Regulation of CNS synapses by neuronal MHC class I
C. Alex Goddard*, Daniel A. Butts†, and Carla J. Shatz‡
Department of Neurobiology, Harvard Medical School, Boston, MA 02115
Contributed by Carla J. Shatz, March 6, 2007 (sent for review January 9, 2007)
Until recently, neurons in the healthy brain were considered
immune-privileged because they did not appear to express MHC
class I (MHCI). However, MHCI mRNA was found to be regulated by
neural activity in the developing visual system and has been
detected in other regions of the uninjured brain. Here we show
that MHCI regulates aspects of synaptic function in response to
activity. MHCI protein is colocalized postsynaptically with PSD-95
in dendrites of hippocampal neurons. In vitro, whole-cell record-
ings of hippocampal neurons from ?2m/TAP1 knockout (KO) mice,
which have reduced MHCI surface levels, indicate a 40% increase
in mini-EPSC (mEPSC) frequency. mEPSC frequency is also increased
100% in layer 4 cortical neurons. Similarly, in KO hippocampal
cultures, there is a modest increase in the size of presynaptic
boutons relative to WT, whereas postsynaptic parameters (PSD-95
puncta size and mEPSC amplitude) are normal. In EM of intact
hippocampus, KO synapses show a corresponding increase in
vesicles number. Finally, KO neurons in vitro fail to respond
normally to TTX treatment by scaling up synaptic parameters.
Together, these results suggest that postsynaptically localized
MHCl acts in homeostatic regulation of synaptic function and
morphology during development and in response to activity block-
ade. The results also imply that MHCI acts retrogradely across the
synapse to translate activity into lasting change in structure.
homeostatic ? neuron ? plasticity ? synapsin ? PSD-95
activity remodels synaptic connections during development is
termed ‘‘activity-dependent plasticity,’’ in which electrical sig-
nals induce specific patterns of gene transcription to alter
synaptic properties and structural connectivity. Genes including
BDNF and CamKII are known to be critical for this plasticity
(2–4); however, many other molecules are likely involved as well.
MHC class I (MHCI) family members are well known for their
roles in cellular immunity, but a neuronal function has not been
generally appreciated. In the immune system, MHCI genes act
in concert with T cell receptors to discriminate self- versus
non-self-proteins. The CNS was considered ‘‘immune privi-
leged,’’ in part, because it was thought that healthy neurons do
not express MHCI protein (5, 6). Recently, MHCI gene family
members have been found at low levels in CNS neurons (7–11).
MHCI mRNA is expressed and regulated in cortical and tha-
lamic neurons during development and is down-regulated by
chronic activity blockade with Tetrodotoxin (TTX) in vivo (7).
MHCI is also a downstream target of the transcription factor
CREB, required for Hebbian synaptic plasticity (8, 12, 13).
MHCI is thus implicated in several forms of activity-dependent
The proteins encoded are heavy chains comprising the largest
portion of the MHCI protein complex. Functional MHCI is
usually a trimer consisting of the heavy chain, ?-2-microglobulin
(?2m), and a 9–11 aa peptide generated from proteosomal
degradation (15). The transporter associated with antigen pro-
cessing [(TAP) a heterodimer of TAP1 and TAP2] is required
for transport of peptide fragments from cytoplasm into the
lumen of the endoplasmic reticulum for assembly (16). For most
MHCI proteins, cell surface expression of heavy chain only
xperience transduced into neural activity is required for
proper brain development (1). The process by which neural
occurs if ?2m and peptide are present (16, 17). In their absence,
both surface and intracellular levels of MHCI are down-
regulated (18). Therefore, brains of mice deficient in ?2m and
TAP1 were studied here as MHCI ‘‘loss of function.’’ These mice
have altered Hebbian synaptic plasticity in the hippocampus and
abnormal patterning of visual system connections (19), reminis-
cent of animals that have undergone blockade of neural activity
(20–22). Despite this new appreciation of MHCI function in
neuronal plasticity and the discovery of a candidate neuronal
receptor (23, 24), however, it is not known whether MHCI
protein is present at CNS synapses or whether it is part of
molecular machinery regulating synaptic function and structure.
Here we investigate the subcellular localization of MHCI and
show that neurons with low levels of MHCI have altered synaptic
function and structure. Moreover, MHCI appears to play a role
in homeostatically regulating aspects of synaptic structure and
function in response to low levels of neural activity.
The subcellular localization of MHCI protein was examined by
immunostaining cultures of hippocampal neurons with a pan-
specific MHCI antibody, Ox18 (7, 25). Punctate immunostaining
is present in soma and dendrites (Fig. 1a); at higher magnifica-
tion, MHCI immunostaining is in spine-like dendritic protuber-
ances (Fig. 1b). To determine prior postsynaptic location for
MHCI, PSD-95 [present at postsynaptic densities of excitatory
synapses (26)] or synapsin [associated with presynaptic vesicles
(27)] was also detected immunofluorescently (Fig. 1b). Signal for
MHCI protein overlaps extensively with PSD-95 signal; 57% of
Ox18 immunoreactive pixels overlap with PSD-95 pixels (Fig.
1c). In contrast, the distribution of synapsin immunoreactivity is
one of close apposition and minimal overlap with MHCI (Fig. 1
b and c), suggesting that the two proteins are in separate,
adjacent pre- and postsynaptic compartments. Thus, MHCI
appears to be located postsynaptically at excitatory synapses,
consistent with a recent report of dendritic localization of MHCI
mRNAs in hippocampal neurons (28).
Given the presence of MHCI at synapses, as well as previously
reported alterations in activity-dependent plasticity in ?2m/
TAP1 knockout (KO) mice (19), it is possible that cultured KO
neurons have altered basal synaptic transmission. Spontaneous
mini-EPSCs (mEPSCs) from WT or KO hippocampal cultures
were recorded by using whole-cell patch-clamp (Fig. 2a). mEP-
SCs from WT neurons have a median instantaneous frequency
of 1.8 Hz and a median amplitude of 7.1 pA. In contrast, the
Author contributions: C.A.G., D.A.B., and C.J.S. designed research; C.A.G. performed re-
search; C.A.G. and D.A.B. contributed new reagents/analytic tools; C.A.G. analyzed data;
and C.A.G. and C.J.S. wrote the paper.
The authors declare no conflict of interest.
Abbreviations: KO, knockout; MHCI, MHC class I; PSD, postsynaptic densities.
*Present address: Department of Neurobiology, Stanford University, Stanford, CA 94305.
†Present address: Department of Physiology and Biophysics, Institute of Computational
Biomedicine, Weill Medical College of Cornell University, New York, NY 10021.
‡To whom correspondence should be addressed. E-mail: email@example.com.
This article contains supporting information online at www.pnas.org/cgi/content/full/
© 2007 by The National Academy of Sciences of the USA
April 17, 2007 ?
vol. 104 ?
Hz). However, mEPSC median amplitude is similar at KO and
WT synapses (Fig. 2a). These results imply that there is an
abnormality in basal synaptic transmission in KO neurons.
MHCI mRNA is observed in many brain regions, including
visual cortical neurons (11, 19, 29). To assess whether sponta-
neous release is abnormal elsewhere in CNS and in a more intact
preparation, mEPSCs were recorded from layer 4 neurons in
frequency of mEPSCs recorded from KO cortical neurons is
100% greater than that from WT neurons, reminiscent of the
hippocampal cultures. As in cultures, mEPSC amplitude is
similar in WT and KO. These findings suggest that alterations in
basal synaptic transmission occur across various brain regions in
mice lacking stable cell surface expression of MHCI.
The increase in mEPSC frequency in KO hippocampal cul-
tures and cortical slices suggests that synaptic organization may
also be altered. An increase in mEPSC frequency could be due
to an increase in the number or density of synapses (30) or to an
increase in probability of release (31, 32). We therefore exam-
ined the structural organization of synapses in KO hippocampal
cultures. First, we verified that MHCI protein levels are reduced
in KO neurons by immunostaining hippocampal cultures with
Ox18 (Fig. 3a). Consistent with many previous studies of non-
neuronal cell types, KO neurons have low levels of MHCI
WT and KO hippocampal cultures were then immunostained
for synapsin I to examine presynaptic terminals. There is a
modest increase in size of synapsin-immunoreactive boutons in
KO cultures: They are ?6% larger than WT (P ? 0.03) (Fig. 3).
Furthermore, boutons immunostained for vGluT1 and vGluT2,
glutamate transporter (34, 35), are also roughly 6% larger in KO
(P ? 0.02) (Fig. 3 b and c). Together these observations suggest
a general alteration in excitatory presynaptic boutons, rather
than a change specific to either protein alone.
Postsynaptic organization of WT and KO neurons was as-
sessed by immunostaining for PSD-95, known to correlate with
AMPA receptor levels in spines (30, 36, 37). Unlike the observed
presynaptic changes, PSD-95 immunostained puncta in KO
neurons are not significantly different in size from WT (Fig. 3 b
and c), nor did synaptic density differ between WT and KO
[supporting information (SI) Fig. 6]. Thus, it appears that
diminished postsynaptic levels of MHCI protein are associated
with a modest enlargement of presynaptic boutons. The signif-
icant increase in mEPSC frequency is probably due to this
presynaptic change, rather than to alterations in synaptic density.
Because synapsin levels, and likely mEPSC frequency, covary
with vesicle number and release (31, 32), it is possible that
differences between WT and KO hippocampal synapses are
evident in the electron microscope (Fig. 4). Samples were
prepared from 44- to 45-day-old WT and KO intact hippocampi;
the number of vesicles in a cluster and the length of PSDs were
analyzed from single-plane sections of stratum radiatum in CA1.
KO presynaptic nerve terminals contain 10% more synaptic
vesicles than WT terminals (P ? 0.04) (Fig. 4b). In contrast,
measurements of PSD length in hippocampus do not differ
significantly between genotypes (Fig. 4c), suggesting that
postsynaptic parameters are not drastically changed, consistent
with hippocampal neurons in vitro. However, postsynaptic orga-
hippocampal neurons (2 weeks in vitro) detected with Ox18 compared with
equimolar amount of control mouse IgG (Inset). Immunoreactivity is present
in soma and dendrites. (Scale bar: 20 ?m.) (b) High-magnification views of
MHCI and synaptic protein immunoreactivity. Antisynapsin antibodies were
detected with Cy-2 linked secondary, anti-MHCI with Cy3, and anti-PSD-95
with Cy5. Synaptic proteins are pseudocolored green for comparison with
MHCI (red). (Top) PSD-95 immunostained puncta marked by arrows; merged
PSD-95 and MHCI signals, with nearly complete colocalization. (Middle) MHCI
immunostaining of dendrites and spines. (Bottom) Synapsin immunostaining
apposes but does not overlap MHCI immunostaining. (Scale bar: 10 ?m.) (c)
Quantification of the percentage overlap of MHCI immunostaining (Ox18
pixels overlap with PSD-95 pixels. However, only 26.1 ? 9.8% MHCI pixels
is statistically significant (P ? 0.001, t test). Overlap between synapsin and
PSD-95, 19.3 ? 6.9%. The amount of overlap between synapsin–MHCI vs.
synapsin–PSD-95 was not significantly different (P ? 0.2, t test) (n ? 7 fields of
325 ? 325 ?m at 512 ? 512 pixel resolution).
MHCI is expressed at or near synaptic sites. (a) MHCI expression in
Whole-cell recordings from hippocampal neurons in culture. (Left) Frequency
of mEPSCs recorded in the basal state from KO (2.5 Hz, n ? 17 neurons) is
greater than that of WT (1.8 Hz, P ? 0.001, n ? 22 neurons), but amplitudes
do not differ (WT, 7.1; KO, 7.0; P ? 0.98). Data presented as ratio of KO/WT
median instantaneous frequency (1 per interevent interval) or median ampli-
tude.*, significance as calculated by Kolmogorov–Smirnov test of cumulative
distribution of events (see Fig. 5 a and b). (Right) Representative traces of
mEPSC recordings from WT or KO neurons (2.5 weeks in vitro). (Scale bar: 10
10.7 ? 1.6; P ? 0.92).*, significance by t test. (Right) Representative traces of
mEPSC recordings from WT or KO neurons. (Scale bar: 10 pA; 400 msec.)
Basal synaptic function is altered in ?2m/TAP1 KO neurons. (a)
Goddard et al. PNAS ?
April 17, 2007 ?
vol. 104 ?
no. 16 ?
nization at KO hippocampal synapses is not entirely normal: The
percent perforated PSDs is reduced in KO to about half the
number in WT (P ? 0.006) (Fig. 4d). Together these observa-
tions demonstrate that the synaptic changes in ?2m/TAP1
mutant mice are both structural and functional.
Neuronal MHCI was discovered in a screen for genes regu-
lated by blockade of neural activity with Tetrodotoxin (TTX) in
vivo (7). In hippocampal cultures, TTX treatment is known to
increase both mEPSC amplitude and frequency (38, 39). Fur-
thermore, ?2m/TAP1 KO mice have abnormal retinogeniculate
projections, reminiscent of animals that have undergone chronic
activity blockade (1, 20); these mice also have altered hippocam-
pal synaptic plasticity (19). Together these findings suggest that
MHCI may regulate a neuron’s response to changes in levels of
activity. To test this hypothesis, hippocampal cultures of both
genotypes were grown in 1 ?M TTX for 3–6 days and mEPSCs
were recorded. Consistent with other reports, TTX treatment
increased both mEPSC amplitude and frequency in WT hip-
pocampal cultures. Median mEPSC frequency recorded from
WT neurons treated with TTX increases 22% over WT neurons
in vehicle, from 1.8–2.2 Hz. Median mEPSC amplitude increases
more modestly from 7.0 pA in vehicle to 7.8 pA in TTX (Fig. 5
a and b). However, mEPSCs recorded from KO neurons grown
in TTX do not increase significantly either instantaneous fre-
quency or amplitude compared with KO neurons in vehicle (Fig.
5 a and b). This failure to regulate synaptic function in response
to TTX blockade is not due to a problem with spiking activity in
the KO cultures: Spontaneous spiking activity recorded with
cell-attached patch is indistinguishable from WT cultures (Fig.
5c; SI Fig. 7).
Chronic TTX blockade regulates synaptic protein levels and
vesicle release in cultured neurons (39–41). Consistent with the
physiological observations above and previous reports, WT
boutons immunostained for synapsin following TTX were 15 ?
5.0% larger than in vehicle control (SI Fig. 8 a and b). In these
WT cultures, the size of PSD-95 immunoreactive puncta also
increased (9.6 ? 5.8%) (SI Fig. 8c). This increase in PSD-95
levels following TTX treatment (30, 36, 37, 42). Unlike the
increase in WT neurons following TTX treatment, PSD-95
puncta size in KO neurons does not increase (SI Fig. 8). No
detectable change in synapse density was observed in either
genotype following TTX treatment (data not shown). Thus,
action potential activity blockade in WT cultures produces the
expected changes in both pre- and postsynaptic morphological
parameters, but the changes are more modest than reported
previously. We attribute this difference to the automated mea-
surements performed here by using stringent thresholding cri-
teria (see SI Methods). Our observations also imply that KO
synapses are unable to regulate synaptic structure in response to
TTX: Presynaptic boutons are already enlarged and do not
increase further with TTX, whereas postsynaptic properties such
as PSD-95 puncta fail to adjust at all.
Together, the morphological and physiological abnormalities
of KO neurons are reminiscent of synaptic changes following
Intensity of MHCI immunostaining decreases in KO neurons (Right), whereas
levels of synapsin increase compared with WT (2 weeks in vitro) (Left). (Scale
bar: 20 ?m.) (b) Synapsin (Left) and VGluT1/2 (Center) immunostained puncta
appear larger in KO compared with WT; PSD-95 (Right) appears unchanged.
(Scale bars: 10 ?m.) (c) Quantification of synapsin, vGluT1/2, and PSD-95
immunoreactive puncta. Synapsin bouton size (average no. of pixels per
bouton) is 6.2 ? 3.3% larger in KO relative to WT control (P ? 0.03, n ? 6
experiments). vGluT1/2 immunoreactive boutons in KO are 6.3 ? 1.6% larger
than WT control (P ? 0.02, n ? 3 experiments). PSD-95 puncta size in KO is not
SD.*, statistical significance by t test.
Synaptic structure is altered in KO hippocampal neurons in vitro. (a)
resentative EM sections from WT and KO hippocampus. Black arrowhead,
example of a perforated synaptic contact. (Scale bar: 50 nm.) (b) Increase in
Data represent average of mean vesicle number per synaptic plane for three
animals of each genotype ? SEM. A total of 522 WT synapses and 563 KO
synapses were counted. (c) Postsynaptic densities (PSD) are not significantly
(d) Fewer perforations at KO hippocampal synapses than WT (3.8 ? 0.6%
average perforated per total synapses per animal) compared with WT (6.8 ?
0.3%) (P ? 0.006).
Hippocampal synaptic ultrastructure is altered in KO mice. (a) Rep-
www.pnas.org?cgi?doi?10.1073?pnas.0702023104Goddard et al.
homeostatic synaptic plasticity or ‘‘synaptic scaling’’ (43, 44).
Presynaptic bouton size and mEPSC frequency scale up follow-
ing chronic activity blockade with TTX (38, 40, 45), reaching
values similar to untreated cultures of ?2m/TAP1 KO neurons.
Moreover, MHCI was discovered in a differential display screen
for genes regulated by TTX (7), suggesting it may contribute to
the homeostatic response to changes in neuronal activity. If so,
then treating WT hippocampal neurons with TTX should down-
regulate MHCI protein levels. Indeed, levels of MHCI immu-
noreactivity in WT cultures treated with TTX decreased to 31%
of vehicle-treated cultures (SI Fig. 9). Thus, the increase in
bouton size in WT hippocampal cultures grown in TTX is
correlated with a decrease in neuronal MHCI protein. These
observations suggest a function for MHCI as a key regulator of
the presynaptic nerve terminal’s response to sustained changes
in neural activity.
A major finding of this study is that MHCI is part of the
molecular machinery regulating synaptic morphology and func-
tion under basal conditions and following action potential
blockade. Four independent methods, whole-cell recordings and
immunohistochemistry from hippocampal neurons in vitro,
whole-cell recordings from cortical neurons in slices, and EM in
intact hippocampus, have been used to demonstrate that syn-
aptic structure and function are altered in ?2m/TAP1 mutant
mice lacking stable cell surface expression of MHCI. The
functional changes assessed with physiology (e.g., increased
mEPSC frequency) are more robust than the observed alter-
ations in presynaptic morphology (e.g., size of immunostained
boutons, EM vesicle numbers), but both are well correlated and
support a role for MHCI in the control of synaptic function.
is due to an increase in the number of silent synapses (39, 46),
we did not observe changes in synaptic density as estimated by
immunostaining for PSD-95 or synapsin. Note also that the
synaptic defects in KOs observed here are not in the initial
assembly and function of synapses. Although synaptic function is
altered, it is modest and not pathological, and action potential
generation within these cultures, as well as in the intact animal
(19), is within a normal range.
MHCI protein is detected at postsynaptic membranes of WT
neurons, but presynaptic abnormalities are observed at KO
synapses in the basal state, implying that MHCI may function as
part of a retrograde signaling system. At present, it is not known
whether this retrograde effect is direct, via transsynaptic signal-
ing with a presynaptic MHCI receptor such as PIRB (23) that
might modulate synaptic vesicle dynamics, or instead whether it
is indirect, via interactions with postsynaptic components such as
glutamate receptors. Because both MHCI mRNA and postsyn-
aptically localized proteins are regulated by neural activity,
MHCI could link presynaptic function with postsynaptic activity.
It should be noted that normally pre- and postsynaptic elements
are coregulated (30). However, in KO mice, there appears to be
a mismatch: Presynaptic boutons, but not PSD-95 puncta, are
enlarged in the basal state (and neither pre- nor postsynaptic
elements respond normally to activity blockade). Our study
suggests that MHCI may function to coordinate communication
across the two sides of the synapse.
There is evidently also a problem in the homeostatic adjust-
ment of synapses to match pre- with postsynaptic size in response
to changes in neural activity within developing circuits in KO
mice. Several experimental manipulations known to induce
homeostatic synaptic scaling also increase both mEPSC fre-
quency and the probability of release at excitatory synapses (12,
40, 47). Because the physiological relationship between sponta-
neous mEPSCs and evoked synaptic transmission is not entirely
in mEPSC frequency seen here indicate an increase in the
probability of release at KO synapses. However, Murthy et al.
(42) observed an increase in vesicle number following TTX
treatment, consistent with our observed increase in synapsin
immunoreactivity. The increases in synapsin and vGluT immu-
noreactivity also correlate with the increase in vesicle number
observed in our EM analysis of intact KO hippocampus. Al-
though the increase in vesicle number is modest (?10%), it is
similar in magnitude to the changes observed here in synapsin
and vGluT immunostaining. Note also that the EM analysis is
likely to be an underestimate because synapses were assessed in
single-plane images, only allowing 2D observation of the 3D
vesicle pool. Serial section comparison of WT and KO neurons
will be necessary to obtain more accurate vesicle counts.
We report that MHCI is needed for homeostatic scaling of
synapses in response to activity blockade in vitro. Because our
focus was on glutamatergic synapses, we do not know whether
MHCI is required for homeostatic adjustments at inhibitory
synapses, which could involve different mechanisms (50), nor do
we know which of the ?70 MHCI family members might
contribute. Other molecules with immune function, such as
TNF?, are also involved in homeostatic scaling at excitatory
synapses (51). TNF? is known to regulate MHCI levels in
hippocampal neurons in vitro (25) and thus may act via an
MHCI-dependent pathway. Here we also show that MHCI
protein levels are regulated by TTX. This finding is consistent
with the decrease in MHCI mRNA observed after infusion of
TTX into the developing brain (7) and with the increase in
MHCI protein following glutamate receptor activation in cul-
tured neurons (9). In the hippocampus, MHCI is downstream of
the transcription factor CREB (8), suggesting that it is part of a
molecular program for synapse scaling that also includes other
in either 1 ?M TTX or citrate control and then mEPSCs were recorded. (a)
Cumulative distribution of instantaneous frequencies recorded from WT or
KO cultures. In WT, TTX treatment increases mEPSC frequency compared with
citrate control (control, 1.8 Hz; n ? 22; TTX treated, 2.2 Hz; n ? 20; P ? 0.09).
(P ? 0.55) from KO control (control, 2.5 Hz; n ? 17; TTX treated, 2.7 Hz, n ? 22
neurons). (b) Cumulative distribution of mEPSC amplitudes recorded from
neurons in WT or KO cultures. In WT, TTX treatment increases mEPSC ampli-
tude compared with control (control, 7.0 pA; TTX treated, 7.8 pA; P ? 0.001).
In KO, TTX treatment does not increase mEPSC amplitude (7.2 pA) over either
WT or KO (7.0 pA) controls (P ? 0.62). (c) Action potential activity is normal in
cultures of KO hippocampal neurons as assessed by cell-attached patch re-
0.91 Hz ? 0.25 (n ? 13 neurons).
mEPSCs fail to scale up with activity blockade in KO neurons.
Goddard et al. PNAS ?
April 17, 2007 ?
vol. 104 ?
no. 16 ?
key members of a Ca2?-regulated signaling pathway including
CamKII and BDNF (12, 52, 53).
Although homeostatic scaling occurs in response to the
amount of activity across the entire neuron, Hebbian mecha-
nisms of LTP and LTD act to modulate strength at specific
synapses. Models that explore changes in synaptic strength in a
competitive, Hebbian manner generally need to invoke a ‘‘syn-
aptic normalization,’’ akin to synaptic scaling, to prevent uncon-
trolled positive feedback of potentiation and/or depression (44).
Consequently, neuronal circuits unable to scale synaptic strength
might also be expected to exhibit abnormalities in synaptic
plasticity; this phenotype is observed in ?2m/TAP1 mice, which
have enhanced LTP and lack LTD (19). Together our observa-
tions reveal a dual role for MHCI in both forms of activity-
dependent synaptic plasticity: MHCI, whose expression is mod-
ulated by overall levels of action potentials both in vivo (7) and
in vitro, contributes to homeostatic plasticity, which in turn may
set limits on the magnitude and direction of Hebbian synaptic
More details are found in SI Methods.
Hippocampal Cultures. All animals were treated in accordance with
institutional guidelines. Cultures were derived from postnatal day 0
to postnatal day 1 mouse hippocampi by using standard protocol
(54). Cultures were grown (2–3 weeks) on coverslips over a glial
feeder layer in 12-well plates in 2 ml of Neurobasal media with B27
supplement (Invitrogen, Carlsbad, CA) and 3.7 ?g/ml glutamate
4 days in vitro and again at 10 days to administer drug. Low-density
cultures (Figs. 1 and 2a) were plated at 20,000 cells per 15-mm
coverslip; high-density cultures were plated at 100,000 cells per
as described (19). WT mice were of two types: derived from
production of ?2m/TAP1 KO (19) or from C57BL/6 from Charles
River Breeding Laboratories (Portage, MI). When possible, WT
and KO cultures were prepared on the same day; occasionally, WT
and KO cultures were prepared 3–4 days apart. If derivation of
cultures was separated by 1 day, cultures were treated and fixed
simultaneously. For separations of ?1 day, cultures were treated
and fixed independently.
For activity manipulations, action potentials were blocked by
adding 1 ?M TTX (T5651; Sigma–Aldrich, St. Louis, MO;
supplied in sodium citrate vehicle) to 13- or 14-days in vitro
cultures; vehicle control was 5 ?M sodium citrate (Sigma–
Aldrich). One milliliter of culture media was removed and
replaced with drug/vehicle to a final concentration of 1 ?M for
5 days, refreshing on the third day.
Hippocampal Culture Physiology. High-density cultures (100,000
cells plated) were used after 14 days in vitro. [TTX treatments
(3–6 days) began at day 13 or 14.] Whole-cell recordings were
obtained at room temperature in standard artificial cerebrospi-
nal fluid (in mM): 136 NaCl, 2.5 KCl, 10 glucose, 10 Hepes, 2
CaCl2, 1.3 MgCl2, containing 0.5 ?M TTX, 20 ?M bicuculline,
and 50 ?M AP5. Data were collected with an Axopatch 200B
amplifier and pClamp 9.2. Pipets were pulled by using a Sutter
P-97, with tip resistances of 4–9 M?. Internal solution contained
(in mM): 130 potassium gluconate, 10 NaCl, 1 EGTA, 0.133
CaCl2, 2 MgCl2, 10 Hepes, 3.5 MgATP, and 1 NaGTP.
Cortical Slice Physiology. Coronal brain slices were cut (300-?m
thick) from WT (derived from the generation of ?2m/TAP1 KO
mice) and ?2m/TAP1 KO mice at P19–21 in ice-cold ACSF (in
mM) 130 NaCl, 3 KCl, 1.25 NaH2PO4, 10 glucose, 20 NaHCO3,
1.3 MgSO4, and 2.5 CaCl2) bubbled with 95% 02/5% CO2. Slices
were immediately transferred to 37°C ACSF bath for 30 min and
at 31–32°C in oxygenated ACSF ? 0.25 ?M TTX, 25 ?M APV,
and 10 ?M bicuculline. Pipets were pulled by using a Sutter P-97,
with tip resistances of 4–8 M?. Internal solution consisted of (in
mM) 115 CsMeSO4, 5 NaF, 10 EGTA, 10 Hepes, 15 CsCl, 3.5
MgATP, 3 QX-314, and Lucifer yellow. Slices were kept for up
to 6 h.
MHCI Immunostaining. For all experiments, age-matched WT and
KO cultures were immunostained and imaged together; imaging
and analysis were done blind to genotype and treatment condition.
Cultures were removed from media and immersed in paraformal-
dehyde (4% in PBS) at 37°C for 8 min. Anti-MHCI antibodies
(Ox18, ERHR52) (7, 55) were added in 5% donkey serum (The
Jackson Laboratory, Bar Harbor, ME) in PBS ? 0.1% Tween
(PBST) at 1 ?g/ml. Sister cultures were immunostained with
equimolar concentrations of isotype IgG as control (Ox18, Serotec,
Oxford, U.K.; mouse IgG1, Sigma–Aldrich; M-5284, ERHR52,
Bachem, Bubendorf, Switzerland; Peninsula Laboratories, Bel-
mont, CA; Rat IgG2a, eBiosciences, San Diego, CA; 14–4321).
was added at 1/200 (anti-mouse) or 1/500 (anti-rat) for 1 h.
Avidin-biotin amplification (Vector Laboratories, Burlingame,
CA) was prepared in PBS and applied for 30 min. Cy3-conjugated
tyramide (NEN Life Science Products, Boston, MA) was prepared
in included diluent at 1/100 and added to cultures for 5 min. Then
synapsin immunostaining was performed as described below by
using rabbit anti-synapsin and goat anti-PSD-95 (for Ox18) or
mouse anti-PSD-95 (for ERHR52).
Synaptic Marker Immunostaining.Primaryandsecondaryantibodies
were diluted in 5% donkey serum (The Jackson Laboratory) in
PBST. Rabbit anti-synapsin (AB1543P; Chemicon International,
Bioreagents, Golden, CO), 15 ?g/ml; guinea pig anti-rat vGluT1
(AB5905; Chemicon) and vGluT2 (AB5907; Chemicon), 1/2000
(serum); and mouse anti-?-III-tubulin (MAB1637; Chemicon), 1
?l per culture (concentration not determined by manufacturer).
Donkey Cy2 anti-rabbit and Cy3 anti-mouse (The Jackson Labo-
Probes, Eugene, OR) was used to image tubulin; reagents were
prepared following documentation from Molecular Probes. Cover
slips were mounted onto slides with N-propyl-galate (Sigma–
Aldrich) and sealed with DPX (Electron Microscopy Sciences,
Hatfield, PA). Cultures were imaged on a LSM 510 (Carl Zeiss,
Thornwood, NY) confocal microscopy system; 15–20 images were
collected per cover slip. Images were taken at 1,024 ? 1,024
resolution by using a Plan-Apochromat ?63/1.4 N.A. oil objective
plus ?1.4 software zoom.
EM Methods. Age P44 ?2m/TAP1 KO mice or WT mice (n ? 3
each genotype; WT derived from generation of KO) were
anesthetized with halothane and given a lethal injection of
euthasol. Mice were perfused transcardially with normal 0.9%
saline at 37°C, followed by at least 30 ml of 2.5% glutaraldehyde
and 2% paraformaldehyde in 0.1 M sodium cacodylate buffer at
37°C. Brains were removed and postfixed overnight in the same
fixative at 4°C; vibratome sections were cut (300-?m thick
coronal) to expose hippocampus, and then blocks were cut from
stratum radiatium. Blocks were treated with 1% osmium in
potassium ferrocyanate and uranyl acetate. Thin sections (90
nm) were cut and placed on formvar grids. Grids were treated
with uranyl acetate and Pb acetate. EM images were taken at
?10,000 magnification in stratum radiatum at a distance of
50–100 ?m from stratum pyramidale.
www.pnas.org?cgi?doi?10.1073?pnas.0702023104Goddard et al.
We thank the Harvard Center for Neurodegeneration and Repair for
access to the Confocal Microscopy Core facility. This work was sup-
ported by National Institutes of Health Grant R01 MH071666 and the
Dana Foundation (to C.J.S.), a Goldenson Research Fellowship (to
D.A.B.), and National Institutes of Health Grant T32 MH20017 and a
Victoria and Stuart Quan fellowship (to C.A.G.).
1. Katz LC, Shatz CJ (1996) Science 274:1133–1138.
2. Lein ES, Hohn A, Shatz CJ (2000) J Comp Neurol 420:1–18.
3. Cabelli RJ, Shelton DL, Segal RA, Shatz CJ (1997) Neuron 19:63–76.
4. Lisman J, Schulman H, Cline H (2002) Nat Rev Neurosci 3:175–190.
5. Wong GH, Bartlett PF, Clark-Lewis I, Battye F, Schrader JW (1984) Nature
6. Williams KA, Hart DN, Fabre JW, Morris PJ (1980) Transplantation 29:274–
7. Corriveau RA, Huh GS, Shatz CJ (1998) Neuron 21:505–520.
8. Barco A, Patterson S, Alarcon JM, Gromova P, Mata-Roig M, Morozov A,
Kandel ER (2005) Neuron 48:123–137.
9. Kaltschmidt C, Kaltschmidt B, Baeuerle PA (1995) Proc Natl Acad Sci USA
10. Ishii T, Hirota J, Mombaerts P (2003) Curr Biol 13:394–400.
11. Loconto J, Papes F, Chang E, Stowers L, Jones EP, Takada T, Kumanovics A,
Fischer Lindahl K, Dulac C (2003) Cell 112:607–618.
12. Thiagarajan TC, Piedras-Renteria ES, Tsien RW (2002) Neuron 36:1103–1114.
13. Deisseroth K, Bito H, Schulman H, Tsien RW (1995) Curr Biol 5:1334–1338.
14. Fischer Lindahl K (1997) Immunogenetics 46:53–62.
15. Bijlmakers MJ, Ploegh HL (1993) Curr Opin Immunol 5:21–26.
16. Van Kaer L, Ashton-Rickardt PG, Ploegh HL, Tonegawa S (1992) Cell
17. Zijlstra M, Bix M, Simister NE, Loring JM, Raulet DH, Jaenisch R (1990)
18. Neefjes JJ, Momburg F (1993) Curr Opin Immunol 5:27–34.
19. Huh GS, Boulanger LM, Du H, Riquelme PA, Brotz TM, Shatz CJ (2000)
20. Upton AL, Salichon N, Lebrand C, Ravary A, Blakely R, Seif I, Gaspar P
(1999) J Neurosci 19:7007–7024.
21. Godement P, Salaun J, Imbert M (1984) J Comp Neurol 230:552–575.
22. Stellwagen D, Shatz CJ (2002) Neuron 33:357–367.
23. Syken J, Grandpre T, Kanold PO, Shatz CJ (2006) Science 313:1795–1800.
24. Boulanger LM, Shatz CJ (2004) Nat Rev Neurosci 5:521–531.
25. Neumann H, Schmidt H, Cavalie A, Jenne D, Wekerle H (1997) J Exp Med
26. Kennedy MB (1997) Trends Neurosci 20:264–268.
27. Greengard P, Browning MD, McGuinness TL, Llinas R (1987) Adv Exp Med
28. Zhong J, Zhang T, Bloch LM (2006) BMC Neurosci 7:17–29.
29. Linda H, Hammarberg H, Cullheim S, Levinovitz A, Khademi M, Olsson T
(1998) Exp Neurol 150:282–295.
30. El-Husseini AE, Schnell E, Chetkovich DM, Nicoll RA, Bredt DS (2000)
31. Hopf FW, Waters J, Mehta S, Smith SJ (2002) J Neurosci 22:775–781.
32. Pieribone VA, Shupliakov O, Brodin L, Hilfiker-Rothenfluh S, Czernik AJ,
Greengard P (1995) Nature 375:493–497.
33. Ploegh HL, Cannon LE, Strominger JL (1979) Proc Natl Acad Sci USA
34. Fremeau RT, Jr, Kam K, Qureshi T, Johnson J, Copenhagen DR, Storm-
Mathisen J, Chaudhry FA, Nicoll RA, Edwards RH (2004) Science 304:1815–
35. Fremeau RT, Jr, Voglmaier S, Seal RP, Edwards RH (2004) Trends Neurosci
36. Kim E, Sheng M (2004) Nat Rev Neurosci 5:771–781.
37. Nakagawa T, Futai K, Lashuel HA, Lo I, Okamoto K, Walz T, Hayashi Y,
Sheng M (2004) Neuron 44:453–467.
38. Burrone J, O’Byrne M, Murthy VN (2002) Nature 420:414–418.
39. Nakayama K, Kiyosue K, Taguchi T (2005) J Neurosci 25:4040–4051.
40. Murthy VN, Schikorski T, Stevens CF, Zhu Y (2001) Neuron 32:673–682.
41. De Gois S, Schafer MK, Defamie N, Chen C, Ricci A, Weihe E, Varoqui H,
Erickson JD (2005) J Neurosci 25:7121–7133.
42. Beattie EC, Stellwagen D, Morishita W, Bresnahan JC, Ha BK, Von Zastrow
M, Beattie MS, Malenka RC (2002) Science 295:2282–2285.
43. Burrone J, Murthy VN (2003) Curr Opin Neurobiol 13:560–567.
44. Turrigiano GG (1999) Trends Neurosci 22:221–227.
45. Turrigiano GG, Leslie KR, Desai NS, Rutherford LC, Nelson SB (1998) Nature
46. Malenka RC, Nicoll RA (1997) Neuron 19:473–476.
47. Wierenga CJ, Walsh MF, Turrigiano GG (2006) J Neurophysiol 96:2127–2133.
48. Miller GL, Knudsen EI (2001) J Neurophysiol 85:2184–2194.
49. Bacci A, Huguenard JR (2006) Neuron 49:119–130.
50. Hartman KN, Pal SK, Burrone J, Murthy VN (2006) Nat Neurosci.
51. Stellwagen D, Malenka RC (2006) Nature 440:1054–1059.
52. Tyler WJ, Pozzo-Miller LD (2001) J Neurosci 21:4249–4258.
53. Rutherford LC, Nelson SB, Turrigiano GG (1998) Neuron 21:521–530.
54. Banker G, Goslin K (1998) (MIT Press, Cambridge, MA).
55. Neumann H, Cavalie A, Jenne DE, Wekerle H (1995) Science 269:549–552.
Goddard et al. PNAS ?
April 17, 2007 ?
vol. 104 ?
no. 16 ?