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8. Gillebert, T. C., S. U. Sys, and D. L. Brutsaert. Influence of loading patterns
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slows the rate of isovolumic pressure fall in patients with normal coronary
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15. Zile, M. R., and W. H. Gaasch. Mechanical loads and the isovolumic and filling
indices of left ventricular relaxation. Progr. Cardiovasc. Dis. 32: 333–346, 1990.
Peter Jonas
Glutamate is the main excitatory transmitter in the mammalian CNS, mediating fast synaptic
transmission primarily by activation of AMPA-type glutamate receptor channels. Both synaptic structure and a cell-specific molecular switch in the AMPA receptor subunit expression are involved in the regulation of the synaptic signaling time course.
C
hemical synaptic transmission involves the release of a
transmitter into the synaptic cleft and the subsequent activation of postsynaptic receptors. The most extensively characterized synapse is the neuromuscular junction, formed between
motor axons and skeletal muscle fibers. The transmitter acetylcholine (ACh) is released from synaptic vesicles in multimolecular packets, or quanta, and activates nicotinic ACh receptors
(AChRs) in the postsynaptic membrane. This generates postsynaptic conductance changes of fast time course and large
amplitude, which induce reliable activation of the muscle fiber.
In the mammalian central nervous system (CNS), glutamate is
the main excitatory transmitter. The principles of synaptic communication at glutamatergic synapses, however, appear to be
more complex than at the neuromuscular junction. First, glutamatergic synapses differ in morphological properties, such as
number of release sites and presence of dendritic spines. Second, synapses in different circuitries differ substantially in impact
and time course of synaptic signaling. Finally, glutamate activates several different types of ionotropic and metabotropic
receptors: α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate
receptors (AMPARs), N-methyl-D-aspartate receptors (NMDARs),
kainate receptors (KARs), and metabotropic glutamate receptors
coupled to either inositol 1,4,5-trisphosphate (IP3; class 1
mGluRs) or cAMP-signaling cascades (class 2 and 3 mGluRs)
(10). On the basis of extensive work in the last ten years, we are
beginning to understand how cellular and molecular factors
shape functional differences between glutamatergic synapses.
P. Jonas is in the Physiologisches Institut der Universität Freiburg, D-79104
Freiburg, Germany.
0886-1714/99 5.00 © 2000 Int. Union Physiol. Sci./Am.Physiol. Soc.
The time course of the excitatory postsynaptic currents
The patch-clamp technique in brain slices developed by
Bert Sakmann and co-workers allows us to record from neurons in the CNS with high resolution in an intact synaptic
environment. Analysis of several synapses revealed that fast
glutamate-mediated signaling is predominantly mediated by
AMPARs (5, 9, 15). However, the time course of signaling differs substantially between synapses. This is illustrated in Fig. 1,
which shows AMPAR-mediated evoked excitatory postsynaptic currents (EPSCs) at three well-characterized synapses: the
mossy fiber-CA3 pyramidal cell synapse in the hippocampus
(9), the granule cell-basket cell synapse in the dentate
gyrus (5), and the calyx synapse on neurons in the medial
nucleus of the trapezoid body (MNTB) in the brain stem (1).
Throughout the CNS, slow AMPAR-mediated evoked EPSCs
are generated at excitatory synapses on hippocampal and
neocortical principal neurons (decay time constant ~5 ms at
22°C). In contrast, fast AMPAR-mediated evoked EPSCs are
generated at glutamatergic synapses on hippocampal and
neocortical inhibitory interneurons and on neurons in the
auditory pathway (decay time constant ~1–2 ms).
Synchrony vs. asynchrony of quantal release
Which factors determine the time course of the AMPARmediated postsynaptic conductance change? First, asynchrony
of quantal release needs to be considered (Fig. 2A; Refs. 3, 5,
and 8). Evoked release of transmitter involves several steps:
invasion of the action potential into the presynaptic terminal,
activation of voltage-gated Ca2+ channels, Ca2+ entry into the
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The Time Course of Signaling at Central
Glutamatergic Synapses
presynaptic terminal, and binding of Ca2+ to the putative Ca2+
sensor (for example see Ref. 1). Thus it is not surprising that the
synaptic delay is somewhat variable from trial to trial at the
same release site and between different release sites.
Similar to the neuromuscular junction, it appears that
quantal release is highly synchronized at many glutamatergic
synapses. The duration of the early release period at physiological temperature is only a few hundred microseconds at
the granule cell-basket cell synapse (5) and the calyx synapse
on cochlear nucleus neurons (8), despite the presence of
multiple release sites at both synapses. Whether these results
can be extrapolated to all glutamatergic synapses remains to
be determined (see Ref. 3 for information on a higher asynchrony of release in cultured neurons).
Time course of glutamate concentration in the synaptic
cleft
Second, the time course of transmitter concentration in the
synaptic cleft is potentially important for the time course of
the postsynaptic conductance change (Fig. 2B). At the neuromuscular junction, enzymatic transmitter degradation limits
the lifetime of transmitter in the synaptic cleft. At glutamatergic synapses, the mechanisms of transmitter clearance must
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be different, because a high-turnover enzyme analogous to
the ACh esterase is lacking.
Different approaches have been used to determine the time
course of glutamate concentration in the synaptic cleft. The
first approach uses low-affinity competitive antagonists of
AMPARs; the longer the transmitter is present, the less blockage of the EPSC by the competitive antagonist will occur, due
to antagonist displacement (4). A second approach is based
on a comparison of the time course of the quantal EPSC at a
real synapse with that of currents evoked by glutamate pulses
at a surrogate synapse where “presynaptic” parameters can be
readily controlled. Such surrogate synapses are based on the
technique of fast application of agonists to isolated outsideout patches introduced by Josef Dudel and colleagues (2, 5, 6,
7, 12, 15). Finally, computer simulation of transmitter diffusion and receptor activation has provided insights into the factors that determine the shape of the synaptic glutamate pulse.
The results converge on the conclusion that diffusion is the
main factor that promotes transmitter elimination at glutamatergic synapses. If the instantaneous release of 5000 glutamate molecules into an infinite synaptic cleft is simulated
(assuming that the diffusion coefficient for glutamate is half of
that in aqueous solution), a complex time course of the
synaptic glutamate pulse is predicted (Fig. 2B). The initial rate
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FIGURE 1. Differences in time course of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate receptor (AMPAR)-mediated excitatory postsynaptic currents (EPSCs)
between glutamatergic synapses. A: unitary EPSCs generated at mossy fiber-CA3 pyramidal cell synapse in hippocampus, a prototypic synapse between glutamatergic principal neurons. Extracellular electrical stimulation of a single granule cell is shown; time of stimulus is indicated by arrow. From Ref. 9. B: unitary
EPSCs generated at granule cell-basket cell synapse in dentate gyrus, a prototypic synapse between a glutamatergic principal neuron and a GABAergic interneuron. C: unitary EPSCs generated at an auditory synapse (between a neuron in cochlear nucleus and a neuron in medial nucleus of trapezoid body). From Ref. 1.
Paired recording configurations are shown in B and C; presynaptic action potentials are shown above EPSC traces. Recording conditions are similar (membrane
potential near –70 mV and recording temperature ~22 °C). Furthermore, synapses are located close to soma of postsynaptic neuron, indicating that voltage-clamp
errors are negligible and do not contribute to differences observed. Schematic drawings illustrate location of synapses investigated; excitatory neurons are depicted
in green and inhibitory neurons are depicted in red.
the release probability is increased. In principle, cross talk
could occur between release sites-postsynaptic density units
of the same bouton or between those of different synapses.
High probability of release and close spacing of release sites
will favor cross talk, whereas low release probability and large
distance between release sites will minimize cross talk.
At glutamatergic synapses, the effects of cross talk are
much smaller than at the neuromuscular junction. Several
glutamatergic synapses do not show significant cross talk (4,
5). However, at calyx synapses on auditory neurons that have
several hundred release sites, the slight prolongation of the
EPSC with increasing release probability and the presence of
a slowly decaying component of the AMPAR-mediated
evoked EPSC were attributed to cross talk (12, 15).
Gating properties of postsynaptic AMPARs
Gating of the postsynaptic receptors is a third factor that is
likely to be relevant for the time course of the postsynaptic
conductance change (Fig. 2C); at the neuromuscular junction,
receptor gating is thought to be the main determinant of the
decay of the synaptic current. Fast application experiments
using a variety of paradigms indicate that AMPAR gating is
complex. The agonist affinity of AMPARs is relatively low, with
the half-maximal activating glutamate concentrations in the
range of 500 µM–2 mM for the peak current. With saturating
FIGURE 2. Factors that determine time course of postsynaptic conductance change at glutamatergic synapses. A: presynaptic action potential and normalized
release probability distribution are shown superimposed at top. Individual EPSCs in conditions of reduced Ca2+/Mg2+ concentration ratio (1.5 mM Ca2+/3 mM
Mg2+) are shown at bottom. Note high percentage of failures under these conditions. This indicates that majority of events were quantal EPSCs, caused by release
of contents of a single synaptic vesicle. Recording temperature was 34°C. From Ref. 5 (copyright held by Cell press). B: time course of glutamate concentration in
synaptic cleft. Glutamate concentration is plotted logarithmically against radial location (for times 0, 10, 50, 100, and 1000 µs; top) and against time (for center
of synapse at release site and in periphery at 0.2 µm radial distance from release site; bottom). Curves were obtained by analytical solution of diffusion equations.
Assumptions were 5000 glutamate molecules per vesicle, instantaneous release from a point source, infinite synaptic cleft of 20 nM width, and diffusion coefficient for glutamate D = 3 10–6 ·cm2 ·s–1, approximately half of that tabulated for glutamine in aqueous solution. C: a hypothetical gating scheme of native AMPARs,
with two binding sites for glutamate, is shown at top. C0, C1, and C2 are closed, available states, C3, C4, and C5 are closed, desensitized states, and O is the
open state. From Ref. 9. Glutamate-activated currents in an outside-out patch isolated from a CA3 pyramidal neuron are illustrated at bottom. Currents evoked by
1- and 100-ms pulses of 1 mM glutamate from same patch are shown superimposed. Recording temperature was 22°C. From Ref. 5 (copyright held by Cell press).
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of glutamate clearance (from millimolar to micromolar levels) is extremely rapid, but the later phases of elimination
(from micromolar to submicromolar) are much slower (4).
The exact time course of the synaptic glutamate pulse may
depend on microanatomical properties of the synapse and
glutamate transporter density. Small synaptic contact size and
simple synaptic morphology will facilitate rapid transmitter
clearance. In contrast, large synaptic contact size, the presence of spines on the dendrite of the postsynaptic neuron, and
glial wrapping of the contact will slow transmitter clearance.
Glutamate transporters bind glutamate rapidly, whereas the
subsequent translocation step appears to be slow; the cycling
time is ~50 ms at 22°C. Thus binding of glutamate to transporters could shape the early phase of the synaptic glutamate
pulse, whereas the subsequent translocation step could shape
the late phase. Both the high density of transporters and the
location in proximity to the release site (in both neuronal and
glial membranes) would be consistent with the hypothesis that
glutamate transporters act as buffers (4).
Interactions between release site-postsynaptic density units,
referred to as cross talk (or spillover), complicate the picture.
Cross talk was first described at the neuromuscular junction in
the presence of ACh esterase inhibitors. If several vesicles fuse
at neighboring release sites, the transmitter pulses in the
synaptic cleft will summate. This may result in a gradual prolongation of transmitter clearance and evoked EPSC decay as
concentrations of glutamate, AMPARs are activated within a
few hundred microseconds. When the glutamate pulse is brief
(e.g., 1 ms), AMPARs close rapidly after the end of the pulse
because of AMPAR deactivation following removal of the agonist. When the glutamate pulse is more prolonged, AMPARs
close more slowly because of AMPAR desensitization in the
maintained presence of the agonist (Fig. 2C; Refs. 2, 6, 7, 12,
and 15). The extent of desensitization is substantial (>90% with
saturating glutamate concentrations). Somewhat unexpectedly,
AMPARs enter desensitized states not only during prolonged
applications of glutamate but also after brief glutamate pulses
(a phenomenon referred to as “brief-pulse desensitization”;
Refs. 2 and 7). Furthermore, low concentrations of glutamate
(2–10 µM), when present for several seconds, can induce
“equilibrium desensitization” without significant receptor activation (2). These gating properties can be accounted for by
cyclic reaction schemes, similar to that shown in Fig. 2C (9).
The complex gating of AMPARs raises the question of which
steps are limiting during the EPSC decay. Consistent with the
rapid decline of glutamate concentration during the early
phase of transmitter clearance, the decay of the quantal EPSC
is very similar to the deactivation time constant at many glutamatergic synapses. The granule cell-basket cell synapse,
where both time constants are identical, represents a particularly remarkable example (Fig. 1B; Ref. 5). However, in conditions that favor slow transmitter clearance (large contact
area, spines, glial wrapping) or cross talk (high release probability, high number of closely spaced release sites), the synaptic glutamate pulse may be prolonged and the decay of the
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evoked EPSC may approach the desensitization time constant.
The calyx synapse on auditory neurons (15) and the glutamatergic synapses on cerebellar Purkinje cells exemplify such
a scenario. The low concentration of glutamate during the
slow phase of transmitter clearance, although insufficient for
receptor activation, may lead to receptor desensitization.
If AMPAR desensitization does not shape the EPSC
directly at many synapses, one wonders whether it could
have any other functional roles. One possibility is that
desensitization contributes to paired-pulse depression during repetitive activation of a synapse. This may occur in
auditory synapses, in which paired-pulse depression is sensitive to the desensitization inhibitor cyclothiazide (15). A
second possibility is that desensitization induced by ambient
glutamate regulates the amplitude of the EPSC. Ambient glutamate concentrations in the extracellular space are thought
to be in the range of a few micromoles, which may be sufficiently high to induce equilibrium desensitization. Finally,
AMPAR desensitization will minimize cross talk effects and
thus ensure specificity of transmission at individual sites.
If gating kinetics shape the decay of the postsynaptic conductance change, regulation of gating kinetics may generate
synapse-specific differences in signaling time course. Analysis
by fast application techniques indicates that AMPARs expressed
in different types of neurons differ substantially in their functional properties (Fig. 3A). Throughout the CNS, AMPARs with
slow deactivation and desensitization are expressed in hippocampal and neocortical principal neurons (deactivation time
constant ~3 ms and desensitization time constant ~15 ms at
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FIGURE 3. Regulation of AMPAR gating by subunit composition. A: currents evoked by 1- and 100-ms pulses of 1 mM glutamate in outside-out patches isolated
from a CA3 pyramidal neuron, a dentate gyrus basket cell, and a neuron in medial nucleus of trapezoid body (MNTB). Mean values of deactivation and desensitization time constants are indicated below traces. All recordings were made at ~22°C. B: abundance bar graphs, indicating relative abundance of glutamate receptors (GluR)-A, -B, -C, and -D mRNA in 3 types of neurons. Data are averages from 10, 10, and 8 individual cells, respectively. Open portions of bars indicate flip
splice variants, and filled portions depict flop splice variants. For GluR-C, splice variants were not analyzed. Putative subunit composition of GluR transcripts was
determined by single-cell RT-PCR with AMPAR-specific primers. Quantitation of relative abundance of GluR-A, -B, -C, and -D transcripts in mixed PCR product
was performed by subcloning and phage plaque counting. Modified from Ref. 5 (copyright held by Cell press).
22°C), whereas AMPARs with faster gating are present in
inhibitory interneurons (2, 6, 7). The fastest AMPARs throughout
the CNS are expressed in neurons of the MNTB and the
cochlear nucleus (deactivation time constant 0.3–0.9 ms and
desensitization time constant ~1.7 ms; Refs. 6, 12, and 15).
Molecular mechanisms underlying differences between
synapses
In the past few years, our understanding of glutamatergic
synaptic transmission has gradually improved, reaching a
level of depth comparable to that at the neuromuscular
junction. Several links between the time course of synaptic
signaling, receptor function, and receptor subunit composition have been discovered. First, AMPAR gating is the primary
determinant of the decay of the postsynaptic conductance
change, but synchrony of release and time course of transmitter concentration in the synaptic cleft are also relevant at
certain synapses. Second, AMPAR gating is regulated by the
subunit composition, with certain subunits determining
slow gating and others conferring fast gating. Differential
AMPAR subunit expression not only generates differences
between AMPARs expressed in different regions of the brain
but also between those in different types of neurons of the
same circuitry.
Why do central neurons put effort into the generation of
synapse-specific postsynaptic conductance changes by running
distinct genetic programs? The answer is probably that the time
course of the postsynaptic conductance change at glutamater-
“AMPAR is the main type of glutamate receptor that
mediates fast excitatory synaptic transmission.”
gic synapses is of critical importance for neuronal function.
One aspect is that the time course of the postsynaptic conductance change shapes the rising phase of the excitatory postsynaptic potential (EPSP). This could be of critical importance
for the timing of action potentials evoked in the postsynaptic
neuron, since activation of voltage-gated Na+ channels requires
fast voltage changes. Furthermore, the time course of the postsynaptic conductance change shapes the decay phase of the
local dendritic EPSP and, if the synapse is located close to the
soma, the somatic EPSP (5). This could determine the extent of
the temporal and spatial summation of synaptic signals.
What is the functional significance of the synapse-specific
differences in the time course of the postsynaptic conductance change for the operation of neuronal networks? A final
conclusion cannot be reached yet. It is remarkable, however, that slow AMPAR-mediated EPSCs are generated in
neurons specialized to integrate synaptic signals, such as
cortical principal neurons, whereas fast EPSCs are generated
in neurons that operate as coincidence detectors, such as
MNTB neurons that are specialized on sound localization.
This may suggest that the time course of the AMPAR-mediated conductance change determines whether a neuron
operates as an integrator, coincidence detector, or fast relay
of synaptic input signals.
Basics of glutamatergic synaptic
transmission in the CNS
AMPAR: The main type of glutamate receptor that mediates
fast excitatory synaptic transmission. As the name indicates, it is selectively activated by the agonist a-amino-3News Physiol. Sci. • Volume 15 • April 2000
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The substantial differences between AMPARs expressed in
different types of neurons raise the question of the molecular
basis of these differences. Native AMPARs are heteromers
assembled from GluR-A to -D subunits. Additional structural
diversity is created by alternative splicing of the flip/flop module and by RNA editing at different sites (10).
Studies on recombinant AMPARs expressed in host cells
show that gating kinetics are regulated by subunit composition. In oocytes and mammalian cell lines, both homomeric
and heteromeric channels can be expressed heterologously,
indicating a high degree of flexibility in AMPAR assembly.
This distinguishes AMPARs from nicotinic AChRs in skeletal
muscle, which are assembled in constant subunit stoichiometry. In homomeric AMPARs, the desensitization time constant
follows the sequence: GluR-Ai ~ GluR-Ao ~ GluR-Ci ~ GluRDi > GluR-Co ~ GluR-Do, with ‘o’ designating the flop and
‘i’ the flip splice form (11, 13). Furthermore, heteromeric
AMPARs formed by GluR-Bi in combination with other
subunits have slower desensitization than the respective
homomers, whereas GluR-Bo has little effect (11, 13). Thus
GluR-Bi is a determinant of slow desensitization, whereas
GluR-Co and GluR-Do are determinants of fast desensitization of recombinant AMPARs.
Similar principles of regulation may apply to native
AMPARs (Fig. 3B). Correlated analysis by fast application and
single-cell RT-PCR (a technique introduced by Jean Rossier
and colleagues) has shed light onto the molecular determinants of native receptor function. Analysis in different types
of cells revealed that the desensitization time constant is positively correlated with the relative abundance of GluR-Bi
mRNA and negatively correlated with the relative abundance
of GluR-D mRNA (6). Similar correlations are found for the
deactivation time constant. Thus the results from both recombinant and native AMPARs indicate that AMPAR gating
(deactivation and desensitization) is regulated reciprocally by
GluR-Bi and GluR-D subunits. Other molecular factors, such
as phosphorylation, may also affect AMPAR gating, but direct
experimental evidence is not yet available.
Different AMPAR subunit composition may not only generate functionally distinct AMPARs in different types of neurons but also at different synapses on the same neuron. In
bipolar cells of the cochlear nucleus, GluR-D subunit protein
is targeted specifically to the basal dendrites, the region
where the auditory afferents terminate (14). This could imply
that glutamatergic synapses established on different dendrites
of these neurons generate postsynaptic conductance changes
with very different time courses. Whether sorting also occurs
in other types of neurons and for AMPAR subunits other than
GluR-D remains to be elucidated.
Functional significance
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News Physiol. Sci. • Volume 15 • April 2000
editing sites of AMPAR subunit mRNA have been identified: the Q/R site in the pore-forming region of the GluRB subunit and the R/G site preceding the flip/flop module of GluR-B, -C, and -D subunits. Q/R-site editing
changes the Ca2+ permeability of AMPARs by more than
an order of magnitude, whereas R/G-site editing is
involved in fine tuning of AMPAR gating.
Paired-pulse facilitation or depression: When two action potentials are elicited in a presynaptic neuron, separated by a
short time interval, the amplitude of the second EPSC may
be either enhanced or reduced as compared to that of the
first, depending on the type of synapse and the probability
of release. Paired-pulse modulation appears to be generated mainly by presynaptic mechanisms, but postsynaptic
factors (such as desensitization) may also contribute at certain synapses.
Postsynaptic density: Specialized postsynaptic, electrondense site that presumably contains the transmitter
receptors.
Release site: Specialized presynaptic site where vesicle
fusion is thought to occur.
Spine: Small, thorn-like protrusion of dendritic membrane,
which may subserve the purpose of increasing surface
area and producing compartmentalization. In the hippocampus and the neocortex, glutamatergic synapses
between principal neurons are mainly formed on spines,
whereas principal neuron-interneuron synapses are
established on dendritic shafts (inhibitory interneurons
characteristically lack spines). Narrow spine necks may
delay the escape of glutamate from the synaptic cleft.
Synaptic delay: Time interval from the steepest point in the rising
phase of the presynaptic action potential (recorded close to
the site of release) to the beginning of the evoked EPSC.
I thank Drs. J. R. P. Geiger, K. Haverkampf, and B. Taskin for helpful
discussions.
This review is based on a lecture given in Lisbon in March 1998 on the
occasion of the first award of the Medinfar European Prize in Physiology.
My studies are supported by the Deutsche Forschungsgemeinschaft, the German Israeli Foundation, and the Human Frontiers Science Program Organization.
I regret that many relevant papers could not be cited due to editorial restrictions on the maximum number of references.
References
1. Borst, J. G. G., F. Helmchen, and B. Sakmann. Pre- and postsynaptic
whole-cell recordings in the medial nucleus of the trapezoid body of the
rat. J. Physiol. (Lond.) 489: 825–840, 1995.
2. Colquhoun, D., P. Jonas, and B. Sakmann. Action of brief pulses of glutamate on AMPA/kainate receptors in patches from different neurones of rat
hippocampal slices. J. Physiol. (Lond.) 458: 261–287, 1992.
3. Diamond, J. S., and C. E. Jahr. Asynchronous release of synaptic vesicles
determines the time course of the AMPA receptor-mediated EPSC. Neuron
15: 1097–1107, 1995.
4. Diamond, J. S., and C. E. Jahr. Transporters buffer synaptically released glutamate on a submillisecond time scale. J. Neurosci. 17: 4672–4687, 1997.
5. Geiger, J. R. P., J. Lübke, A. Roth, M. Frotscher, and P. Jonas. Submillisecond AMPA receptor-mediated signaling at a principal neuron-interneuron
synapse. Neuron 18: 1009–1023, 1997.
6. Geiger, J. R. P., T. Melcher, D.-S. Koh, B. Sakmann, P. H. Seeburg, P. Jonas,
and H. Monyer. Relative abundance of subunit mRNAs determines gating
and Ca2+ permeability of AMPA receptors in principal neurons and
interneurons in rat CNS. Neuron 15: 193–204, 1995.
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hydroxy-5-methyl-4-isoxazolepropionate. Some authors
designate this receptor as the AMPA/kainate receptor,
since it is also activated by kainate. Unlike the responses
to AMPA and glutamate, those to kainate do not desensitize. Several years ago this receptor had been referred to
as the “quisqualate receptor” but was renamed because
quisqualate turned out to be a nonselective agonist, also
activating class 1 mGluRs.
Cross talk: When transmitter is released simultaneously at
closely spaced sites, the transmitter pulses in the synaptic
cleft summate. This may lead to a slowing in transmitter
clearance, which would result in a prolongation of the postsynaptic current.
Deactivation: Closure of AMPAR channels after removal of
the agonist.
Desensitization: Closure of AMPAR channels in the maintained presence of the agonist, which is approximately
threefold slower than deactivation. Although desensitization is a general characteristic of ligand-gated ion
channels, it is particularly fast for AMPARs.
EPSC, evoked: EPSC evoked by stimulation of presynaptic
neurons. If a single neuron is stimulated (for example, in
a paired recording configuration), the EPSC would be
denoted as “unitary.” If several presynaptic neurons are
stimulated (by extracellular stimulation of axonal tracts),
the EPSC would be referred to as “composite.”
EPSC, quantal: EPSC generated by the release of the contents
of a single synaptic vesicle. The term “quantum” was
introduced by Bernard Katz for events at the neuromuscular junction, and is also used at glutamatergic synapses.
One approach to isolating quantal events is to record
evoked EPSCs in conditions of low release probability. An
alternative approach is to record miniature EPSCs, i.e.,
spontaneously occurring EPSCs after suppression of presynaptic action potential activity (with tetrodotoxin and/or
blockers of voltage-gated Ca2+ channels).
Failure: Evoked EPSCs show statistical fluctuation in peak
amplitude from trial to trial. Occasionally the presynaptic action potential does not evoke an EPSC in the
postsynaptic neuron at all. This is denoted as failure of
synaptic transmission.
GluR-A to -D: AMPAR subunits identified by molecular
cloning. In an alternative nomenclature, these subunits
are referred to as GluR1 to 4. Each of the subunits consists of ~900 amino acids (sequence identity between
subunits ~70 %). It is thought that each subunit has four
membrane segments (three transmembrane segments
and a loop that forms the aqueous pore). The functional
channel appears to be an oligomer assembled from four
or five subunits.
Flip, Flop: AMPAR subunit variants generated by alternative
splicing of the flip/flop module. This module is comprised
of 115 base pairs (corresponding to 38 amino acids, 9-11
of which are different between flip and flop variants).
RNA editing: Form of posttranscriptional modification of
AMPAR RNA, discovered by Peter Seeburg and colleagues. Desamination of adenosine to inosine is
thought to be the underlying molecular mechanism. Two
7. Hestrin, S. Different glutamate receptor channels mediate fast excitatory
synaptic currents in inhibitory and excitatory cortical neurons. Neuron 11:
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8. Isaacson, J. S., and B. Walmsley. Counting quanta: direct measurements of
transmitter release at a central synapse. Neuron 15: 875–884, 1995.
9. Jonas, P., G. Major, and B. Sakmann. Quantal components of unitary
EPSCs at the mossy fibre synapse on CA3 pyramidal cells of rat hippocampus. J. Physiol. (Lond.) 472: 615–663, 1993.
10. Jonas, P., and H. Monyer (eds.). Ionotropic glutamate receptors in the CNS.
In: Handbook of Experimental Pharmacology. Berlin: Springer, 1999.
11. Mosbacher, J., R. Schoepfer, H. Monyer, N. Burnashev, P. H. Seeburg,
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Peter J. Meier and Bruno Stieger
Active canalicular secretion of bile salts and non-bile salt organic anions represents the major driving
force of hepatic bile formation. The most important carriers involved have now been cloned on
both the basolateral and canalicular sides of hepatocytes. Elucidation of their structure,
transport properties, and regulation is an important step forward in the ultimate
understanding of the molecular physiology of bile formation.
C
ontinuous bile formation is an important function of the
liver, and bile is used as a vehicle for the secretion of
bile acids and the excretion of lipophilic endo- and xenobiotics. It is an osmotic process that is driven by active secretion
of predominantly organic solutes into the minute channels
of bile canaliculi, followed by passive inflow of water, electrolytes, and nonelectrolytes (e.g., glucose) across hepatocytes and tight junctions. Bile salts are quantitatively the
major constituents of bile and are circulated in the enterohepatic circulation between the liver and the small intestine
with high efficiency. Bile is also rich in phospholipids (predominantly phosphatidylcholine) that form mixed vesicles
and micelles with bile salts and thereby provide a means for
the disposal of highly lipophilic endogenous and exogenous
compounds, such as cholesterol, from the body. Canalicular
bile is produced by the highly polarized hepatocytes that
localize a distinct set of transporters at their basolateral
(sinusoidal) and apical (canalicular) plasma membrane
domains (Fig. 1) (8, 11). Primary hepatic bile fluid is then
modified by the bile ductular and gall bladder epithelial
cells before it reaches the small intestine. Because active
canalicular secretion of bile salts represents the major driving force of hepatic bile formation, and because the
canalicular bile salt export pump (bsep) of mammalian liver
has recently been identified and cloned, this overview concentrates on the molecular mechanisms involved in the
hepatocellular transport of bile salts from blood into bile.
P. J. Meier and B. Stieger are in the Division of Clinical Pharmacology and
Toxicology, 1st Department of Medicine, University Hospital Zurich, CH8091 Zurich, Switzerland.
0886-1714/99 5.00 © 2000 Int. Union Physiol. Sci./Am.Physiol. Soc.
Basic mechanisms of hepatic bile formation
Bile salts are concentrated up to 1000-fold in bile compared with portal blood plasma and the hepatocyte cytosol.
This vectorial and concentrative transport process requires a
polar arrangement of the hepatocellular carriers involved
(Fig. 1). The basolateral plasma membrane of the hepatocytes
faces the space of Disse, which is in direct contact with the
portal blood via the fenestrae of the endothelial cells lining
the sinusoids. The basolateral plasma membrane represents
the first barrier to be crossed by cholephilic substances, i.e.,
substances destined to be secreted or excreted into bile. The
canalicular plasma membrane of hepatocytes limits the bile
canaliculi that form the exit site for all cholephilic compounds
from the hepatocytes. The canalicular plasma membrane is
separated from the basolateral plasma membrane by tight
junctions. They form the only anatomical barrier between the
portal blood and the primary bile fluid produced by hepatocytes and are hence of great importance for the maintenance
of the concentration gradients between blood and bile.
Hepatocellular uptake of bile salts across the basolateral
plasma membrane occurs against a concentration gradient and
is driven predominantly by a secondary active sodium-dependent transport system (8, 9, 11). This transport system has been
well characterized in a variety of experimental systems and
exhibits apparent Michaelis-Menten constant (Km) values for
taurocholate between 15 and 56 µM. The system is electrogenic, with a sodium-to-taurocholate stoichiometry of >1. In
addition to sodium/bile salt cotransport, sodium-independent
bile salt transport pathways are also present at the basolateral
plasma membranes of hepatocytes (8, 9, 11). These sodiumindependent bile salt uptake systems have been less well
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Molecular Mechanisms in Bile Formation