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NEUROSCIENCE
Astrocyte dysfunction increases cortical
dendritic excitability and promotes cranial pain
in familial migraine
Jennifer Romanos1,2, Dietmar Benke1,2, Daniela Pietrobon3,4,
Hanns Ulrich Zeilhofer1,2,5, Mirko Santello1,2*
Astrocytes are essential contributors to neuronal function. As a consequence, disturbed astrocyte-neuron interactions
are involved in the pathophysiology of several neurological disorders, with a strong impact on brain circuits and
behavior. Here, we describe altered cortical physiology in a genetic mouse model of familial hemiplegic migraine
type 2 (FHM2), with reduced expression of astrocytic Na+,K+-ATPases. We used whole-cell electrophysiology,
two-photon microscopy, and astrocyte gene rescue to demonstrate that an impairment in astrocytic glutamate
uptake promotes NMDA spike generation in dendrites of cingulate cortex pyramidal neurons and enhances output
firing of these neurons. Astrocyte compensation of the defective ATPase in the cingulate cortex rescued glutamate
uptake, prevented abnormal NMDA spikes, and reduced sensitivity to cranial pain triggers. Together, our results
demonstrate that impaired astrocyte function alters neuronal activity in the cingulate cortex and facilitates
migraine-like cranial pain states in a mouse model of migraine.
INTRODUCTION
Astrocytes closely interact with neurons and strongly affect their
functions at the synaptic, cellular, and circuit levels (1). Defective
neuron-astrocyte interactions have been implicated in the establish-
ment and development of several neurological disorders (1–3).
Migraine is an extremely debilitating disease characterized by re-
current unilateral and severe headaches, frequently accompanied
with several other neurological symptoms (4). However, migraine is
much more than an episodic pain disorder. Several findings indeed
suggest that migraine is a disease affecting a large part of the central
nervous system and characterized by a global dysfunction in sensory
information processing and integration, which also occurs between
migraine episodes (interictal period) (5). For example, patients with
migraine exhibit increased cortical responses to noxious and non-
noxious sensory stimuli during the interictal period (6,7). At present,
the cellular mechanisms responsible for these alterations are largely
unknown. Astrocytes have been proposed to play a role in some
inherited forms of migraine, including familial hemiplegic migraine
type 2 (FHM2), an autosomal dominant form of migraine with aura.
FHM2 is caused by mutations in the Atp1a2 gene, which encodes
the 2 subunit of the Na+, K+-dependent adenosine triphosphatase
(ATPase) (2 NKA) (8), an isoform that is almost exclusively ex-
pressed in astrocytes in the adult brain (9). Expression of the 2
NKA is reduced in the heterozygous Atp1a2+/R887 FHM2 knock-in
(KI) mice, which carry an Atp1a2 missense mutation, causing a
complete loss of function of recombinant 2 NKA (8,10). We pre-
viously showed that these FHM2 mice display impaired glutamate
and K+ clearance by astrocytes of the primary somatosensory cortex,
which, in turn, promotes cortical spreading depression (CSD), the
neuronal correlate of the aura symptoms that precede migraine
headache (3).
In this study, we took advantage of the FHM2 KI mouse model
to understand whether and how a mutation in an astrocyte-specific
protein affects neuronal functions in the cingulate cortex (Cg). This
cortical region is crucial for pain processing and displays altered
functionality in patients with migraine (7,11–13). We show that
impairment in astrocytic glutamate uptake in this region strongly
enhances cortical dendritic excitability, especially the generation
of N-methyl-d-aspartate (NMDA) spikes in layer 5 (L5) pyramidal
neurons, and enhances their output firing. Moreover, we reveal that
FHM2 mice display increased sensitivity to head pain triggers. Last,
we show that rescuing the disease-causing mutation in astrocytes of
the Cg recovers neuronal function and reduces the pain phenotype.
Our results provide a clear example of how astrocyte dysfunction
produced by a genetic defect affects neuronal activity in the Cg and
affects sensitivity to head pain triggers.
RESULTS
Astrocytic dysfunctions in the Cg of FHM2 mice
The 2 NKA is physically and functionally coupled to glutamate trans-
porters (GluTs) expressed on perisynaptic astrocyte processes (9,14).
The reduced expression of 2 NKA in heterozygous Atp1a2+/R887
mice (FHM2 mice) results in a reduction in glutamate and K+ buffering
capacity of astrocytes in the primary somatosensory cortex (3,10).
To investigate FHM2-associated alterations in glutamate and K+
uptake in the Cg, a brain region crucially involved in pain process-
ing, we performed whole-cell patch-clamp recordings from astro-
cytes in acute cortical slices (Fig.1A). We recorded synaptically evoked
GluT currents (STCs) and K+ uptake currents induced by electrical
stimuli to L1 afferent fibers with an extracellular electrode and
applied different stimulation paradigms (single pulses and pulse
trains at 50 and 100 Hz; Fig.1,AandB). Glutamate uptake was
significantly slower in the Cg of FHM2 mice compared to their
wild-type (WT) littermates, which was reflected by higher STC decay
time constants () [Fig.1,CandD; single pulse: WT decay=3.11±0.15 ms
1Institute of Pharmacology and Toxicology, University of Zurich, CH-8057 Zurich,
Switzerland. 2Neuroscience Center Zurich, University of Zurich and ETH Zurich, CH-8057
Zurich, Switzerland. 3Department of Biomedical Sciences and Padova Neuroscience
Center, University of Padova, 35131 Padova, Italy. 4CNR Institute of Neuroscience,
Via Ugo Bassi 58/B, 35131 Padova, Italy. 5Institute of Pharmaceutical Sciences, ETH
Zurich, CH-8093 Zurich, Switzerland.
*Corresponding author. Email: mirko.santello@pharma.uzh.ch
Copyright © 2020
The Authors, some
rights reserved;
exclusive licensee
American Association
for the Advancement
of Science. No claim to
original U.S. Government
Works. Distributed
under a Creative
Commons Attribution
License 4.0 (CC BY).
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(n=10 cells, N=5 mice) and FHM2 decay=3.69±0.15 ms (n=14,
N=8; *P=0.017); 50 Hz: WT decay=2.81±0.11 ms (n=10) and
FHM2 decay=3.34±0.18 ms (n=13; *P=0.029); 100 Hz: WT
decay=2.54± 0.13 ms (n=10) and FHM2 decay=3.22±0.18 ms
(n=13; **P=0.009)]. The decay kinetics of the K+ currents following
a train of pulses at 100Hz were also significantly slower in the FHM2
mice [Fig.1E; WT decay=1.72±0.08 s (n=7 cells, N=4 mice) and
FHM2 decay=2.25± 0.18 s (n=10, N=5 mice; *P =0.03)]. We
observed no difference in astrocytic resting membrane potential
nor input resistance between the two groups (fig. S1). These data
indicate that in the Cg of adult FHM2 mice, astrocytic uptake of
neuron-derived glutamate and K+ was impaired.
Slower glutamate clearance by astrocytes may lead to prolonged
and increased glutamate levels in the extracellular space. To directly
= 3.49 ms
= 3.71 ms
= 3.81 ms
Single pulse
10 ms
20 pA
= 3.5 ms
Train of pulses
at 50 Hz or 100 Hz
STC (ms)
*P = 0.029 **P = 0.009
*P = 0.01
7
STC (ms)
STC (ms)
K+(s)
*P = 0.03
1
ROI
Stimulation
GFAP.iGluSnFR SR-101
L1 L2/3
GFAP.iGluSnFR: Extrasynaptic glutamate
Decay (ms)
**P = 0.002
A
WT
100 Hz
= 3.53 ms
= 2.75 ms
50 Hz
= 2.81 ms
WT
WT
0
1
2
3
0
2
4
6
0
2
4
6
0
2
4
6
WT
FHM2
WT
WT
STC 11th pulse STC 11th pulse 100 Hz
B
G
E
WT
FHM2
Synaptically activated
glutamate transporter currents (STC)
D
WT
FHM2
AAV.GFAP.iGluSnFr
WT or FHM2 KI mice
F
Unilateral in cingulate cortex
10 × 50 Hz 10 × 100 Hz
C
0
50
100
150
0
50
100
150 **P = 0.006
FHM2
FHM2
FHM2
FHM2
FHM
2
Single pulse
200 ms
Decay (ms)
10 ms
20 pA
From FHM2 KI (FHM2) mice
and WT littermates
SR-101
Cingulate
cortex
Stim.
electrode
Patch pipette
Layer 1
Fig. 1. Aberrant astrocytic glutamate and K+ uptake in the Cg of FHM2 mice. (A) Schematic representation of the experiment. Scale bar, 30 m. (B) Superimposed
representative traces of the inward current evoked in an astrocyte with different stimulation patterns. (C) The decay time of inward currents evoked by single-pulse stim-
ulation is slower in FHM2 mice (red) compared to WT mice (black). (D) The average STC decay times of the last pulse of the trains at 50 (left) and 100 Hz (right) are signifi-
cantly slower in FHM2 mice. Each point represents the STC decay time in one astrocyte. (E) Decay kinetics of the K+ inward current following trains of 100 Hz stimulation
are slower in FHM2 mice. (F) Injection of AAV.GFAP.iGluSnFr unilaterally in the Cg of WT and FHM2 mice. A typical two-photon experiment showing the expression of
iGluSnFr on astrocytes in the Cg (green) and sulforhodamine 101 dye (SR-101; red) is shown. The theta glass electrode for synaptic stimulation is placed in the inner L1,
and glutamate is imaged from an ROI adjacent to the electrode. Scale bar, 40 m. (G) Upon trains of synaptic stimulation (10 × 50 Hz and 10 × 100 Hz), robust and reliable
increases in iGluSnFr emission could be detected. The decay kinetics of the averaged transients are slower in FHM2 mice. Representative traces are the average of at least
five sweeps. Data are means ± SEM. Two-tailed unpaired t test was used.
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test this prediction, we took advantage of the intensity-based glutamate
fluorescent sensor iGluSnFr (15) that we expressed on the extracellular
side of the astrocytic plasma membrane (Fig.1F and see Materials
and Methods). Two-photon imaging of iGluSnFr glutamate signals
allowed the study of the time course of extracellular glutamate follow-
ing synaptic stimulation. Synaptic activity was evoked by focal electrical
stimulation in L1 of the Cg, and glutamate was imaged in a region of
interest (ROI) in the proximity of the stimulation electrode (Fig.1F).
To estimate the speed of glutamate clearance, we fitted the decay of
the averaged evoked glutamate transients (10 trials) with a monoex-
ponential curve (16). We found that synaptically evoked iGluSnFr
transients in the adult Cg were sensitive to minor changes in the
activity of astrocytic GluTs. Accordingly, partial blockade of GluTs
with subsaturating concentrations of threo-beta-Benzyloxyaspartate
(DL-TBOA) (3 M), a concentration that mimics in WT mice the
slowing of STC decay kinetics produced by the FHM2 mutation (3),
increased the decay time constant of extracellular glutamate transients
in WT mice (fig. S2).
We subsequently recorded iGluSnFr signals in the Cg of FHM2
mice and compared it with WT littermates. The iGluSnFr signals
displayed a longer time course following trains of 50 and 100Hz
stimulations in the FHM2 mice, which were 32 and 29% slower,
respectively [Fig.1G; 50 Hz: WT decay= 75.56±3.62 ms (n =13
slices, N=5 mice) and FHM2 decay=100.9±5.33 ms (n=22, N=7;
**P=0.0019); 100 Hz: WT decay = 65.6±3.32 ms (n=14) and FHM2
decay=84.57 ± 4.77 ms (n=22; **P= 0.006)]. Note that we and
others have previously demonstrated that the size and location of
ROIs, stimulation intensity, and sulforhodamine 101 dye (SR-101)
do not influence the iGluSnFr decay kinetics (16,17). Overall, these
experiments demonstrate that in the adult Cg, astrocytes carrying
the W887R 2 NKA mutation responsible for FHM2 display defective
glutamate and K+ buffering capacity upon repetitive synaptic stim-
ulation, which results in temporally prolonged glutamate spillover.
Facilitation of NMDA spike generation in L5 pyramidal
neurons of FHM2 mice
Pharmacological reduction of astrocytic glutamate uptake capacity
in the cortex increases extracellular buildup of glutamate, which
directly affects NMDA receptor activation in L5 pyramidal neurons
(17). In addition, glutamate spillover promotes the occurrence of
NMDA dendritic spikes (18). These spikes are local events (de-
polarizations) caused by the regenerative and voltage-dependent
activation of NMDA receptors in specific dendritic branches and
have been shown to strongly promote pyramidal cell firing invivo
(19). We tested whether the alteration of astrocytic glutamate clearance
in FHM2 mice would affect NMDA spike generation. We evoked
NMDA spikes in the distal dendrites of L5 pyramidal neurons by
focal synaptic stimulation (paired pulse, 50 Hz) in close proximity
to single branches of tuft dendrites in L1 of the Cg (Fig.2A). In-
creasing stimulation intensities caused a nonlinear increase in the
amplitude and area under the curve (AUC) of the second pulse,
which is characteristic of NMDA spikes (20). This nonlinear increase
was abolished in the presence of NMDA receptor blocker d,l-2-
amino-5-phosphonovaleric acid (AP-V) (Fig.2B).
As predicted from the reduced glutamate clearance in FHM2 mice,
these mice displayed facilitated NMDA spike generation reflected
by higher amplitude and AUC of the voltage responses evoked by
L1 synaptic stimulations [Fig.2C; second pulse amplitude: WT,
1.56±0.4 mV (n=12 cells, N=8 mice) and FHM2, 9.13±2.48 mV (n=13,
N=6; ***P<0.0001); second pulse AUC: WT, 40.64±13.9 mV ms
(n=11) and FHM2, 411.5±131 mV ms (n=12; *P<0.05); stimula-
tion intensity, 2 mA]. Partial blockade of GluTs with subsaturating
concentrations of the GluT blocker dl-TBOA (3 M) significantly
enhanced NMDA spike generation in L5 pyramidal neurons in WT
mice (fig. S3), mimicking the effect of the FHM2 genetic mutation.
Dendritic NMDA spikes have been reported to promote somatic
firing in neurons of the somatosensory cortex (19). The main apical
dendrite of L5 pyramidal neurons of the Cg displays a remarkably
low dendrite-to-soma attenuation of slow synaptic inputs compared to
other cortical regions (21), suggesting that NMDA-mediated dendritic
depolarizations may have an even stronger influence on somatic depo-
larization and action potential (AP) firing in this region. We find that
our NMDA spike induction protocol easily triggered somatic firing
(Fig.2D). Consequently, we predicted that the facilitation of NMDA
spikes that we identified in FHM2 Cg L5 pyramidal neurons should
lead to increased firing of these neurons. We found that the probability
of AP firing following NMDA spikes was significantly higher in FHM2
mice compared to their WT littermates (Fig.2D). These data demon-
strate that in the Cg, defective glutamate uptake and the prolonged
presence of synaptically released glutamate are accompanied by a fa-
cilitation in NMDA spikes and somatic firing of L5 pyramidal cells.
We then investigated whether synaptic activity is altered in FHM2
mice. We first recorded miniature excitatory postsynaptic currents
(mEPSCs) from L5 pyramidal cells of WT and FHM2 mice
(Fig.3,A, B, and C) and found no change in baseline synaptic activity
(mEPSC frequency and amplitude) between the two groups [mEPSC
frequency: WT, 3.02± 0.71Hz (n =7 cells) and FHM2 KI,
4.53±1.54Hz (n=9 cells; P=0.43); mEPSC amplitude: WT,
14.81±1.35 pA and FHM2 KI, 13.09±0.82 pA (P=0.27)]. We
additionally recorded AMPA-mediated EPSCs evoked with single-
pulse and paired-pulse extracellular synaptic stimulations (Fig.3,D and E).
Upon single-pulse stimulations, both the amplitude and decay
kinetics of EPSCs were similar in WT and FHM2 mice [amplitude:
WT, −117±18.04 pA (n=7 cells) and FHM2 KI, −118.6±12.03 pA
(n= 6 cells; P=0.94); decay: WT, 17.7 ±2.1 ms (n=7 cells) and
FHM2 KI, 18.29±1.4 ms (n=6 cells; P=0.82)]. Similarly, upon
paired-pulse stimulation at 20 Hz, the second-over-first EPSC
amplitude ratio was comparable in WT and FHM2 mice [second/
first amplitude ratio: WT, 1.73±0.09 (n=7 cells) and FHM2,
1.54±0.08 (n=7 cells; P=0.17)]. These datasets strongly argue
against alterations in glutamate release or in AMPA-mediated
synaptic transmission in the Cg of FHM2 mice.
Local astrocytic defects are responsible for NMDA-mediated
neuronal dysfunctions
Excitatory neuronal activity appears to be highly increased in the
Cg of FHM2 mice. Nevertheless, to what extent the local astrocytic
malfunction is responsible for the modifications in neuronal activity
is not clear. To address this question, we compensated for the astro-
cyte dysfunction in the Cg region of FHM2 mice by expressing the
WT form of 2 NKA (Atp1a2) in astrocytes. To deliver Atp1a2,
we used an adeno-associated virus (AAV) of the 5/2 serotype that
preferentially targets astrocytes and took advantage of the astrocyte
promoter hGFAP (AAV.hGFAP.ATP1A2) (22). As a control, AAV
of the same serotype and with the same vector backbone, but
containing only enhanced green fluorescent protein (eGFP)
(AAV.hGFAP.eGFP), was injected (Fig.4A; also see Materials and
Methods). Since AAV vectors’ size is limited to ~4.7 kb, it was not po ssib le
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to include a reporter gene in the virus with ATP1A2. Therefore, to visual ize
the injection site, we injected a mixture of 0.5l of the two viruses
with a ratio of 2:1 of AAV.hGFAP.ATP1A2 to the control virus
(AAV.hGFAP.eGFP). Two to 3 weeks following injections, immuno-
histochemistry experiments showed that AAV.hGFAP targeted
astrocytes with no apparent neuronal expression (fig. S4). Consistent
with previous findings on other brain regions (10), the Cg of FHM2
mice showed a substantial reduction of 2 NKA expression levels
(about 70%) compared to WT littermates (fig. S5A). This reduction
was significantly, albeit not fully, restored in FHM2 mice injected
with the rescue virus (fig. S5A). The Western blot experiments were
performed 15 days post-injection (d.p.i.) of the viruses. Accordingly,
the functional recovery of the STC decay kinetics was also incomplete
at 15 d.p.i. (fig. S6). This may, at least in part, account for the incomplete
restoration of 2 NKA expression. For this reason, the functional and
behavioral experiments (see later) were performed mostly at 21 d.p.i., wh en
the STC decay kinetics became similar to WT levels (fig. S6). In addition,
it is also likely that the Cg tissue extracted for Western blot analysis con-
tained regions with a mixture of high and low (or no) virus expression.
We additionally evaluated the expression levels of the astrocytic
GluTs, GLT-1 and GLAST, that both play a crucial role in glutamate
uptake in the Cg (17). We observed a 25% reduction of GLAST ex-
pression in the Cg of FHM2 mice, which was restored to WT levels in
FHM2 mice injected with the rescue virus (fig. S5B). On the other
hand, no changes in GLT-1 expression levels were observed between
the different groups (fig. S5C).
We then performed electrophysiological recordings in the same
manner as in Fig.1 from FHM2 astrocytes either expressing WT Atp1a2
or the eGFP control. Our data show that both K+ [Fig.4B; control decay=
2.66±0.16 s (n=9 cells, N=4 mice) and rescue decay=2.14±0.13 s
(n=10, N=4; *P=0.02)] and active glutamate clearance by astrocytes
became significantly faster in FHM2 mice in which WT Atp1a2 was
expressed compared to FHM2 mice that were injected with the control
virus [Fig.4C; single pulse: control decay=3.3±0.06 ms (n=10 cells,
N= 4 mice) and rescue decay= 2.6±0.13 ms (n=12, N= 4;
***P=0.0003); 50 Hz: control decay = 3.24±0.06 ms (n=10) and
rescue decay=2.46±0.18 ms (n=12, **P=0.001); 100 Hz: control
decay = 3.11±0.04 ms (n=10) and rescue decay =2.31± 0.12 ms
(n=12; ***P=0.0001)]. The decay kinetics of the uptake currents in the
rescued FHM2 mice became comparable to those observed in WT mice
(fig. S7A). The astrocytic resting membrane potential was slightly
but significantly hyperpolarized in rescued FHM2 mice compared
to those injected with control virus, with no difference in input
resistance (fig. S8).
AP-V
2nd pulse amplitude (mV)
Stim. intensity (mA)
***
*
Stim. intensity (mA)
2nd pulse area (mV.ms)
*
**
Stim. intensity (mA)
Probability of AP ring
3.5
2 mA
2.5
3
4
–70 mV –72 mV
WT FHM2
50 ms
10 mV
*
***
WT
FHM2
5 mV
20 ms
–73 mV
–72 mV
WT
FHM2
BC
D
WT
FHM2
1.0
1.5
2.0
2.5
0
500
1000
1.0
1.5
2.0
2.5
0
5
10
15
1
1.5 2
2.5 3
3.5 4
4.5 5
5.5
0.0
0.5
1.0
5 mV
20 ms
A
Stimulation
L1
L2-L3
L5
Biocytin
Fig. 2. Facilitation of NMDA spike generation in L5 pyramidal neurons in the Cg of FHM2 mice. (A) Image of whole-cell recording from the soma of a biocytin-labeled
L5 pyramidal cell in the Cg showing the location of the recording pipette (L5) and stimulation electrode (in close proximity to single branches of tuft dendrites in L1 of the
Cg). Scale bar, 50 m. (B) Representative traces of NMDA spikes evoked by focal synaptic stimulation (paired pulse, 50 Hz) of increasing stimulation intensities that cause
an abrupt and nonlinear increase in amplitude and AUC of the second pulse, which is characteristic of NMDA spikes. This nonlinear increase is abolished in the presence
of NMDA receptor blocker AP-V (50 M; box). (C) The amplitude and the AUC of the second pulse are significantly higher in FHM2 mice compared to WT mice upon lower
stimulation intensities. (D) Paired-pulse stimulation had a higher probability to evoke a somatic action potential (AP) in FHM2 mice. Data are means ± SEM. Two-way
analysis of variance (ANOVA) with Bonferroni post hoc test and Z score were used.
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In light of these results, we investigated whether the rescue of the
astrocytic dysfunctions could indeed affect the neuronal defects
observed in FHM2 mice. To this end, we evoked NMDA spikes in the
distal dendrites of L5 pyramidal neurons by focal synaptic stimula-
tion as in previous experiments (Fig.4,DandE). The amplitude and
AUC of NMDA spikes were significantly lower in FHM2 mice in-
jected with WT Atp1a2 compared to FHM2 mice injected with the
control virus [Fig.4F; second pulse amplitude: control, 12.56±2.11 mV
(n=7 cells, N=3 mice) and rescue, 4.8±1.52 mV (n=11, N=4;
***P<0.0001); second pulse AUC: control, 432.7±87.7 mV ms
(n=7) and FHM2, 198.15±69 mV ms (n=12; **P<0.01); stimu-
lation intensity, 2.5 mA]. The values in the rescued FHM2 mice were
comparable to those in WT mice (fig. S7B). This rescue was also
accompanied by a significantly lower probability of AP firing follow-
ing NMDA spikes in the rescued FHM2 mice (Fig.4G).
Local astrocyte dysfunction in the Cg influences orofacial
pain in FHM2 mice
The Cg is a critical cortical region in encoding cephalic pain. Altered
neuronal activity in this brain area has been reported to influence
the activation and sensitization of pain pathways in pathological pain
conditions (23). Whether local astrocyte dysfunction in the Cg can
10 pA
P = 0.43 P = 0.27
Wild type
FHM2 KI
mEPSCs
0
500
1000
1500
2000
2500
0
50
100
150
IEI (ms)
Cumulative frequency (%)
0
20
40
60
0
50
100
150
Amplitude (pA)
0
5
10
15
20
25
Amplitude (pA)
WT
0
5
10
15
20
Frequency (Hz)
WT
150 ms
25 pA
10 ms
WT
KI
WT
KI
Amplitude (pA)
–250
–200
–150
–100
–50
0
Decay time (ms)
0
10
20
30
0.0
0.5
1.0
1.5
2.0
2.5
2nd/1st EPSC
amplitude ratio
P = 0.17
FHM2
WT
FHM2
WT
FHM2
WT
WT FHM2
P = 0.94 P = 0.82
20 ms
50 pA
AMPA-mediated evoked EPSCs
FHM2
FHM2
Cumulative frequency (%)
20 pA
10 ms
WT
Single pulse
FHM2
Paired pulse at 20 Hz
A
B
C
D
E
Fig. 3. Baseline synaptic activity is similar in L5 pyramidal cells of FHM2 and WT mice. (A) Representative example traces (6 s) of whole-cell mEPSCs recordings from
WT mice (left) and FHM2 KI mice (right). Average mEPSCs are shown above the traces. (B) Cumulative frequency plots of interevent intervals (IEI) (left) from WT and FHM2
KI and cumulative frequency plot of mEPSCs amplitudes from the same cells (right) calculated from the presented traces above. (C) No difference is observed in mEPSCs
frequency between WT and FHM2 KI slices. Data points represent individual cells. (D) Left: Example traces of AMPA-mediated evoked EPSCs following a single pulse in WT
(black) and FHM2 mice (red). Right: The amplitude of EPSCs following a single pulse is similar in WT and FHM2 mice. Decay kinetics are also similar in WT and FHM2 KI mice.
(E) Left: Example traces of AMPA-mediated evoked EPSCs following a paired-pulse stimulation at 20 Hz in WT and FHM2 mice. Right: The second-to-first EPSC amplitude
ratio is not different in WT compared to FHM2 mice. Data are means ± SEM. Two-tailed unpaired t test was used.
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100 pA
8 ms
0
1
2
3
4
0
1
2
3
4
0
1
2
3
4
= 3.04 ms
STC (ms)
STC (ms)
STC (ms)
AAV.ATP1A2
100 Hz
AAV.eGFP
K+ (s)
0
1
2
3
4*P = 0.02
***P = 0.0001
**P = 0.001
***P = 0.0003
Single pulse 50 Hz 100 Hz
= 3.3 ms
= 2.12 ms
= 3.22 ms
= 1.93 ms = 1.91 ms
STC 11th pulse STC 11th pulse
AAV.ATP1A2
AAV.eGFP
Stim. intensity (mA)
Stim. intensity (mA)
**
***
*
*******
Probability of AP ring
Stim. intensity (mA)
Stim.
electrode
Patch pipette
Biocytin AAV.GFAP.eGFP
L1 L2
AAV5/2.hGFAP.eGFP (control)
or
AAV5/2.hGFAP. ATP1A2 (rescue)
FHM2 KI mice
21 d.p.i.
AAV.eGFP
AAV.ATP1A2
AB
C
DE F
G
Unilateral in cingulate cortex
AAV.eGFP
AAV.eGFP
AAV.ATP1A2
AAV.eGFP
AAV.ATP1A2
AAV.ATP1A2
1.0
1.5
2.0
2.5
0
5
10
15
20
1.0
1.5
2.0
2.5
0
200
400
600
2nd pulse amplitude (mV)
2nd pulse area (mV.ms)
0.0
0.5
1.0
1
1.5 2
2.5 3
3.5 4
4.5 5
5.5
20 mV
20 ms
1.5 mA
2.5
3.5
3
2
5 mV
10 ms
1 s
AAV.eGFP
AAV.ATP1A2
L1
L2-L3
L5
Stimulation
AAV.eGFP
AAV.ATP1A2
Fig. 4. Local astrocytic defects are responsible for NMDA-mediated neuronal dysfunctions. (A) AAVs containing the WT form of Atp1a2 (AAV5/2.hGFAP.ATP1A2) or
a control virus (AAV5/2.hGFAP.eGFP) is unilaterally injected in the Cg of FHM2 mice. At 21 d.p.i., astrocytes expressing either GFP or Atp1a2 in L1 of the Cg cortex are
targeted for recording in acute brain slices while stimulating nearby neurons. Scale bar, 40 m. (B) Example traces of the K+ current following trains of 100 Hz stimulations
in FHM2 mice injected with the control virus (red) and those injected with the rescue virus (blue). The decay kinetics of K+ currents are significantly faster in rescued FHM2
mice compared to control FHM2 mice. (C) Upon all stimulation patterns (single pulse and trains of 50 and 100 Hz), the decay kinetics of STCs are faster in rescued mice
compared to the control group. (D) Image of whole-cell recording from the soma of a biocytin-labeled L5 pyramidal cell in the Cg surrounded by eGFP-expressing astro-
cytes. Scale bar, 40 m. (E) Representative traces of NMDA spikes evoked by focal synaptic stimulation (paired pulse, 50 Hz) in control FHM2 mice (red) and in rescued
FHM2 mice (blue). (F) Both the amplitude and the AUC of the second pulse are significantly lower in rescued FHM2 mice compared to the control group. (G) Paired-pulse
stimulation had a lower probability to evoke a somatic AP in rescued FHM2 mice (blue) compared to control mice (red). Data are means ± SEM. Two-tailed unpaired t test,
two-way ANOVA with Bonferroni post hoc test, and Z score were used.
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facilitate cranial pain in FHM2 mice is unknown. To explore this
possibility, we activated the cranial pain pathway via a single systemic
injection of the nitric oxide donor nitroglycerin (NTG). NTG is con-
sidered a reliable cranial pain trigger especially in migraine-susceptible
patients. Only in patients with migraine, NTG induces a delayed
migraine-like headache with associated features (e.g., premonitory
symptoms) that resemble their own spontaneous migraine attacks
(24). Systemic injections of NO donors have also been used in rodents
(25) and have been shown to evoke hypersensitivity to touch (typical
migraine symptom), particularly in mice carrying a mutation associated
with migraine with aura (26). To assess the development of facial
mechanical sensitization upon NTG injections, we gently poked the
mice in the orofacial region with von Frey filaments and scored the
evoked nocifensive behavior (orofacial pain score; see Materials and
Methods). This scoring system has been previously used to assess
trigeminal neuropathic pain in rodent models (27). We first found
that NTG [10mg kg−1, intraperitoneally (i.p.)] triggered facial
mechanical hypersensitivity in both WT and FHM2 mice at 30, 60, and
120min post NTG injection compared to mice injected with saline
[Fig.5, Band C; AUC for WT: saline, 67.56± 12.50 (N=6 mice)
and NTG, 165.5±19.08 (N=6; **P=0.0016); AUC for FHM2:
saline, 71.32± 11.72 (N=6) and NTG, 190.3±23.92 (N =6;
**P=0.0012)]. FHM2 mice developed orofacial hypersensitivity upon
NTG doses that were ineffective in WT littermates [5mg kg−1;
Fig.5,DandE; AUC for WT: saline, 91.51±19.11 (N=6 mice) and
NTG, 58.90± 8.88 (N=8; P=0.12); AUC for FHM2: saline,
67.16±20.29 (N=7) and NTG, 146.0±13.13 (N=7; **P=0.0068)].
These results suggest that the FHM2 mutation promotes the develop-
ment of NTG-induced facial mechanical hypersensitivity. The
same phenotype was observed with a lighter von Frey filament of
0.025 g (fig. S9). Activity tests showed no difference between
saline- and NTG-treated mice or between FHM2 and WT litter-
mates, suggesting that the different responses to mechanical
stimulation were not accompanied by alterations in motor func-
tion (fig. S10, A and B).
Since the Cg is implicated in pain signaling and development of
mechanical hypersensitivity in several pathological pain syndromes,
we wondered whether rescuing local astrocyte dysfunction in this
brain area could ameliorate the increased nocifensive responses
detected in FHM2 mice upon NTG treatment (23). To that end, we
expressed WT Atp1a2 (AAV.hGFAP.ATP1A2) in Cg astrocytes
of FHM2 mice. As a control, FHM2 mice were injected with AAV.
hGFAP.eGFP. Three weeks after viral injections, we triggered facial
mechanical sensitivity by NTG injections (5mg kg−1) and per-
formed facial von Frey tests as described above (Fig.5,FandG).
We found that rescuing the astrocytic loss-of-function FHM2
mutation locally in the Cg was sufficient to attenuate the acute
hypersensitive phenotype induced by NTG in FHM2 mice (Fig.5H
and fig. S9), suggesting that the local astrocyte dysfunction in
the Cg is implicated in orofacial pain sensitivity [AUC: control,
148.8±9.62 (N=14 mice) and rescue, 91.40±13.98 (N=13 mice;
**P=0.0021)].
Together, our results demonstrate that astrocytes in a genetic
migraine model display altered glutamate and K+ clearance in
the Cg, which facilitate neuronal NMDA spike generation and
synaptically evoked AP firing. We report that these alterations are
pathologically relevant since local rescue of the astrocytic dysfunc-
tions reduces facial hypersensitivity induced by a migraine-relevant
pain trigger.
Inj. Inj. **P = 0.001
AUC
Saline
NTG (10 mg kg−1)
Wild type FHM2 KI mice
A
B
NTG (10 mg kg−1)
Saline
AUC
AUC
Saline
NTG (5 mg kg−1)
Inj. Inj.
**P = 0.006
P = 0.118
***
***
***
*
D
C
Wild type
AUC
**P
= 0.002
***
E
F
Time (min)
0
50
100
150
200
250
BL1
BL2
30
60
120
180
0
2
4
6
Orofacial pain score
Time (min)
Orofacial pain score
BL1
BL2
30
60
120
180
0
2
4
6
***
*
AUC
Saline
NTG (10 mg kg−1)
NTG (10 mg kg−1)
NTG (5 mg kg−1)
NTG (5 mg kg−1)
0
50
100
150
200
250 **P = 0.002
Saline
0
50
100
150
200
250
0
50
100
150
200
250
Time (min)
BL1
BL230
60
120
180
0
2
4
6
Orofacial pain score
Time (min)
Orofacial pain score
BL1
BL2
30
60
120
180
0
2
4
6
NTG (5 mg kg−1)
AAV5/2.hGFAP.eGFP (control)
AAV5/2.hGFAP.ATP1A2 (rescue)
**
0
50
100
150
200
250
AAV5/2.hGFAP.eGFP (control)
or
AAV5/2.hGFAP.AT P1A2 (rescue)
FHM2 KI mice
21 d.p.i.
Wild-type or FHM2 KI mice
NTG (NO donor) or saline (i.p.)
Bilateral in cingulate cortex
Saline
NTG (10 mg kg−1)
Saline
NTG (5 mg kg−1)
Saline
FHM2 KI mice
Mechanical stimulation
AAV.eGFP
AAV.ATP1A
2
NTG (5 mg kg−1)
0.1-g filament
NTG (5 mg kg−1)
0.1-g filament
Mechanical stimulation
H
G
Time (min)
Orofacial pain score
BL1
BL2
30
60
120
180
0
2
4
6
0.1-g filament
Fig. 5. Local rescue of astrocyte dysfunction in the Cg reduces facial pain in FHM2
mice. (A) Schematic illustration of the experimental design. (B) Time course showing
the orofacial pain score in WT mice injected with either saline (gray) or NTG (black).
NTG (10 mg kg−1) elicits hypersensitivity to touch at 30, 60, and 120 min following in-
jection. (C) Same as (B) but for FHM2 mice injected with saline (gray) or NTG (red). NTG
(10 mg kg−1) evokes higher orofacial pain scores at 30, 60, and 120 min following injec-
tion. (D) A lower dose of NTG (5 mg kg−1) does not elicit a higher sensitivity in WT mice.
(E) In FHM2 mice, NTG (5 mg kg−1) evokes a higher orofacial pain score at 30 min after
injection. (F) Schematic illustration of the experimental design. (G) Three-dimensional
reconstruction of a cleared brain imaged with light-sheet microscopy showing the
volume and site of the injection in Cg after ex vivo fixation. Scale bar, 300 m. The bottom
left image shows a single coronal section, and dots indicate the center of the injection
volume of the viruses from six brains. (H) Time course showing the orofacial pain score
in FHM2 mice injected with the control virus (red) or the rescue virus (blue). NTG
(5 mg kg−1) only evoked a higher orofacial pain score at 30 min after injection in
contro l FHM2 mice and not in rescued FHM2. Data are means ± SEM. Two-way ANOVA
with Bonferroni post hoc test and two-tailed unpaired t test were used.
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DISCUSSION
Astrocytic dysfunction enhances NMDA-mediated dendritic
excitability in Cg of FHM2 mice
In this study, we first confirmed that in the adult Cg of FHM2 mice,
similar to the developing somatosensory cortex (3), 2 NKA dys-
function impairs both K+ and synaptically released glutamate up-
take by astrocytes. Western blot analysis performed in FHM2 mice
also revealed a significant reduction in the expression levels of 2
NKA and the GluT GLAST, which we previously reported to be
implicated in astrocyte-mediated glutamate uptake in this cortical
region (17). The reduced GLAST expression in FHM2 Cg could
suggest potential physical coupling between 2 NKA and GLAST in
Cg astrocytes, as was previously shown to occur between 2 NKA
and GLT-1in perisynaptic astrocytic processes of the somatosensory
cortex (9). As a consequence of this tight coupling, a reduced density
of GLT-1 transporters in perisynaptic astrocytic processes could be
previously detected with electron microscopy in the developing barrel
cortex of FHM2 mice (3). A local reduction of GLT-1 density spe-
cifically around synapses may remain undetected in Western blots
and be consistent with the unaltered expression of GLT-1in FHM2
Cg (28). With regard to the reported impairment of K+ clearance in
the Cg of FHM2 mice, this is likely to be directly ascribed to the re-
duced 2 NKA expression and function, which plays a major role in
K+ clearance following trains of high-frequency stimulation (29).
The slowdown in glutamate clearance by astrocytes prolonged
the presence of elevated glutamate levels in the extracellular space.
Increase in glutamate spillover enhances cortical excitatory neuro-
transmission, particularly NMDA receptor–mediated transmission
(17). Consistently, we found a facilitation of NMDA spike genera-
tion in tuft dendrites of L5 pyramidal cells both in FHM2 mice and
upon subsaturating concentrations of the GluT blocker dl-TBOA
(that reduces glutamate clearance to an extent similar to that pro-
duced by the FHM2 mutation) in WT mice. The probability of AP
firing following NMDA spikes was also increased in FHM2 mice. In
contrast, no changes were observed in spontaneous and evoked synaptic
transmission in FHM2 mice.
To establish a causal relationship between the astrocyte malfunc-
tion and the observed neuronal modifications in FHM2, we com-
pensated for the astrocyte dysfunction by expressing the WT form
of the Atp1a2 gene in the Cg of FHM2 mice. This intervention re-
versed the defective glutamate and K+ clearance by astrocytes.
Compensating the astrocyte dysfunction reduced the facilitation
of dendritic NMDA spike generation, thereby lowering the output
firing induced by these spikes. These findings demonstrate that NMDA
spike generation on tuft dendrites of L5 pyramidal cells and the sub-
sequent neuronal output are directly and dynamically affected by
astrocytic dysfunction.
NMDA spikes are believed to be the dominant mechanism by which
distal synaptic inputs lead to firing of pyramidal neurons in the cortex
(30). Since dendritic spikes increase the computational properties of
individual neurons (19,30), the facilitation of NMDA spike genera-
tion and the resulting firing of L5 pyramidal cells (the main output
cells of the Cg) could greatly influence network activity in down-
stream cortical and subcortical areas involved in pain processing.
The altered cellular functions in the Cg of FHM2 mice lead
to hypersensitivity to a migraine-relevant pain trigger
In migraineurs, NTG administration induces a delayed migraine-
like headache with associated features such as premonitory symptoms
and allodynia (24). NTG-induced hyperalgesia in animals provides
a behavioral model of the NTG-induced allodynia observed in
migraineurs during the attack (25). Using this model, we found that
relatively low doses of NTG trigger hypersensitivity to facial mechanical
stimulation in FHM2 mice, while they are ineffective in WT mice. A
similar finding was previously reported for another genetic mouse
model of migraine (26). Local rescue of the astrocytic defect by
expressing the WT Atp1a2 gene in the Cg of FHM2 mice strongly
reduced their increased nociceptive response upon NTG treatment.
This finding is consistent with the conclusion that the hypersensitivity
of FHM2 mice to a migraine-relevant trigger is largely due to altered
neural function in the Cg.
The Cg involvement in migraine pathophysiology
The anterior Cg (ACC) and midcingulate cortex (MCC) play a key
role in pain processing (23,31). The ACC is consistently activated
in humans and animal models upon nociceptive stimuli (32), and
neuronal plasticity in the ACC is correlated with the development
of chronic pain (21,23). The ACC is also among the regions that are
activated during spontaneous migraine attacks (33) and during the
premonitory phase of the delayed migraine-like headache induced
by NTG infusion in migraineurs (34). Moreover, functional imaging
studies show increased activation of both the ACC (12) and the MCC
(13,35) in response to noxious (including trigeminal) stimulation
in migraineurs during the interictal period. Migraineurs show in-
creased activation of the MCC that correlates with increased pain
rating (sensitization) during repeated trigeminal noxious stimula-
tion, while healthy controls show decreased MCC activation and
pain rating (habituation) (13). However, the underlying mechanism
of this hyperactivity and its potential involvement in cranial pain
induction was still elusive. We report an astrocyte-mediated facili-
tation of NMDA spike generation and a subsequent increase of L5
pyramidal cell firing in the Cg of FHM2 mice, which may be potentially
involved in pain generation and/or sensitization of the pain process-
ing system in familial migraine. The evidence that astrocyte dysfunc-
tion in the Cg can specifically increase facial tactile sensitivity to a
pain trigger supports the notion that the Cg is a critical hub in pain
processing and may additionally gate the activation of cranial pain
pathways. Whether the ACC, the MCC, or both mediate this remains
unclear. The ACC is broadly connected to the salience network and
projects to the periaqueductal gray, rostral ventromedial medulla,
and dorsal horn (5). An interesting pathway from the MCC to the
posterior insula (and involving descending serotoninergic facilitation
of nociception from the raphe magnus nucleus) has been shown to
be necessary and sufficient for the induction and maintenance of
pain sensitization and could also be involved in the observed behavior
(31). Therefore, heightened activity in the ACC and/or MCC could
have broad effects on migraine-relevant pain perception and descend-
ing modulatory circuits, which could lead to a loss of pain inhibition
and/or to pain facilitation. It would therefore be intriguing to test
how the rescue of the astrocytic mutation in the ACC or MCC affects
the activity of those downstream regions crucial for the development
or sensitization of cranial pain.
Our data support the idea that FHM2-associated astrocytic dys-
function in particular brain regions may engender different migraine-
relevant functional consequences. In the somatosensory cortex, it
lowers the CSD threshold with a potential impact on aura occurrence
and cranial nociceptor activation (3,10). CSD facilitation is a common
feature of all genetic models of migraine that have been investigated
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so far (26,36,37). In the hippocampus, the FHM2 mutation causes
abnormal region-dependent synaptic plasticity, which might underlie
some of the memory deficits observed in FHM2 patients (38). We
demonstrate instead that Cg astrocyte dysfunction in FHM2 mice
leads to hypersensitivity to a migraine-relevant trigger.
In conclusion, we provide evidence that astrocyte dysfunction in
the Cg, a nonsensory cortical area, is implicated in heightened
sensitivity to head pain triggers and may be involved in pain gener-
ation and/or sensitization of the pain processing system in familial
migraine. Understanding the cellular and molecular nature of
circuit-specific network dysfunctions associated with familial migraine
might be key to shed light on incompletely understood aspects of
migraine pathophysiology.
MATERIALS AND METHODS
Animals
Experiments were performed using adult (5 to 9 weeks old) hetero-
zygous KI mice harboring the W887R FHM2 mutation [Atp1a2+/R887
mice; (10)] and their WT littermates (background C57BL/6J; male
and female in equal or near-equal number). Adult mice were group-
housed up to five in filter-top cages with a standard 12-hour lig ht/ 12-h our
dark cycle and food and water available ad libitum. Permission for
animal experiments was obtained from the Tierversuchskommission
of the canton of Zurich, Zurich, Switzerland. All animal experiments
complied with the relevant ethical regulations.
Chemicals and drugs
Reagents for artificial cerebrospinal fluid (ACSF) and internal solution s,
biocytin, 6-nitro-7-sulfamoylbenzo[f]quinoxaline-2,3-dione
(NBQX), and picrotoxin were obtained from Sigma-Aldrich. 6-cyano-
7-nitroquinoxaline-2,3-dione (CNQX), AP-V, dl-TBOA, and d-serine
were obtained from Tocris. NBQX, CNQX, and dl-TBOA were dis-
solved in dimethyl sulfoxide. Tetrodotoxin (TTX) was obtained from
Abcam. NTG was obtained from Sigma-Aldrich. Picrotoxin was disso lved
in ethanol (EtOH). AP-V, d-serine, and TTX were dissolved in ddH2O.
Acute brain slice preparation
Mice were briefly anesthetized with isoflurane and decapitated. The
brain was quickly removed and transferred to an ice-cold solution
containing 65 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 25 mM
NaHCO3, 7 mM MgCl2, 0.5 mM CaCl2, 25 mM glucose, and 105 mM
sucrose saturated with 95% O2 and 5% CO2; coronal slices (350 m thick)
containing the ACC were cut from the tissue block with a vibratome
(HM 650, Microm). Slices were then transferred to a recovery solu-
tion containing 130 mM K-gluconate, 15 mM KCl, 0.2 mM EGTA,
20 mM Hepes, and 25 mM glucose for 2min before being kept in
oxygenated ACSF (315 mosm) saturated with 95% O2 and 5% CO2
and containing 125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 25 mM
NaHCO3, 1 mM MgCl2, 2 mM CaCl2, and 25 mM glucose at 34°C
for 25min and then at room temperature until use. Slices used for
two-photon glutamate imaging and astrocytic patch-clamp recordings
were loaded with SR-101 (1M) for 15min at 34°C before being
kept in ACSF at room temperature.
Electrophysiological recordings
Whole-cell recordings from astrocytes
Recordings were performed as previously described in (17). Individual
slices were transferred to a recording chamber perfused with oxygenated
ACSF, at a flow rate of 1 to 2 ml/min at 32° to 34°C. Whole-cell re-
cordings were taken from L1 astrocytes in the Cg. Unless otherwise
stated, cell bodies of astrocytes were visualized using an astrocyte-
specific dye (SR-101; see above) that was excited with wLS broad-
band light-emitting diode illumination (460 nm), and images were
acquired with Retiga R1 camera using Ocular software (QImaging,
Germany) with a 40× water immersion objective. In addition, astro-
cytes were recognized by their hyperpolarized resting membrane
potential, their linear current-voltage relationship, their inability to
generate APs, and their low input resistance. Recordings were taken
with borosilicate glass pipettes (4 to 8megohm) containing the fol-
lowing internal solution: 115 mM K-gluconate, 6 mM KCl, 5 mM
glucose, 7.8 mM Na-phosphocreatine, 4 mM Mg-ATP (adenosine
triphosphate), 0.4 mM Na-GTP (guanosine triphosphate) [pH 7.25
with KOH; osmolarity, 295 mosm (readjusted with sucrose when
necessary)]. Recordings were performed using MultiClamp 700B
amplifier, and data were acquired with a Digidata 1550A 16-bit
board (all from Molecular Devices). For the recording of STCs, the
extracellular solution contained antagonists of NMDA receptors
(AP-V; 50 M), AMPA receptors (CNQX or NBQX; 10 M), and
-aminobutyric acid type A (GABAA) receptors (picrotoxin; 100 M).
Astrocytes were held at −80 mV, and STCs were evoked by single-
pulse stimulation or by trains of 10 and 11 pulses at high frequen-
cies (50and then 100 Hz) every 20 s. The K+ inward current was
characterized as the slowly decaying inward current elicited in as-
trocytes upon high-frequency synaptic stimulations. Every protocol
was repeated at least five times and then averaged and analyzed.
Currents were evoked by focal electrical stimulation (bipolar, 100 s,
8.5 V) through a theta glass pipette placed in L1, in the proximity of the
recorded astrocyte. Access resistance was monitored (<16megohm),
and recordings with an access resistance changing more than 30%
between the beginning and the end of the recording were discarded.
Resting membrane potential and input resistance were monitored
for analysis of electrophysiological properties of astrocytes. The de-
cay kinetics of the last pulse of the trains were analyzed by subtract-
ing the current elicited by 10 pulses from that elicited by the 11th
pulse.
Whole-cell recordings from L5 pyramidal neurons
Somas were patched with borosilicate glass pipettes (2.2 to 4megohm).
Cells were clamped at −70 mV, and focal synaptic stimulation was
performed through a theta patch pipette located close to the selected
apical tuft dendritic segments in L1. NMDA spikes were evoked by
applying paired-pulse stimulations of 50Hz of increasing stimula-
tion intensities (from 1 to 2.5 mA or 5.5 mA). Recordings with an
access resistance >15 megohm were discarded. The following internal
solution was used: 130 mM K-gluconate, 5 mM KCl, 10 mM Hepes,
10 mM phosphocreatine, 4 mM Mg-ATP, 0.3 mM GTP, and biocytin
(1.5 mg/ml) (pH 7.3 with KOH; osmolarity, 294 mosm). To visualize the
dendrites, patch pipettes also contained Alexa Fluor 488. Recordings
were performed in the presence of GABAA receptor blocker (picro-
toxin; 100 M) and d-serine (10 M).
For mEPSCs, somata were targeted for recording with borosilicate
glass pipettes (2.2 to 4megohm) containing the following: 130 mM
gluconic acid, 130 mM CsOH, 5 mM CsCl, 10 mM Hepes, 1.1 mM EGTA,
10 mM Na-phosphocreatine, 4 mM Mg-ATP, and 0.3 mM Na-GTP.
The pH of the intracellular solution was adjusted to 7.3 with CsOH,
and biocytin (1.5mg ml−1) was added for the reconstruction of neurons.
Cells were held at −70 mV, and synaptic responses were recorded in
the presence of picrotoxin (100 M) and TTX (1 mM). Recordings
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with an unstable baseline or a holding current less than −400 pA
were rejected. Currents were filtered off-line using a Butterworth
low-pass filter (2 kHz) and analyzed in 1-min epochs using the Mini
Analysis Program 6.0.7 (Synaptosoft Inc., USA). Recordings with
leak current increasing more than 100 pA and access resistance
changing more than 30% between the beginning and the end of the
recording were discarded. At least 100 events were analyzed for every
condition. Events were identified as mEPSCs by setting the event
detection threshold at least twice the baseline noise level and by
checking that events had (i) rise times faster than the decay time, (ii) rise
times greater than 0.5 ms, and (iii) decay times greater than 1.5 ms.
Events not fitting the above parameters were rejected. Event amplitudes,
interevent intervals, and rise and decay times were first averaged
within each experiment and regrouped by condition. The resulting
means were averaged between experiments. Single-cell properties
(access resistance, membrane capacitance, etc.) were analyzed with
Clampfit 10.5 (Axon Instruments, Union City, CA). Graphs were
made using GraphPad Prism and Illustrator 15.1.0 (Adobe). For
evoked AMPA-mediated excitatory postsynaptic currents (eEPSCs),
cells were held at −70 mV and focal synaptic stimulation was
performed through a theta patch pipette located close to the tuft
dendrites in L1. AMPA eEPSCs were evoked by applying either a
single stimulation or paired-pulse stimulations at 20Hz. Recordings
were acquired in the presence of picrotoxin (100 M) and AP-V (50 M)
in ACSF.
Biocytin labeling
For some experiments, internal solutions used for recording of py-
ramidal cells contained biocytin (1.5 mg/ml) that diffused in the
cells for at least 10 min, as previously described in (17). Briefly, slices
(350 m) containing the recorded cell were then fixed in 4% para-
formaldehyde (PFA) at +4°C overnight. The following day, slices were
washed in phosphate-buffered saline (PBS) and 274 mM NaCl be-
fore being transferred to a blocking solution containing 10% nor-
mal goat serum in 0.3% Triton-PBS and 274 mM NaCl for 1 hour.
Afterward, slices were put in a blocking solution containing Alexa
Fluor 647–conjugated streptavidin (1:700; Jackson ImmunoResearch
Europe Ltd.; code: 016-600-084) for 2 hours. Slices were then
washed in PBS and 274 mM NaCl before being mounted on Superfrost
Plus slides. Images were acquired on a Zeiss LSM 710 Pascal confocal
microscope using a 0.9 numerical aperture ×10 Plan-Apochromat
objective (for L5 pyramidal cells) and the ZEN 2012 software (Carl Zei ss).
Whenever applicable, contrast and illumination were adjusted in
ImageJ. Presented images are Z projections.
AAV5/2 generation and injections in vivo
The mAtp1a2 gene (WT form of 2 NKA for compensation exper-
iments) and the EGFP gene (for control experiments) were cloned
into plasmid backbones containing a shorted glial fibrillary acidic
protein (HgfaABC1D) promoter as in (22). These plasmids were
packaged into AAV serotype 5 [AAV-5/2-hGFAP-mATP1A2-bGH p(A)
and AAV5/2- hGFAP-hHBb1/E-EGFP-bGHp(A)] by the Viral Vector
Facility of the University of Zurich. All electronic information con-
cerning the plasmids and viruses is available online in the viral vector
repository (https://vvf.ethz.ch). FHM2 KI mice were used in all
experiments in accordance with institutional guidelines.
All surgical procedures were conducted under general anesthesia
using continuous isoflurane inhalation (induction at 5% and main-
tenance at 1 to 3%). Following induction of anesthesia, the mice
were placed into a motorized stereotaxic frame (David Kopf Instru-
ments and NeuroStar) using adjustable ear bars and an anesthesia
mask with an incisor bar. The animal was maintained at a physio-
logical temperature using a heating mat placed between the animal
and the frame. Vitamin C ointment was applied to the eyes to prevent
corneal drying during the operation. Mice were subcutaneously
administered buprenorphine (0.2 mg/kg) before surgery. The fur
between the ears was shaved, scrubbed with EtOH and betadine,
and dried, and a longitudinal incision of approximately 3 to 5mm
was made in the skin above the skull. For unilateral injections, a single
hole was drilled through the skull directly above the Cg on one
hemisphere (stereotaxic coordinates with respect to bregma: 0.75mm
anterior, 0.3mm lateral, and 1.25mm ventral). For control experiments,
0.5 l of the control virus AAV5/2- hGFAP-hHBb1/E-EGFP-bGHp(A)
was injected through a glass pipette. Since AAV vectors size is limited
to ~4.7 kb, it was not possible to include a reporter gene in the rescue
virus AAV-5/2-hGFAP-mATP1A2-bGHp(A). Therefore, to visualize
the injection site, we injected a mixture of 0.5 l of the two viruses
with a ratio of 2:1 of the rescue virus to the control virus containing
eGFP. Glass pipettes were left in place for at least 10min follow-
ing infusion of the virus. Surgical wounds were closed with single
5-0 nylon sutures. Following surgery, animals were closely monitored.
Mice were euthanized 15 to 25 days after surgery for electrophysiology
experiments or were used for behavioral experiments. When assess-
ing behavior, AAVs were injected bilaterally. The experimenter was
blinded to which AAV was injected.
Two-photon glutamate imaging
Mice aged between 4 and 6 weeks were injected with 0.3 to 0.5 l of
AAV2/1.GFAP.iGluSnFr.WPRE.SV40 (Penn Vector; provided by
L. Looger, Janelia Farm) unilaterally into the Cg through a glass
pipette as described above. Fifteen to 21 days following the virus
injections, coronal brain slices (350 m) containing the Cg were
obtained as previously described (21). Imaging was performed as
described in (17). Briefly, a galvanometer-based two-photon laser
scanning system was used to image extracellular glutamate (16×
objective; zoom, 6; excitation wavelength, 900 nm; 64pixels by 64 pixels
per image; acquisition rate, 19.2 Hz). Astrocytes were visualized using
SR-101. Synaptic glutamate release was elicited by trains of 10 pulses
at high frequency (50 or 100 Hz) every 20 s delivered via a theta
glass pipette (bipolar, 100 s; stimulation intensity, 3 to 5 V) placed
in the inner L1. To visualize the theta glass pipette, it was filled with
ACSF containing 1 M SR-101. Note that SR-101 staining does not
affect the kinetics of iGluSnFr transients nor of STCs as shown in
(17). Moreover, when we doubled the acquisition rate to 38.4 Hz,
the iGluSnFr decay kinetics remained unaltered, indicating that the
image acquisition rate used was sufficient to detect the monitored
changes. All solutions contained 10M NBQX or CNQX, 50M
AP-V, and 100M picrotoxin, and temperature was kept between
32° and 34°C during imaging. Ten consecutive sweeps were acquired
and subsequently analyzed using ImageJ. Fluorescence emission
was collected from an ROI (diameter, 34 m) 10 to 40 m away
from the stimulation pipette. The average background value was
derived from a region within the field of view that was free
of clearly visible iGluSnFr (typically in the contralateral hemi-
sphere) and subtracted from the fluorescence intensity of the ROIs
for each frame. Traces were then averaged, and decay tau was cal-
culated by fitting a single exponential function using Igor Pro
(WaveMetrics).
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Behavioral analysis
Orofacial von Frey
Behavioral testing was performed in a dimly lit and quiet room by
the same female experimenter. Throughout the week before the first
behavioral testing, mice were habituated to handling by the experi-
menter. Each mouse was placed in Plexiglas cages of about 20cm by
20cm sitting on a metal grid ground floor. Mice were allowed to
accommodate to the cage for 1 hour before the testing. In all experime nts,
0.025- and 0.01-g von Frey filaments were used. The first tests were
performed using the lower-force filament. Filaments were applied
to the orofacial area close to the whisker pad or close to a 90° angle
until bent, in three series of four pokes, in the middle or on either
left or right side of the snout. After a total of 12 pokes with the
0.025-g filament, the stimulation with the 0.01-g filament was
applied in the same way. The responses recorded were as follows:
unilateral or bilateral forepaw swipes across the face (1 point each),
continuous forepaw swipes (three or more: 1.5 points), aggression/
biting of the probe following stimulus (0.25 points), and clear with-
drawal of the head from the stimulus (0.25 points) as described
in (27). Before the injection of NTG (5 or 10 mg/kg) or 0.9% saline
(intraperitoneally), baseline 1 was recorded several hours or a day
before the experiment and baseline 2 was immediately before intra-
peritoneal injections. Testing was carried out at 30, 60, 120, and
180min after injection. The experimenter was blinded to the treat-
ment in all behavioral experiments.
Locomotor activity
NTG (5 mg/kg, i.p.) or saline was administered 30min before testing.
Locomotor activity was measured in an open-field arena (radius,
10 cm) equipped with four pairs of light beams and photosensors
and analyzed for the time interval between 10 and 60min after NTG
or saline administration.
Brain clearing and light-sheet microscopy
After fixation and dehydration
Mice were anesthetized with isoflurane and decapitated. Entire brains
were extracted, briefly washed in PBS to remove excessive blood, and
postfixed in 4% PFA for 2 days at 4°C. After fixation, brains were washed
two times in PBS and subsequently dehydrated in increasing alcohol
concentrations [30, 50, 70, 80, 90, 96, and 100% EtOH (each adjusted
to pH 9.5)] for a day each (39). Tissue shrinkage of up to 50% was
observed during the dehydration process. After dehydration, brains were
transferred into a clearing solution of benzyl alcohol and benzyl benzoate
(1:2) solution (BABB) in separate glass vials on a gentle shaking or rotat-
ing cycle under a chemical hood for a minimum of 1 day until transparen t.
Imaging
Cleared brains were transferred to an immersion cuvette containing
BABB and placed in the imaging reservoir of the microscope. Images
were acquired using the mesoscale single-plane illumination micro-
scope mesoSPIM system (www.mesospim.org) with an Olympus MVX10
macroscope in the detection path and an MVPLAPO 1× objective
(40). The fluorescent signal in the sample was recorded at ×1.6 magnifica-
tion by moving the sample through the light sheet in 4-m steps. The
sample was illuminated from one side using Toptica multi-laser engine
laser at excitation wavelengths of 488 and 640nm. Postprocessing of
images was carried out using ImageJ and Imaris software (Bitplane).
Western blots
Two weeks following stereotaxic viral injections in the Cg, tissue
of the Cg was rapidly dissected and immediately frozen on dry ice
and stored at −80°C until used. On the day of the experiment, the
tissue (10 to 20 mg) was thawed and homogenized by sonication in
20 volumes of 20 mM tris (pH 7.4) containing the protease inhibitor
cocktail cOmplete Mini (Roche Diagnostics). The homogenate was
centrifuged for 10min at 1000g, and the supernatant was recovered.
After total protein quantification using the Bradford protein assay
(Bio-Rad), the samples were incubated with Laemmli sample buffer
(Bio-Rad) for 30min at 37°C. Aliquots containing 10g (GLAST)
or 15 g of protein (ATPase 2 and GLT-1) were subjected to SDS–
polyacrylamide gel electrophoresis using 10% (GLAST and GLT-1)
or 7.5% (ATPase 2) mini gels (Mini-PROTEAN 3, Bio-Rad). Pro-
teins were transferred onto nitrocellulose membranes in a Trans-
Blot semi-dry transfer cell (Bio-Rad) at 15 V for 60min using 39 mM
glycine, 48 mM tris, 1.3 mM SDS, and 20% methanol as transfer
buffer. After blotting, the transferred proteins were stained with
REVERT Total Protein Stain (LI-COR) and immediately imaged
using the Odyssey CLx imager (LI-COR). For immunodetection,
the blots were blocked for 1 hour in PBS containing 5% nonfat dry
milk at room temperature, followed by incubation at 4°C overnight
with anti–2 NKA antibodies (1:1000; rabbit polyclonal, Sigma-
Aldrich, catalog no. 07-674), anti–GLT-1 antibodies (1:1000; rabbit
polyclonal, knockout verified, Synaptic Systems, catalog no.
250203), or anti-GLAST antibodies (1:10,000; rabbit polyclonal,
knockout verified, Synaptic Systems, catalog no. 250113), diluted in
PBST [PBS (pH 7.4) and 0.05% Tween 20] containing 5% nonfat
dry milk. The blots were then washed five times for 5 min with Tris
Buffered Saline + Tween 20 (TBST) (10 mM Tris, pH 7.4, 150 mM
NaCl, 0.05% Tween 20) and incubated with secondary antibodies
(1:20,000; goat anti-rabbit Alexa Fluor Plus 800) for 1 hour at room
temperature. Following extensive washing (see above), immuno-
reactivity was detected using the Odyssey CLx imager (LI-COR).
Immunoreactivity was quantified with the Image Studio software
(LI-COR) and normalized to total protein in the corresponding
lanes. Each antibody was tested for its linear detection range using
different protein and antibody concentrations.
Immunohistochemistry and image analysis
Three weeks after stereotaxic viral injections in the Cg, FHM2 KI mice
were anaesthetized with pentobarbital (160mg kg−1, i.p.) before
transcardiac perfusion with 20ml of phosphate buffer followed by
100ml of 4% ice-cold PFA [in 0.1M sodium phosphate buffer (pH 7.4)].
Brain tissue was postfixed for 4 hours with 4% PFA on ice and cryo-
protected in 30% sucrose solution (in 0.1M sodium phosphate buffer)
overnight at 4°C. Brains were embedded in NEG50 frozen section
medium (Richard-Allan Scientific) and cut into 40-m free-floating
sections (Hyrax KS 34 microtome, Carl Zeiss). Brain sections were
left in antifreezing solution at −20°C until use. Following incubation
in blocking solution (PBS, 0.3% Triton X-100, and 10% normal
donkey serum) for 1 hour, brain sections were incubated at 4°C
overnight in a primary antibody solution (PBS, 0.3% Triton X-100,
and 10% normal donkey serum) containing combinations of the
following antibodies: chicken anti-GFP (1:1000; LifeTech,
AB_2534023), guinea pig anti-NeuN (1:1000; Synaptic Systems,
AB_2619988), and rabbit anti-S100B (1:700; Abcam, AB_52642).
Three washing steps of 10min each in PBS were performed before
incubating brain sections with secondary antibodies (1:500): Cyanine
Cy3 donkey anti-rabbit (Jackson ImmunoResearch, AB_2307443),
Alexa Fluor 647 donkey anti-guinea pig (Jackson ImmunoResearch,
AB_2340477), and Alexa Fluor 488 donkey anti-chicken (Jackson
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SCIENCE ADVANCES | RESEARCH ARTICLE
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ImmunoResearch, AB_2340376) for 90min at room temperature in
PBS supplemented with 0.3% Triton X-100. Images were taken with an
LSM 800 with Airyscan confocal microscopes (Carl Zeiss) controlled
with ZEN 2.3 (blue edition) software and using a Plan-Apochromat
×40/1.4 Oil DIC M27 oil-immersion objective. Z stack images of
10 optical sections and 1.5-m step size were used for the analysis of
fluorescence colocalization and to create maximum intensity pro-
jections images. Images were processed using ImageJ software.
Quantification and statistical analysis
Data are displayed as means±SEM in all experiments except for the
Western blot data that are displayed as means±SD. Statistical details
can be found in Results, figures, and figure legends. Statistical com-
parisons were made with two-tailed paired or unpaired t tests, one-
way analysis of variance (ANOVA) with Bonferroni post hoc test,
one-way ANOVA with Tukey post hoc test, or two-way repeated-
measures ANOVA test. All graphs and statistical tests were performed
using GraphPad Prism, and figures were prepared using Adobe
Illustrator CS5. P values less than 0.05 were considered statistically
significant.
SUPPLEMENTARY MATERIALS
Supplementary material for this article is available at http://advances.sciencemag.org/cgi/
content/full/6/23/eaaz1584/DC1
View/request a protocol for this paper from Bio-protocol.
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Acknowledgments: We thank G. Casari for generating and providing the FHM2 KI mouse
model; J. C. Paterna and M. Rauch from the local Viral Vector Facility for help with the
production of the viruses; H. Johannssen for assembling the two-photon microscope and
helping with two-photon imaging; E. Platonova and the Center of Microscopy and Image
Analysis at UZH for assistance with light-sheet microscopy; T. Grampp for help with Western
blots; G. Albisetti and R. Ganley for advice on immunohistochemistry and proofreading the
manuscript; S. Gudmundsdottir for some electrophysiological experiments; W. B. Gan and
J. Cichon for providing the GFAP-iGluSnFr AAV viruses and training with cortical glutamate
imaging; and members of H.U.Z. group, F. Brandalise, R. Min, P. Bezzi, and B. Weber for
discussion. Funding: This work was supported by the Swiss National Science Foundation
grant PP00P3_176838, Novartis Foundation for medical-biological Research grant no. 17C157,
and the Hartmann-Müller Stiftung no. 2253 to M.S. and a Telethon grant GGP14234 and PRIN
2017ANP5L8 to D.P. Author contributions: J.R. and M.S. conceived the project, designed
the study, wrote the manuscript, performed two-photon imaging experiments, and analyzed
the data. J.R. performed patch-clamp experiments, behavioral experiments, light-sheet
imaging, and immunohistochemistry experiments; contributed to virus production; and
analyzed the data. D.B. performed and analyzed Western blot experiments. H.U.Z. contributed
to reagents and equipment and discussed the interpretation of the data. D.P. provided the
FHM2 KI mouse model, discussed the interpretation of the data, and contributed to the study
design. All authors edited and approved the final manuscript. M.S. supervised the whole
project. Competing interests: The authors declare that they have no competing interests.
Data and materials availability: All data needed to evaluate the conclusions in the paper are
present in the paper and/or the Supplementary Materials. Data, associated protocols, and
additional information regarding this paper are available to the reader upon request from the
authors and are located in the internal server of the Institute of Pharmacology and Toxicology
at the University of Zurich.
Submitted 16 August 2019
Accepted 25 March 2020
Published 5 June 2020
10.1126/sciadv.aaz1584
Citation: J. Romanos, D. Benke, D. Pietrobon, H. U. Zeilhofer, M. Santello, Astrocyte dysfunction
increases cortical dendritic excitability and promotes cranial pain in familial migraine. Sci. Adv.
6, eaaz1584 (2020).
on June 5, 2020http://advances.sciencemag.org/Downloaded from
familial migraine
Astrocyte dysfunction increases cortical dendritic excitability and promotes cranial pain in
Jennifer Romanos, Dietmar Benke, Daniela Pietrobon, Hanns Ulrich Zeilhofer and Mirko Santello
DOI: 10.1126/sciadv.aaz1584
(23), eaaz1584.6Sci Adv
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