www.sciencemag.org/content/344/6188/1173/suppl/DC1

Supplementary Materials for

Sleep promotes branch-specific formation of dendritic spines after learning

Guang Yang, Cora Sau Wan Lai, Joseph Cichon, Lei Ma, Wei Li, Wen-Biao Gan*

*Corresponding author. E-mail: gan@saturn.med.nyu.edu

Published 6 June 2014, Science 344, 1173 (2014) DOI: 10.1126/science.1249098

This PDF file includes

Materials and Methods Figs. S1 to S8 References

Supplementary Materials Materials and Methods Experimental animals. Mice expressing YFP (H-line) or Cre (FVB/N-Tg(Thy1-Cre)1Vln/J) in

Layer V pyramidal neurons were group-housed in NYU Skirball animal facilities. The stress

hormone corticosterone (2.5 mg/kg) or NMDA receptor antagonist MK801 (0.25 mg/kg body

weight) was injected into the peritoneum of mice upon completion of the motor training session.

All experiments were performed in accordance with institutional guidelines.

Rotarod and treadmill training. An EZRod system with a test chamber (44.5 cm × 14 cm × 51

cm dimensions) was used in this study. Animals were placed on the motorized rod (30 mm in

diameter) in the chamber. The rotation speed gradually increased from 0 to 100 r. p. m. over the

course of 3 min. The time latency and rotation speed were recorded when the animal was unable

to keep up with the increasing speed and fell. Rotarod training/testing was performed in one 30-

min or 60-min session (20–40 trials). Performance was measured as the average speed animals

achieved during the training session. A backward running paradigm was introduced to provide

mice with new motor learning experience. In this paradigm, animals were forced to run

backward on the accelerated rod (speed increased gradually from 0 to 50 r. p. m. over 3 min) for

20–40 trials.

A custom built free-floating treadmill (96 cm × 56 cm × 61 cm dimensions) was also

used for motor training in this study. This free-floating treadmill allows head-fixed mice to move

their forelimbs freely to perform motor running tasks (forward or backward). To minimize

motion artifact during imaging, the treadmill was constructed so that all the moving parts (motor,

belt, and drive shaft) would not be in contact with either the microscope stage or the supporting

air-table. Animals were positioned on a custom-made head holder device that would allow the

micro-metal bars to be mounted (37). During motor training, the treadmill motor was driven by a

DC power supply. At the onset of a trial, the motor was turned on and the belt speed gradually

increased from 0 cm/s to 8 cm/s within ~3 sec, and the speed of 8 cm/s was maintained for the

rest of the trial.

For spine imaging in mice subjected to treadmill running, the animals were returned to

the home cage at the end of imaging session, which lasted less than 20 minutes. For calcium

imaging, the animals were head-restrained on the treadmill for about 7 hours with food and water

available on the side of the head-fixed device. To assay gait-running patterns, mouse forelimbs

were coated into ink and animals ran on white construction paper. Footprints were analyzed

offline by average step distance between two ipsilateral forelimb footprints in structured gait

pattern.

Sleep deprivation procedure. Sleep deprivation was achieved through gentle handling over a

period of ~7 hours after the first imaging session and rotarod training. Specifically, mice were

gently touched with a cotton applicator for 1–2 seconds whenever they displayed signs of

drowsiness. On average, mice were touched ~23 times per hour during the period of sleep

deprivation. The animals were not accommodated to this gentle handling protocol.

Imaging dendritic spine plasticity in awake, head-restrained mice. Dendritic spine imaging

was carried out in awake, head restrained Thy1-YFP mice through a thinned-skull preparation

(37). 24 hours before imaging, surgery was performed to attach a head holder and to create a

thinned-skull cranial window. Specifically, mice were deeply anesthetized with an

intraperitoneal injection of ketamine (100 mg/kg) and xylazine (10 mg/kg). The mouse head was

shaved and the skull surface was exposed with a midline scalp incision. The periosteum tissue

over the skull surface was removed without damaging the temporal and occipital muscles. A

head holder composed of two parallel micro-metal bars was attached to the animal’s skull to help

restrain the animal’s head and reduce motion-induced artifact during imaging. A small skull

region (~0.2 mm in diameter) was located over the primary motor cortex based on stereotaxic

coordinates (41) (1.0 mm posterior from bregma and 1.5 mm lateral from the midline) and

marked with a pencil. A thin layer of cyanoacrylate-based glue was first applied to the top of

entire skull surface, and the head holder was then mounted on top of the skull with dental acrylic

cement such that the marked skull region was exposed between the two bars. Precaution was

taken not to cover the marked region with dental acrylic cement.

In some experiments, four electrodes were implanted to allow simultaneous imaging and

EEG/EMG recording in the same animal. Two electrodes were used for recording epidural EEG

and two for recording EMG. Each electrode was made by soldering one end of an epoxy coated

silver wire (0.005 inch in diameter) to a connector pin. One EEG electrode was placed over the

left frontal cortex (2 mm lateral to midline, 2 mm anterior to bregma) and another on the

cerebellum (at midline, 1 mm posterior of lambdoid suture). Before the electrode implantation, a

small area of skull (each ~0.2 mm in diameter) was thinned with a high-speed drill and carefully

removed with forceps. The electrodes were bent at 1 mm from the tip of the silver wire and

carefully inserted under the skull above the dural matter. The electrodes were fixed by

cyanoacrylate-based glue and further stabilized by dental cement. Two electrodes for EMG

recording were placed on the nuchal muscle.

After the dental cement was completely dry, the head holder was screwed to two metal

cubes that were attached to a solid metal base, and a cranial window was created over the

previously marked region. The procedures for preparing a thinned-skull cranial window for two-

photon imaging have been described in detail in previous publications (42). Briefly, a high-speed

drill was used to carefully reduce the skull thickness by approximately 50% under a dissecting

microscope. The skull was immersed in artificial cerebrospinal fluid during drilling. Skull

thinning was completed by carefully scraping the cranial surface with a microsurgical blade to

~20 µm in thickness. A high quality picture of the brain vasculature was taken with a CCD

camera attached to a stereo dissecting microscope. Completed cranial window was covered with

silicon elastomer and the animals were returned to their own cages to recover.

Before imaging, mice were given one day to recover from the surgery related anesthesia,

and habituated for a few times (10 min each) in the imaging apparatus to minimize potential

stress effects due to head restraining and awake imaging. To image dendritic spines in un-

anesthetized mice, the head holder was screwed to two metal cubes attached to a solid metal base.

The silicon elastomer covering the thinned skull window was removed and the skull was

immersed in artificial cerebrospinal fluid. The head-restrained animal was then placed on the

stage of a two-photon microscope. The area of interest was selected and marked on the CCD

vasculature map taken previously. The two-photon laser was tuned to the wavelength of ~920

nm and images were acquired using 1.1 NA 60X water-immersion objectives. A low

magnification stack (200 µm × 200 µm; 512 pixel × 512 pixel) of fluorescently labeled neuronal

processes was taken and used as a map for relocation of the same area at later time points, in

addition to the marked brain vasculature map. Two to three stacks of image planes (66.7 µm ×

66.7 µm; 512 pixel × 512 pixel) within a depth of 120 µm from the pial surface were collected at

each time point, yielding a full three-dimensional data set of dendrites in the area of interest. The

animal was head restrained during image acquisition which took ~15 min, and immediately

released to its original cage and stayed there until the next imaging sessions.

In both control and experimental groups, about 50% of the animals were imaged twice to

generate one time point data and the rest were imaged 3–4 times to generate two data points in

Fig. 1B. The data from animals imaged 2 times or 3–4 times were comparable and thus grouped

together in Fig. 1B.

Data analysis of spine structural plasticity. All data analysis of spine remodeling was

performed blind to treatment conditions. Data analysis was performed with NIH ImageJ software

as described previously (18). More than 150 spines were analyzed from each animal. The same

dendritic segments were identified from three-dimensional stacks taken from different time

points with high image quality (ratio of signal to background noise > 4:1). The number and

location of dendritic spines were identified in each view without prior knowledge of

experimental condition. Filopodia were identified as long thin structures (generally larger than

twice the average spine length, ratio of head diameter to neck diameter < 1.2:1 and ratio of

length to neck diameter > 3:1). The remaining dendritic protrusions were classified as spines. No

subtypes of spines were separated. Three-dimensional stacks were used to ensure that tissue

movements and rotation between imaging intervals did not influence spine identification. Spines

were considered the same between views if their positions remain the same distance from

relative adjacent landmarks. Spines were considered different if they were more than 0.7 µm

away from their expected positions based on the first view. The degree of spine formation or

elimination was calculated as the number of spines added or eliminated divided by the number of

pre-existing spines.

EEG/EMG recording and analysis. Between imaging sessions, EEG/EMG was recorded with

band pass setting of 0.1–100 Hz and digitized at 10 KHz. EEG/EMG data were visually scored

for the states of wake and sleep. Wake state was identified by lower amplitude and higher

frequency (> 10 Hz) of EEG activity, and medium to high muscle activity. REM sleep was

identified by lower amplitude and higher frequency (> 10 Hz) of EEG activity, and low muscle

activity. NREM sleep was identified by higher amplitude and lower frequency (< 10 Hz) of EEG

activity, and low muscle activity.

Imaging and analysis of calcium signals from layer V neurons in mice expressing GCaMP6.

Genetically-encoded calcium indicator GCaMP6 Slow (S) was used for calcium imaging of layer

V pyramidal neurons in the primary motor cortex (at bregma and 1.5 mm lateral to midline) (43).

Recombinant adeno-associated viruses AAV-CaMKII-Cre (serotype 9) and AAV-Flex-CAG-

GCaMP6S (serotype 2/1) were used to drive the expression of GCaMP6S in layer V neurons

(>2×1013 (GC/ml) titer; University of Pennsylvania Gene Therapy Program Vector Core). In

some experiments, Thy1-Cre transgenic mice (FVB/N-Tg(Thy1-Cre)1Vln/J; The Jackson

laboratories) were used to label layer V neurons in combination with Cre-dependent GCaMP6S

(AAV-Flex-CAG-GCaMP6S). 0.1–0.2 µl AAV-Cre viruses were diluted 10X in artificial

cerebrospinal fluid (ACSF) and then injected into layer V of the motor cortex. The dilution in

ACSF allowed better spread of viruses through layer V and sparse neuronal labeling. Sparse

expression of GCaMP6 ensures that the contribution of fluorescent signals from the neuropile to

the somata is negligible. Two to three weeks after virus injection, layer V neurons were imaged

with two-photon microscopy.

For imaging somata of layer V pyramidal cells, a circular craniotomy (1.0 mm diameter)

was made above the primary motor cortex following implantation of the head holder and

EEG/EMG electrodes (same as Thy1-YFP animals). The craniotomy was covered with a round

glass coverslip (custom to the size of bone removed) that was glued to the skull to reduce motion

of the exposed brain. 24 h after window implantation, mice were head restrained in the imaging

apparatus on top of a custom-built free-floating treadmill, and imaging was performed with a

two-photon laser tuned to 920 nm. The average laser power on the sample was ~50 mW. All

experiments were performed using a 25× objective immersed in an ACSF solution and with a

1.5–2X digital zoom. All images were acquired at frame rates of 2 Hz (2-μs pixel dwell time)

using FV10-ASW v.2.0 software.

In all experiments, calcium imaging was first performed for 5 min while the mice were

under the quiet awake state positioned on the treadmill. Following this period of quiet

wakefulness, mice were either subjected to forward running for 20 trials (~1 min per trial), with

short resting breaks (20–30 s) in-between running trials, or allowed to sleep (termed pre-run

sleep). In mice undergoing treadmill training, calcium imaging was performed after every 4th

trial, giving a total of 5-min calcium recordings during the 20-min training period. All mice were

allowed to sleep on the two-photon microscope stage, either before training or after the training

session. Head-restrained mice usually experienced NREM sleep after 0.5 h (left undisturbed) and

calcium signals were recorded for 1 min approximately every 20 min (total of 5 min of calcium

recordings for each NREM sleep period). In the first series of experiments, mice were imaged

during quiet wakefulness, pre-running NREM sleep and/or running, then post-running NREM

sleep (Fig. 4C-F). In other mice, calcium recording in post-running NREM sleep were followed

over an extended period of time (a total of 8 h) (Fig. 4G). In the experiments where retraining

was preformed (Fig. 4G), mice were wakened after the first 4 h of sleep (post-run sleep 1) and

retrained with either forward- (F-F) or backward-running task (F-B). Similar to the first training

period, retraining lasted 20 trials and 5 min of calcium recordings were acquired over the second

training block. Following retraining, mice were allowed to sleep (post-run sleep 2) and calcium

imaging was performed during NREM sleep for the next 4 h (a total of 5-min calcium recordings

during sleep 2). For simultaneous calcium imaging and EEG recordings, optical recordings were

only taken during periods of slow-wave sleep (multiple time points during NREM sleep, 5 min

total of recordings).

Changes of neuronal activity during running and sleep, as indicated by GCaMP6

fluorescence changes, were analyzed post hoc using ImageJ software (NIH). Imaging movement

artifact occurred predominantly in the plane of imaging at the onset of running (less than 3

seconds) and these time-lapse frames were discarded from quantification. The lateral movement

of the images was typically less than 1 µm. Infrequent vertical movements were minimized due

to flexible belt design, two micro-metal bars attached to the animal’s skull by dental acrylic, and

a custom-built body support to minimize spinal cord movements. The imaging stacks were

registered using NIH ImageJ plugin StackReg. The cells that could be identified in all imaged

sessions were included in the data set. The fluorescence time course of each cell was measured

with NIH ImageJ by averaging all pixels within the circular ROIs covering the somata. The

ΔF/F0 is calculated as (F-F0)/F0, where F0 is the baseline fluorescence signal averaged over a 2-s

period before the onset of the motor task. Fluorescence changes of individual neurons during

running or NREM sleep was quantified as averaged ΔF/F0 over 5-min period of running or

NREM sleep period, respectively, and normalized to the averaged ΔF/F0 under the quiet awake

state.

All the cells analyzed in Figure 4 were active cells under quiet awake, running and sleep

conditions. Active cells are defined as those with changes of somatic fluorescence (∆F/F0) > 20%

during the 5-min imaging session under each condition. This threshold (20%) is more than 3

times the standard deviation of baseline fluorescence noise (~16.7%). Task specific neurons

were defined as cells that showed a large increase (>50%) of calcium level in somata during

running as compared to the quiet awake state (ΔF running /ΔF quiet > 1.5).

Statistics. All imaging data were presented as mean ± SEM. Tests for differences between

groups were performed using non-parametric tests and one-way ANOVA. Significant levels

were set at P ≤ 0.05. All statistical analyses were performed using the GraphPad Prism 6.

Supplementary Figures

Fig. S1. Motor training induces dendritic spine formation on a subset of dendritic branches.

(A-B) Relative (A) and cumulative (B) distribution of spine formation rates among all dendritic

branches (including sibling and non-sibling branches). 24 h after training, the degree of new

spine formation varies significantly among dendritic branches. About 30% of dendritic branches

in trained mice exhibit a significantly higher rate of spine formation than the branches in the non-

trained control mice. (C-D) There is no significant difference in spine elimination on dendritic

branches between trained and non-trained animals over 24 hours.

Fig. S2. Sibling branch analysis. (A) The sibling branches were defined as two apical tuft

branches that share the same parent branch. For example, branches 1 & 2 are a pair of sibling

branches, branches 3 & 4 are a different pair of sibling branches. (B) Analysis for each sibling

branch started from the branching point of the parent branch and extended towards the distal end

of the branch. A total of 209 sibling branch pairs (418 dendritic branches) were analyzed in this

study. In 52 out of 418 branches (~12%), the entire tuft branches were analyzed (e.g. branch 1).

In 84 out of 418 branches (~20%), the analysis stopped at the next branching point (e.g. branch

4). For the rest of 282 branches, the analysis stopped before it reached to the branch tip or the

next branching point due to the fact that dendrites were out of the imaging field (e.g. branches 2

& 3). The average length of dendritic branches in the study is 62.7 ± 1.3 µm (mean ± SEM). The

average length of high formation branches (HFBs) analyzed in this study is 63.7 ± 1.9 µm, and

the average length of low formation branches (LFBs) is 61.7 ± 1.8 µm.

Fig. S3. ELISA quantification of plasma corticosterone under various conditions. Plasma

corticosterone (Cort) was measured with a commercially available ELISA kit as described

previously (36). Trunk blood was collected from separate cohorts of mice that were (1) trained

with sleep (F/S); (2) trained and then deprived of sleep for 7 hours (F/SD); (3) trained and

restrained for 20 min; (4) trained and then injected with corticosterone at the dose of 2.5 mg/kg.

Blood samples in groups 3 and 4 were collected immediately after 20-min restraint and ~20 min

after corticosterone injection, respectively. Sleep deprivation via gentle handling caused ~150%

increase of plasma corticosterone in sleep-deprived mice (F/SD) than in non-deprived mice (F/S).

The stress caused by physical restraint (20-min restraint), as measured by the level of

corticosterone, is much higher than that induced by gentle handling. 20 minutes after i.p.

injection of corticosterone (2.5 mg/kg), the level of plasma corticosterone is the highest among

all groups. Corticosterone at this dose was chosen to control for the potential effect of the

increased level of corticosterone on branch-specific spine formation over 8 hours.

Fig. S4. The reduction in spine formation after 7-h SD could not be rescued by subsequent

sleep. 24 hours after learning, the rate of spine formation on either HFBs or LFBs was lower in

SD mice as compared to non-SD mice. **P < 0.01, non-parametric test. The survival of new

spines formed on LFBs during 0-8 h period was similar over the next 16 hours in mice with or

without previous sleep deprivation. Notably, new spine formation on LFBs is significantly

higher from 8 to 24 hours in mice without being sleep-deprived previously than in mice sleep-

deprived previously for 7 hours. Whether or not sleep rebound from 8 to 24 hours is involved in

this phenomenon remains to be investigated.

Fig. S5. New persistent spines and performance 24 hours after motor training. (A) The

scatter plot of the total persistent new spines (summation of new spines formed on HFBs and

LFBs over 8 hours and persisted at 24 h) and the performance at 24 h, which was measured as

the average maximum speed that mice achieved for each trial before falling off the rod. (B) The

scatter plot of persistent new spines formed on HFBs and the performance. (C) Plot of persistent

new spines formed on LFBs and the performance 24 hours after training. Each circle represents

an individual animal.

Fig. S6. Treadmill motor training induces branch-specific spine formation as rotarod

motor training. (A) Treadmill motor training/testing was performed in one 40-trial session (~40

min), and mice were given 5-min break in the middle of each session. (B) Treadmill performance

was measured as the average step distance animals achieved during each training session. Both

forward (8 mice) and backward (6 mice) running paradigms were introduced to provide mice

with different motor learning experiences. Similar to rotarod motor skill learning, mice showed

intra- and inter-session improvement in performance for both tasks. Data are presented as mean

± SEM. (C) Similar to rotarod running, forward or backward treadmill running task also induced

branch-specific spine formation over 8 hours. A total of 12 sibling branch pairs (826 spines)

from 4 mice were analyzed for forward running task. For backward running, 11 sibling branch

pairs (714 spines) from 4 mice were quantified. ****P < 0.0001, non-parametric test. (D) There

is no significant difference in spine elimination between sibling branches in both forward (P >

0.74) and backward running task (P > 0.38, non-parametric test).

Fig. S7. Cells active during running are specifically activated during subsequent sleep. To

rule out the possibility that certain cells are active throughout sleep and running epochs, the same

neurons were imaged under the quiet wakefulness, pre-running sleep, forward running and post-

running sleep conditions (138 cells, 3 animals). 18% of cells showed high calcium activity

during pre-running sleep (∆F pre-run sleep /∆F quiet > 1.5) and were removed from the analysis of

neuronal reactivation during post-running sleep in Fig. 4F. As shown in Fig. 4F, cells highly

activated during forward running but not during the pre-running sleep (∆F running / ∆F quiet > 1.5;

∆F pre-run sleep /∆F quiet < 1.5) were reactivated during the post-running sleep. In contrast, cells with

no or moderate increase (< 50%) in somatic calcium level during forward running and the pre-

running sleep (∆F running / ∆F quiet < 1.5; ∆F pre-run sleep /∆F quiet < 1.5) were not active during the

post-running sleep.

Fig. S8. New spine formation after training at different time of the day. Spine formation on

HFBs is slightly but not significantly lower in mice trained at the beginning of the wake cycle

than in mice trained at the beginning of the sleep cycle (P = 0.19). Because mice sleep less

during the wake cycle than during the sleep cycle, this result suggests that the degree of spine

formation is not proportional to the duration of post-training sleep. Because the transcription and

translation of many genes involved in synaptic plasticity express circadian oscillations, the

effectiveness of sleep in promoting spine formation may vary at different time of the day.

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