conducted along
the membrane surface via voltage-gated Na+ channels. When the membrane depolarization
reaches a specialized region, it triggers Ca2+ movement from its intracellular store, the
sarcoplasmic reticulum, into the cytosol; the resultant rise in cytosolic Ca 2+ induces muscle
Two factors greatly assisted in the characterization of the nicotinic acetylcholine receptor. First,
this receptor can be rather easily purified from the electric organs of electric eels and electric
rays; these organs are derived from stacks of muscle cells (minus the contractile proteins) and
thus are richly endowed with this receptor. (In contrast, this receptor constitutes a minute
fraction of the total membrane protein in most nerve and muscle tissues). Second, α-
bungarotoxin, a neurotoxin present in snake venom, binds specifically and irreversibly to
nicotinic acetylcholine receptors. This toxin can be used in purifying the receptor by affinity
chromatography and in localizing it. For instance, in autoradiographs of muscle-cell sections
exposed to radioactive α-bungarotoxin, the toxin is localized in the plasma membrane of
postsynaptic striated muscle cells immediately adjacent to the terminals of presynaptic neurons.
Careful monitoring of the membrane potential of the muscle membrane at a synapse with a
cholinergic motor neuron has demonstrated spontaneous, intermittent, and random ~2-ms
depolarizations for about 0.5-1.0 mV in the absence of stimulation of the motor neuron. Each of
these depolarizations is caused by the spontaneous release of acetylcholine from a single
synaptic vesicle. Indeed, demonstration of such spontaneous small depolarizations led to the
notion of the quantal release of acetylcholine (later applied to other neurotransmitters) and
thereby led to the hypothesis of vesicle exocytosis at synapses. The release of one
acetylcholine-containing synaptic vesicle results in the opening of about 3000 ion channels in
the postsynaptic membrane, far short of the number need to reach the threshold depolarization
that induces an action potential. Clearly, stimulation of muscle contraction by a motor neuron
requires the nearly simultaneous release of acetylcholine from numerous synaptic vesicles.
All Five Subunit in the Nicotinic Acetylcholine Receptor Contribute to the Ion
The acetylcholine receptor from skeletal muscle is a pentameric protein with a subunit
composition of α2βγδ. Each molecule has a diameter of about 9nm and protrudes about 6nm
into the extracellular space and about 2nm into the cytosol (Figure 11-36). The α, β, γ, and δ
subunits have considerable sequence homology; on average, about 35-40 percent of the
residues in any two subunits are similar. The complete receptor has a five-fold symmetry, and
the actual cation channel is a tapered central pore, with a maximum diameter of 2.5 nm, formed
by segments from each of the five subunits (Figure 11-36).
The channel opens when the receptor cooperatively binds two acetylcholine molecules to sites
located at the interfaces of the αδ and αγ subunits. Once acetylcholine is bound to a receptor,
the channel is opened virtually instantaneously, probably within a few microseconds. Studies
measuring the permeability of different small cations suggest that the open ion channels is, at its
narrowest, about 0.65-0.80 nm in diameter, in agreement with estimates from electron
micrographs. This would be sufficient to allow passage of both Na+ and K+ ions with their bound
shell of water molecules.
Although the structure of the central ion channel is not known in molecular detail, much
evidence indicates that it is lined by five transmembrane M2 α helices, one from each of the five
subunits. The M2 helices are composed largely of hydrophobic or uncharged polar amino
acids, but negatively charged aspartate or glutamate residues are located at each end, near the
membrane faces and several serine or threonine residues are near the middle. If a single
negatively charged glutamate or aspartate in one subunit is mutated to a positively charged
lysine, and the mutant mRNA is injected together with mRNAs for the other three wild-type
subunits into frog oocytes, a functional channel is expressed, but its ion conductivity – the
number of ions that can cross it during the open state – is reduced. The greater the number of
glutamate or aspartate residues mutated (in one or multiple subunits), the greater that reduction
in conductivity. These findings suggest that aspartate and glutamate residues – one residue
from each of the five chains – form a ring of negative charges on the external surface of the
pore that help to screen out anions and attract Na+ or K+ ions as they enter the channel (see
Figure 11-36). A similar ring of negative charges lining the cytosolic pore surface also helps
select cations for passage.
The two acetylcholine-binding sites in the extracellular domain of the receptor lie ~4 to 5 nm
from the center of the pore. Binding of acetylcholine thus must trigger conformational changes
in the receptor subunits that can cause channel opening at some distance from the binding
sites. Receptors in isolated postsynaptic membranes can be trapped in the open or closed
state by rapid freezing in liquid nitrogen. Images of such preparations suggests that the five M2
helices rotate relative to the vertical axis of the channel during opening and closing.
Two Types of Glutamate-Gated Cation Channels May Function in a Type of
The hippocampus is the region of the mammalian brain associated with many types of short-
term memory. Certain types of hippocampal neurons, here simply called postsynaptic cells,
receive inputs from hundreds of presynaptic cells. In long-term potentiation a burst of
stimulation of a post-synaptic neuron makes it more responsive to subsequent stimulation by
presynaptic neurons. For example, stimulation of a hippocampal presynaptic nerve with 100
depolarizations acting over only 200 milliseconds causes an increased sensitivity of the
postsynaptic neuron that lasts hours to days. Changes in the responses of postsynaptic cells
may underlie certain types of memory.
Two types of glutamate-gated cation channels in the postsynaptic neuron participate in long-
term potentiation. Unlike other neurotranmitter-gated ion channels, both glutamate receptors
have four subunits, each containing a pore-lining M2 helix; both are excitatory receptors,
causing depolarization of the plasma membrane when activated. Because the two receptors
were initially distinguished by their ability to be activated by the non-natural amino acid N-
methyl-D-aspartate (NMDA), they are called NMDA glutamate receptors and non-NMDA
As illustrated in Figure 11-41, non-NMDA receptors are “conventional” in that binding of
glutamate, released from the presynaptic cell, triggers their opening. NMDA glutamate
receptors are different in two key respects. First, they allow influx of Ca 2+ as well as Na+.
Second, and more important, two conditions must be fulfilled for the ion channel to open:
glutamate must be bound and the membrane must be partly depolarized. In this way, the
NMDA receptor functions as a coincidence detector; that is, it integrates activity of the
postsynaptic cell – reflected in its depolarized plasma membrane – with release of
neurotransmitter from the presynaptic cell, generating a cellular response greater than that
caused by glutamate release alone. Once a post-synaptic cell becomes “sensitized”, it takes
fewer action potentials in the presynaptic neurons to induce a given depolarization in the
postsynaptic neuron; in other words, the synapse “learns” to have an enhanced response to
signals from the presynaptic cells.
Opening of NMDA receptors depends on membrane depolarization because of the voltage-
sensitive blocking of the ion channel by a Mg2+ ion from the extracellular solution. A small
depolarization of the membrane causes the Mg 2+ ion to dissociate from the receptor, thereby
making it possible for glutamate binding to open the channel. Mutagenesis of a single
asparagine residue in the pore-lining M2 helix of the NMDA receptor abolishes the effect of
Mg2+, indicating that Mg2+ binds in the channel.
Since activation of a single synapse, even at high frequency, generally causes only a small
depolarization of the membrane of the postsynaptic cell, long-term potentiation is induced only
when many synapses simultaneously stimulate a single postsynaptic neuron. Thus, the
requirements for membrane depolarization explains why long-term potentiation depends on the
simultaneous activation of a large number of synapses on the postsynaptic cell.