Noninvasive optical inhibition with a red-shifted microbial rhodopsin

Abstract

Optogenetic inhibition of the electrical activity of neurons enables the causal assessment of their contributions to brain functions. Red light penetrates deeper into tissue than other visible wavelengths. We present a red-shifted cruxhalorhodopsin, Jaws, derived from Haloarcula (Halobacterium) salinarum (strain Shark) and engineered to result in red light-induced photocurrents three times those of earlier silencers. Jaws exhibits robust inhibition of sensory-evoked neural activity in the cortex and results in strong light responses when used in retinas of retinitis pigmentosa model mice. We also demonstrate that Jaws can noninvasively mediate transcranial optical inhibition of neurons deep in the brains of awake mice. The noninvasive optogenetic inhibition opened up by Jaws enables a variety of important neuroscience experiments and offers a powerful general-use chloride pump for basic and applied neuroscience.

Full text

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe ADVANCE ONLINE PUBLICATION 1
t e C h n I C a l r e p O r t S
Optogenetic inhibition of the electrical activity of neurons
enables the causal assessment of their contributions to brain
functions. Red light penetrates deeper into tissue than other
visible wavelengths. We present a red-shifted cruxhalorhodopsin,
Jaws, derived from Haloarcula (Halobacterium) salinarum
(strain Shark) and engineered to result in red light–induced
photocurrents three times those of earlier silencers. Jaws
exhibits robust inhibition of sensory-evoked neural activity in the
cortex and results in strong light responses when used in retinas
of retinitis pigmentosa model mice. We also demonstrate that
Jaws can noninvasively mediate transcranial optical inhibition
of neurons deep in the brains of awake mice. The noninvasive
optogenetic inhibition opened up by Jaws enables a variety
of important neuroscience experiments and offers a powerful
general-use chloride pump for basic and applied neuroscience.
Optogenetic inhibition, the use of light-activated ion pumps to enable
transient activity suppression of genetically targeted neurons by
pulses of light
1–3
, is valuable for the causal parsing of neural circuit
component contributions to brain functions and behaviors. A major
limit to the utility of optogenetic inhibition is the addressable quantity
of neural tissue. Previous optogenetic hyperpolarizing proton pumps
(Arch
1
, ArchT
3
, Mac
1
) and chloride pumps (eNpHR
4
, eNpHR3.0 (ref. 2))
have successfully inhibited volumes of approximately a cubic
millimeter, but many neuroscience questions require the ability to sup-
press larger tissue volumes. A number of pharmacogenetic, chemical
and genetic strategies have been used for this purpose
5–7
, but it would
ideally be possible to address these large brain volumes with the
millisecond temporal precision of optogenetic tools.
Another common desire in optogenetic experiments is to
minimize invasiveness from inserting optical fibers into the brain,
which displaces brain tissue and can lead to side effects such as
brain lesion, neural morphology changes, glial inflammation and
motility, or compromise of asepsis
8–10
. Less invasive strategies that do
not require an implanted optical device would also increase experi-
mental convenience and enable longer timescale experiments than
often feasible with fragile implants. While a number of previous studies
using channelrhodopsins have attempted to address this problem
11–16
,
noninvasive optical inhibition has not yet been possible.
To enable noninvasive large-volume optogenetic inhibition, we engi-
neered and characterized Jaws, a spectrally shifted cruxhalorhodopsin
derived from the species H. salinarum (strain Shark)
17
, which mediates
strong red light–driven neural inhibition. Jaws is capable of powerful
optical hyperpolarization in a variety of neuroscientific contexts: it
successfully enabled suppression of visually evoked neural activity in
mice, functioned in cone photoreceptors to restore greater light sensi-
tivity in mouse models than possible with previous opsins and enabled
the noninvasive transcranial inhibition of neurons in brain structures
up to 3 mm deep. This new reagent thus makes a variety of important
experiments amenable to optogenetic investigation.
RESULTS
Engineering a red light–sensitive chloride pump
In earlier work, we identified two cruxhalorhodopsins from the haloarcula
H. marismortui and H. vallismortis that possessed the most red-shifted
action spectra known for any hyperpolarizing opsins
1,18
. While their low
photocurrents made them poor candidates for in vivo use
1
, their spectra
suggested they might be good scaffolds for further engineering. Because red
light is less absorbed by hemoglobin than blue, green or yellow wavelengths,
we reasoned this red-light sensitivity might render deep brain regions more
accessible. We validated this through Monte Carlo modeling and direct
measurement in the live mouse brain (Supplementary Fig. 1).
We therefore screened members of the cruxhalorhodopsin class in
primary neuronal culture to identify molecules with both red-shifted
1
Media Lab, Department of Media Arts and Sciences, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA.
2
McGovern Institute for Brain
Research, Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA.
3
Department of Biological
Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA.
4
Department of Neurobiology, Yale School of Medicine, Yale University,
New Haven, Connecticut, USA.
5
Neural Circuit Laboratories, Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland.
6
Department of Genetics,
Harvard Medical School, Boston, Massachusetts, USA.
7
Picower Institute for Learning and Memory, Department of Brain and Cognitive Sciences, Massachusetts
Institute of Technology, Cambridge, Massachusetts, USA.
8
George W. Woodruff School of Mechanical Engineering, Georgia Institute of Technology, Atlanta, Georgia,
USA.
9
Department of Biomedical Engineering, Boston University, Boston, Massachusetts, USA.
10
Department of Bioengineering, University of Pennsylvania,
Philadelphia, Pennsylvania, USA.
11
Kavli Institute for Neuroscience, Yale University, New Haven, Connecticut, USA.
12
These authors contributed equally to
this work. Correspondence should be addressed to E.S.B. (esb@media.mit.edu).
Received 28 February; accepted 1 June; published online 6 July 2014; doi:10.1038/nn.3752
Noninvasive optical inhibition with a red-shifted
microbial rhodopsin
Amy S Chuong
1–3
, Mitra L Miri
4,12
, Volker Busskamp
5,6,12
, Gillian A C Matthews
7,12
, Leah C Acker
1–3,12
,
Andreas T Sørensen
2
, Andrew Young
2
, Nathan C Klapoetke
1–3
, Mike A Henninger
1–3
, Suhasa B Kodandaramaiah
1–3,8
,
Masaaki Ogawa
1–3
, Shreshtha B Ramanlal
9
, Rachel C Bandler
1
, Brian D Allen
1
, Craig R Forest
8
, Brian Y Chow
10
,
Xue Han
9
, Yingxi Lin
2
, Kay M Tye
7
, Botond Roska
5
, Jessica A Cardin
4,11
& Edward S Boyden
1–3

© 2014 Nature America, Inc. All rights reserved.
2 ADVANCE ONLINE PUBLICATION nature neurOSCIenCe
t e C h n I C a l r e p O r t S
action spectra and robust photocurrents (Fig. 1a and Supplementary
Fig. 2) and identified a hyperpolarizer we named Halo57, the crux-
halorhodopsin from H. salinarum (strain Shark), with less than 60%
homology to the Natronomonas pharaonis halorhodopsin (NpHR/
halo; Supplementary Fig. 3a). We subsequently engineered Halo57 by
identifying K200R and W214F point mutations
19,20
(Fig. 1b), which
significantly boosted photocurrents without altering its red action spec-
trum (n = 9 or 10 cells; P = 0.02, ANOVA with Dunnett’s post hoc test),
and by appending the KGC
21
or ER2 (ref. 22) trafficking sequences
from the potassium channel Kir2.1 to result in the final molecules Jaws
(KGC + ER2) and Jaws-ER2 (ER2) (Fig. 1c and Supplementary Fig. 3b).
In accord with the low sequence conservation, the two homologous
Figure 1 Engineering and in vitro
characterization of Jaws, a red-shifted
cruxhalorhodopsin. (a) Red light photocurrents
(left y-axis, red bars) and photocurrent densities
(right y-axis, white bars) from cruxhalorhodopsins
and the N. pharaonis halorhodopsin in cultured
neurons (632 nm, 5 mW/mm
2
; n = 6 cells for
each). ATCC, American Type Culture Collection;
DSM, German Collection of Microorganisms and
Cell Cultures. (b) The K200R W214F Halo57
mutant (n = 9 cells) demonstrates enhanced
photocurrents over wild-type Halo57 in cultured
neurons (n = 10 cells; 632 nm, 5 mW/mm
2
).
(c) Photocurrents (black bars) and photocurrent
densities (white bars) for wild-type Halo57
(n = 9 cells), Halo57 (K200R W214F) (n = 21
cells), Jaws (n = 11 cells) and Jaws-ER2 in cultured
neurons (n = 5 cells; 5 mW/mm
2
, 543 nm;
P < 0.01 for all variants compared to the
wild-type Halo57). (d) Action spectra for Jaws
(n = 12 cells), ArchT (n = 13 cells) and eNpHR3.0
(n = 15 cells; measured in cultured neurons
using equal photon fluxes of ~3.0 × 10
21
photons s
−1
m
−2
). (e) Red:green photocurrent
ratios for Jaws (n = 33 cells), ArchT (n = 36 cells)
and eNpHR3.0 in cultured neurons (n = 29 cells; 5 mW/mm
2
at 632 nm or 543 nm). Regression lines are shown for each opsin, indicating distinct
spectral shifts. (f) Red-light mediated photocurrents (left y-axis, red bars) and photocurrent densities (right y-axis, white bars) for Jaws (n = 33 cells),
ArchT (n = 36 cells) and eNpHR3.0 in cultured neurons (n = 29 cells; 5 mW/mm
2
at 632 nm; P < 0.001 for Jaws compared to ArchT or eNpHR3.0).
Data in each panel were measured from 2 batches of neuron culture. Values throughout are mean ± s.e.m. In b, *P = 0.020; in c, from left to right,
**P = 0.0017, **P = 0.0047, *P = 0.0183; in f, ***P < 0.001. Unpaired t-tests in b, ANOVA in c with Dunnett’s post hoc test using wild-type Halo57 as
the reference, ANOVA in f with Newman-Keuls post hoc test. t = 2.567 for b; F = 5.473 for c and 14.02 for f.
cb
ON cell
100 µV
Full-eld red light
100 µV
OFF cell
Full-eld red light
a
GFP mCAR
Merged
0 100 200 300 400 500
Min
WT
Max
WT
0
50
100
eNpHR
Halo57
Jaws
ArchT
Mac
Peak spike frequency (Hz)
Fraction of cells (%)
d
eNpHR
0
100
200
300
****
untreated f-RD
spontaneous ring
Mean ganglion
cell spiking in
opsin-expressing
f-RD retinas (Hz)
***
Halo57
Jaws
ArchT
Mac
eNpHR
Halo57
Jaws
ArchT
Mac
470-nm log
light intensity
(photons cm
–2
s
–1
)
Blue-light-driven
ganglion cell spiking
15 16 17 18 19
Green-light-driven
ganglion cell spiking
550-nm log
light intensity
(photons cm
–2
s
–1
)
15 16 17 18 19
***
****
****
e
600-nm log
light intensity
(photons cm
–2
s
–1
)
15 16 17 18 19
0
100
200
***
****
****
Mean ganglion cell spiking
in opsin-expressing
f-RD retinas (Hz)
Red-light-driven
ganglion cell spiking
Figure 2 Jaws-mediated light responses
in mouse retinitis pigmentosa retinas.
(a) Confocal fluorescence images from
Jaws-GFP expressing retina with the fast
form of retinal degeneration (f-RD),
stained for GFP (green) and mouse
cone arrestin (mCAR) (magenta),
4 weeks after injection with
AAV8-mCAR-Jaws-GFP virus. Scale
bar, 20 µm. (b) Raw traces recorded from
ON and OFF retinal ganglion cells in retinas
expressing Jaws in cone photoreceptors, optically stimulated with 9.6 × 10
17
photons cm
−2
s
−1
at 600 nm for 1 s. (c) Comparison of mean spiking in
ganglion cells downstream of neurons expressing eNpHR (n = 21 cells from 1 mouse), Halo57 (n = 14 cells from 1 mouse), ArchT (n = 30 cells from
1 mouse), Mac (n = 13 cells from 2 mice) or Jaws (n = 27 cells from 1 mouse) at opsin peak wavelength sensitivity. Light intensity was 1.2 × 10
18
photons
cm
−2
s
−1
for green light (ArchT, Mac) and 9.6 × 10
17
photons cm
−2
s
−1
for red light (Jaws, eNpHR, Halo57). Dotted line indicates baseline f-RD firing per
ref. 49. (d) Population distribution of retinal ganglion cell peak firing rates at opsin peak wavelength sensitivity. Light intensities as indicated in c; dotted
lines indicate the minimum (min
WT
) and maximum (max
WT
) of the wild-type dynamic range
24
. (e) Retinal ganglion spike rate versus red, green and blue
irradiances, measured in ganglion cells downstream of opsin-expressing cones in f-RD retina. Values are means ± s.e.m.; n values for c–e are as indicated
in c. In c–e, ***P < 0.001, *P < 0.0001. ANOVAs in c,e with Dunnett’s post hoc test using eNpHR as the reference; Kolmogorov-Smirnov test in d. F =
22.13 for c and 37.08, 32.53 and 10.43 for e. In c, ****P < 0.0001 for Jaws, ***P < 0.001 for ArchT, P = 0.9973 for Halo57 and P = 0.4929 for Mac.
In d, as compared to Jaws, P < 0.0001 for eNpHR, P < 0.0001 for Halo57, P = 0.0375 for ArchT and P = 0.0008 for Mac. In e, P = 0.4679, P = 0.001,
P < 0.0001, P < 0.0001 for green and P = 0.2703, P = 0.001, P < 0.0001 and P < 0.0001 for red, from lowest to highest irradiance.
d
400 500 600 700
0
0.5
1.0
Jaws eNpHR3.0
ArchT
Wavelength (nm)
Peak-normalized
photocurrent
Photocurrent
Photocurrent density
a
0
50
100
150
0
1
2
3
Cruxhalorhodopsins
N. pharaonis
, Halo/NpHR
H. marismortui
ATCC 43049
Halomicrobium
mukohataei
DSM 12286
H. californiae
ATCC 33799
H. sinaiiensis
ATCC 33800
H. salinarum
(strain Shark)
H. salinarum
(strain Port)
Photocurrent (pA)
Photocurrent density
(pA pF
–1
)
e
0 500 1,000 1,500
0
500
1,000
Jaws eNpHR3.0
ArchT
632-nm photocurrent (pA)
543-nm photocurrent
(pA)
b
Halo57 (wild type)
Halo57 (K200R W214F)
0
100
200
300
400
0
5
10
*
Photocurrent (pA)
Photocurrent density
(pA pF
–1
)
f
0
100
200
300
400
0
2
4
6
8
Photocurrent (pA)
Photocurrent density
(pA pF
–1
)
***
Jaws
eNpHR3.0
ArchT
Photocurrent
Photocurrent density
c
Halo57 (wild type)
Halo57 (K200R W214F)
Jaws-ER2
Jaws
0
100
200
300
400
500
0
5
10
**
**
*
Photocurrent (pA)
Photocurrent density
(pA pF
–1
)

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe ADVANCE ONLINE PUBLICATION 3
t e C h n I C a l r e p O r t S
point mutations did not enhance the N. pharaonis halorhodopsin
(Supplementary Fig. 3a,c,d). Ion-specific solutions confirmed Jaws
to be a light-driven chloride pump (Supplementary Fig. 3e,f).
In vivo optogenetic inhibition has previously only been possible with
blue, green and yellow light, as existing optogenetic inhibitors oper-
ate at only 10–30% of their peak capacity at red wavelengths. Because
Jaws has a 14-nm red shift relative to the N. pharaonis halorhodopsin
(NpHR/halo, eNpHR, eNpHR3.0), we hypothesized that it might be a
good candidate for potent red-light neural inhibition (Fig. 1d,e). To
further characterize this, we transfected Jaws, eNpHR3.0 or ArchT into
primary neuron cultures along with a secondary cytosolic tdTomato
plasmid and selected cells for whole-cell patch clamp characterization
solely on the basis of the presence of tdTomato (Supplementary Fig. 4),
to prevent selection bias for opsin expression levels and more accu-
rately represent the full range of in vivo expression: we found Jaws
to have significantly higher red light (632 nm) photocurrents
than eNpHR3.0 or ArchT across all tested light powers (Fig. 1f
and Supplementary Fig. 4d; n = 21–30 cells from 2 batches of culture;
P < 0.01 for 0.1–20 mW/mm
2
, ANOVA with Newman-Keuls post hoc
test). The robust in vitro photocurrents we observed seemed promis-
ing, so we next moved to assess Jaws in a variety of in vivo contexts.
Potential for optogenetic visual therapeutics
Optogenetic neural hyperpolarization is being explored not only as
a tool for basic neuroscience, but also as a prototype therapeutic.
As one example, retinitis pigmentosa is a visual disorder that results
first in night blindness and then in overall blindness as a result of
photoreceptor degeneration
23
. A potential therapy for patients with
cone photoreceptor atrophy would be to resensitize the cone cells
to light by genetically expressing light-activated hyperpolarizing ion
pumps in the cone photoreceptors, which in their healthy state hyper-
polarize in response to optical stimulation.
When expressed in retinal cones of retinitis pigmentosa mouse
models, eNpHR has previously been shown to be capable of transduc-
ing spikes in downstream retinal ganglion cells and mediating visually
guided behaviors. However, the resultant spiking rates are limited to
less than 200 Hz, substantially less than the full wild-type range of
15–450 Hz (ref. 24), and thus potentially limit fine visual perception
in future clinical use. Additionally, while eNpHR can be activated
using light powers that are safe for human use
24
, more light-sensitive
hyperpolarizers would require less light for stimulation and thus
provide a greater margin of efficacy and safety given the inevitable
unknowns that might crop up in potential future trials in humans.
Thus, better neural hyperpolarizers remain of great interest for poten-
tial clinical use in the human eye.
We decided to first compare Jaws’s performance against those of
other known hyperpolarizers by injecting adeno-associated virus
(AAV), serotype 8 (AAV8), encoding various light-driven proton or
chloride pumps into the cone photoreceptors of Pde6b
rd1
mice, also
called fast retinal degeneration mice
24,25
(Fig. 2a and Supplementary
Fig. 5b), which are blind by postnatal day (P) 28. We conducted
extracellular retinal ganglion cell recordings (Fig. 2b) 4–6 weeks after
–1 0 1 2
0
1
2
Time (s)
Firing rate (Hz)
Visual stimuli
1 s light
–1 0 1 2
0
2
4
6
Firing rate (Hz)
Visual stimuli
1 s light
–1 0 1 2
0
2
4
6
Firing rate (Hz)
Visual stimuli
a
–1 0 1 2
0
1
2
Time (s)
Firing rate (Hz)
Visual stimuli
b
f
12
8
4
0
10.50–0.5
Time (s)
Firing rate (Hz)
Square pulse
Ramp
eNpHR
eNpHR3.0
Arch
ArchT
Jaws
0
50
100
**
Inhibition of
evoked response (%)
c d
0 25 50 75 100
0
50
100
Contrast strength (%)
Suppression (%)
eNpHR3.0
ArchT
Jaws
e
0
2
4
6
*
Firing rate (Hz)
Spontaneous
Rebound
h
Rebound delay (ms)
g
Peak firing rate (Hz)
0
80
100
60
40
20
**
***
0
100
150
50
Square pulse
Ramp
Square pulse
Ramp
Figure 3 Jaws-mediated inhibition of evoked responses in visual cortex. (a) Representative rasters of visually evoked responses (top left) and
Jaws-mediated inhibition of visually evoked responses (top right), and post-stimulus time histograms for visually evoked (bottom left) and Jaws-inhibited
visually evoked (bottom right) responses, of a representative neuron (n = 27) in the visual cortex of an anesthetized Emx1-cre mouse, as measured by
extracellular tetrode recording (AAV5-FLEX virus; 35 mW/mm
2
at 593 nm using a 200-µm fiber). (b) Population average of five simultaneously recorded
neurons showing visually evoked responses (left) and Jaws-mediated inhibition of visually evoked responses (right). (c) Inhibition of visually evoked
neural responses for eNpHR (n = 8 units from 2 mice), eNpHR3.0 (n = 32 units from 4 mice), Arch (n = 18 units from 3 mice), ArchT (n = 21 units
from 3 mice) and Jaws (n = 27 units from 4 mice; P < 0.01 for Jaws versus eNpHR3.0). (d) Jaws-mediated inhibition of visually evoked responses
(n = 14 units from 2 mice) for different visual input stimulus strengths. (e) Comparison of spontaneous and immediately post-illumination firing rates
for Jaws, eNpHR3.0 and ArchT (n = 32 units from 4 mice for eNpHR3.0, n = 21 units from 3 mice for ArchT, n = 5 units for Jaws from 2 mice;
P < 0.05 for Jaws versus eNpHR3.0). (f) Post-stimulus time histogram for a standard, step light pulse (black line) versus ramped illumination
(yellow line), for a spontaneously firing visual cortex neuron. (g,h) Comparison of peak firing rates (g) and rebound delay rates (h) for step versus
ramped illumination (n = 16 units from 3 mice). Values throughout are mean ± s.e.m. For c,e,g,h, *P < 0.05, **P < 0.01, ***P < 0.001. ANOVAs with
Newman-Keuls post hoc tests in c,e; paired t-tests in g,h. F = 7.379 for c, 2.826 for e; t = 4.485 for g and 9.078 for h.

© 2014 Nature America, Inc. All rights reserved.
4 ADVANCE ONLINE PUBLICATION nature neurOSCIenCe
t e C h n I C a l r e p O r t S
injection (P64–75) and found peak wavelength photostimulation of
Jaws-expressing photoreceptors to induce significantly more retinal
ganglion spiking, indicated by mean spiking frequencies greater than
those for eNpHR, ArchT, Mac or the wild-type Halo57 (Fig. 2c and
Supplementary Fig. 5a). Jaws also enabled a broader peak spike fre-
quency distribution than other opsins (P < 0.05 for Jaws versus all
other opsins, Kolmogorov-Smirnov test), with a mean frequency of
67.8 ± 7.0 Hz with eNpHR versus 189.5 ± 15.9 Hz with Jaws (values
throughout are mean ± s.e.m. unless otherwise stated), increasing
the spiking bandwidth by threefold (Fig. 2d). We additionally
observed Jaws to mediate the highest ganglion spiking rates over
various irradiances of 600 nm, 550 nm and 470 nm light (Fig. 2e
and Supplementary Fig. 5c,d), indicating its high light sensitivity,
which alongside Jaws’s spectral peak at 600 nm may confer benefits
because of the relative safety of 600-nm light
26
. From this comparison
of light-sensitive hyperpolarizers, we conclude that Jaws may repre-
sent an optogenetic reagent with molecular properties better suited
than previous opsins for therapeutic cone reactivation for a subset of
patients with retinitis pigmentosa.
In vivo suppression of visually evoked cortical activity
We next evaluated the performance of Jaws in a common neuroscien-
tific experimental context: the suppression of stimulus-evoked neural
activity. Given the proximity of yellow light to all existing inhibitory
opsins’ peak excitation wavelengths
1,2
, we decided, for this specific
experiment, to directly compare Jaws’s in vivo performance in the mouse
brain against that of other inhibitors, using yellow light. Although
all previous studies characterizing optogenetic hyperpolarizers
have focused on inhibiting spontaneous neural activity
1–4
, many
neuroscientific questions focus on stimulus-evoked or event-associated
neural activity. The ability to suppress the activity of a given cell type
responding to behaviorally relevant inputs is therefore critical.
To assess Jaws’s potential as a general-purpose inhibitor of stimulus-
evoked neural activity, we injected a Cre recombinase–dependent
AAV, serotype 5 (AAV5-FLEX), expressing eNpHR, eNpHR3.0, Arch,
ArchT or Jaws into the primary visual cortex of Emx1-cre transgenic
mice and delivered visual stimuli while delivering yellow light with an
optetrode (593 nm; 35 mW/mm
2
out of a 200 µm fiber tip). Inhibition
of visually evoked neural activity was strong (Fig. 3a,b), with an
86 ± 3% reduction of visually evoked activity in Jaws-expressing
cortex (Fig. 3c; from 4 mice, n = 27 units with a significant reduction in
firing; n = 3 units showed no change and n = 3 units showed a signifi-
cant increase in firing) over a range of evoked firing rates (1.8–12.5 Hz;
Supplementary Fig. 6a), and consistent across a range of input
contrast strengths (Fig. 3d). This comparison of neural inhibitors
thus reveals that Jaws is capable of mediating excellent inhibition of
evoked neural activity in a biologically meaningful context.
A rebound burst of action potentials is common after illumination
of cells expressing halorhodopsins
18,27–30
or archaerhodopsins
18,27,31
,
which respectively pump chloride inward and protons outward.
A variety of possible mechanisms have been proposed, including
hyperpolarization-activated I
h
currents
29,30,32
or changes in chloride
reversal potential due to intracellular chloride accumulation
33
. We
similarly observed post-illumination rebound in Jaws-, eNpHR3.0-
and ArchT-transduced neurons, and Jaws produced greater effects
than eNpHR3.0 or ArchT (Fig. 3e; n = 8–32 units; P < 0.05, ANOVA
with Newman-Keuls post hoc test). We attempted to ameliorate this by
gradually ramping down illumination over a duration of 200–1,000 ms
Figure 4 Jaws-mediated red light inhibition in
rodent cortex. (a) Confocal fluorescence images
from Jaws-GFP–expressing motor cortex,
6 weeks after injection. Outline indicates brain
boundary
50
. Scale bar, 1 mm (top) or 100 µm
(bottom). (b) Representative current-clamp
recording of Jaws-expressing neuron undergoing
optically evoked (top, 632 nm, 5 mW/mm
2
) or
electrically evoked (bottom) hyperpolarization
in acute cortical slice 6 weeks after injection.
Red bar indicates optical illumination; blue
trace indicates electrical current injection.
(c) Quantification of Jaws photocurrents as
a function of red light irradiance in acute
slice (n = 16 cells from 4 mice). (d) Confocal
fluorescence images of Jaws-GFP-, Jaws-
GFP-ER2- or eNpHR3.0-expressing motor
cortex. Scale bars, 100 µm. (e) Side-by-side
comparison of red light–driven inhibition of
spontaneous neural activity in motor cortex
(n = 6 units from 2 mice for each opsin; light
powers were measured at the tip of a 200-µm
fiber; 637 nm). (f) Red-light inhibition was
equivalent to yellow-light inhibition for Jaws
(n = 5 units from 2 mice; P = 0.4685)
and Jaws-ER2 (n = 6 units from 2 mice;
P = 0.1952), but substantially less potent
for eNpHR3.0 (n = 6 units from 2 mice;
P = 0.0121). (g) Red light efficaciously
inhibited neurons expressing Jaws (n = 5 units
from 2 mice) and Jaws-ER2 (n = 6 units
from 2 mice) but not eNpHR3.0 (n = 8 units
from 2 mice) over a range of firing rates. Values throughout are mean ± s.e.m. In e,f, n.s., not significant; *P < 0.05, **P < 0.01, ***P < 0.001,
****P < 0.0001. Unpaired t-test in e, paired t-test in f. From left to right, t = 0.2902, 3.264, 3.494, 2.422, 5.422, 11.4, 9.275 and 4.338 for e and
0.6205, 3.843 and 1.496 for f. In e, P = 0.031, 0.0068, 0.0452, 0.0016, <0.0001, <0.0001 and 0.0025 from lowest to highest irradiance. In f,
P = 0.4685 for Jaws, P = 0.1952 for Jaws-ER2 and *P = 0.0121 for eNpHR3.0.
c
0.01 0.1 1 10 100
0
500
1,000
632-nm irradiance (mW/mm
2
)
Photocurrent (pA)
e
1 10 100 1,000
0
50
100
eNpHR3.0Jaws
*
**
**
*
****
****
**
Red light power out of fiber tip
(mW/mm
2
)
Inhibition (%)
eNpHR3.0
*
n.s.
0
25
50
75
100
593 nm
130 mW/mm
2
637 nm
130 mW/mm
2
593 nm
130 mW/mm
2
593 nm
400 mW/mm
2
637 nm
450 mW/mm
2
Jaws
Inhibition (%)
f
n.s.
Jaws-ER2
0 5 10 15
0
5
10
15
Jaws (130 mW/mm
2
)
Jaws-ER2 (130 mW/mm
2
)
eNpHR3.0 (450 mW/mm
2
)
eNpHR3.0 (130 mW/mm
2
)
Baseline firing (Hz)
Red-illumination firing
rate (Hz)
g
a
d
Jaws eNpHR3.0Jaws-ER2
b
1 s
20 mV
–67 mV
–67 mV
300 pA
637 nm
130 mW/mm
2

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe ADVANCE ONLINE PUBLICATION 5
t e C h n I C a l r e p O r t S
and found that the mean rebound instantaneous firing rate dropped
significantly, from 52 ± 12 Hz to 23 ± 6 Hz (n = 16 units; P = 0.0065,
paired t-test), suggesting that sculpting of light pulses may enable
rebound reduction. Notably, the rebound delay also significantly
lengthened, from 18 ± 6 ms to 76 ± 6 ms (n = 16 units; P < 0.001,
paired t-test) and became more temporally dispersed (Fig. 3f–h and
Supplementary Fig. 6b). Like all previous inhibitors, Jaws does yield
post-illumination rebound, which likely has implications for careful
experiment design comparable to those for earlier inhibitors such as
eNpHR3.0. Further attention to light-pulse shape may be merited as
a way to reduce rebound for all optogenetic inhibitors.
Red-light performance of Jaws
We next assessed Jaws’s red-light properties via whole-cell patch-clamp
recordings in acute cortical slices, as well as extracellular recordings
in awake, head-fixed mice. We injected AAV8 encoding Jaws under
the Ca
2+
/calmodulin-dependent protein kinase (Camk2a) promoter
or a truncated human synapsin (SYN1) promoter, hereafter referred
to as hSyn
16
, or Jaws-ER2 under the hSyn promoter, into motor cortex
and observed robust red light photocurrents, hyperpolarization and
neural inhibition (Fig. 4a–c and Supplementary Fig. 7a,c), with
~95% reduction of neural activity achievable in neurons illuminated
by a 200-µm fiber (130 mW/mm
2
fiber tip irradiance at 635 nm;
Supplementary Fig. 8). Jaws’s kinetic properties were comparable to
those of other optogenetic inhibitors (Supplementary Figs. 4e and 7c),
and its expression did not alter basal cell properties in cultured cortical
neurons (Supplementary Fig. 4g), or in cortical or dentate granule
neurons in acute brain slice (Supplementary Figs. 7d and 9).
Having established Jaws as a potent red-light-drivable inhibitor, we
next assessed its in vivo performance compared to that of eNpHR3.0.
We did titer-matched injections of 2 × 10
9
viral particles of AAV8-
hSyn-Jaws, AAV8-hSyn-Jaws-ER2 or AAV8-hSyn-eNpHR3.0 into
mouse motor cortex (Fig. 4d) and performed awake head-fixed record-
ings to characterize each hyperpolarizer over different red (637-nm)
light powers (Fig. 4e). Yellow light (593 nm) was used as a positive
control to identify opsin-expressing neurons, owing to its proximity to
both Jaws’s and eNpHR3.0’s spectral peaks, and we conducted paired
recordings of individual opsin-expressing neurons to assess their
b
c
a
f
2 mV
e
2.01.51.00.50–0.5–1.0
Time (s)
Average spike
frequency (Hz)
4
3
2
1
0
10 sweeps
d
i
Transcranial
1 mV
100 µV
1,800 µm
1,300 µm
5 s 5 s
5 s 5 s
200-µm fiber
g
0 1,000 2,000 3,000
0
25
50
75
100
Transcranial
200-µm fiber
Distance from brain surface (µm)
Suppression (%)
h
0 10 20
0
10
20
Non-opsin-expressing cells
(transcranial illumination)
Transcranial
Baseline firing rate (Hz)
Illumination firing
rate (Hz)
200-µm fiber
j
Transcranial
0
25
50
75
100
n.s.
Suppression (%)
200-
µm fiber
0 1,000 2,000 3,000
0
25
50
75
100
Transcranial
Suppression (%)
200-µm fiber
Distance from brain surface (µm)
Bregma + 1.78 mm
1 mm
0 5 10 15 20 25
0
10
20
30
Time (s)
Average spike
frequency (Hz)
Figure 5 Noninvasive red-light inhibition of neural activity.
(a) Schematic of noninvasive red light delivery through the
intact skull. (b,c) Fluorescence images from mouse insular cortex (b; scale bar, 1 mm)
and motor cortex (c; scale bars, 25 µm) 6 weeks after injection with AAV8-hSyn-Jaws
virus. Outline indicates brain boundary
50
. (d) Raster plot (top) and population average
(bottom) of transdural inhibition of neurons in the medial prefrontal cortex of anesthetized
mice, showing light-induced suppression 6–8 weeks after injection with AAV8-hSyn-
Jaws virus (n = 26 units from 2 mice; red bar, 1 s pulse, 25 mW/mm
2
fiber tip irradiance,
635 nm). Red bars denote red light illumination. Vertical red lines in f also demarcate the illumination period. (e) Representative glass-pipette
extracellular recording (n = 13) of a transcranially illuminated cortical neuron 2,700 µm below the brain surface in an awake mouse (red bar,
10 mW/mm
2
fiber tip irradiance, 635 nm; 4 weeks after injection with AAV8-hSyn-Jaws virus). (f) Spike rasters taken from a representative neuron (top)
and population average (bottom; n = 13 units from 6 mice) of instantaneous firing rate in neurons from awake mice undergoing transcranial light-induced
suppression 4–8 weeks after injection with AAV8-hSyn-Jaws virus (5 s pulse, 10 mW/mm
2
635 nm fiber tip irradiance; black line, mean; gray lines, mean
± s.e.m.). (g) Percentage reduction in neural activity for neurons recorded at different depths in awake mouse brain, 6–8 weeks after injection
with AAV8-hSyn-Jaws virus. Red light delivered via traditional 200-µm fiber (n = 37 units from 11 mice) or transcranially (n = 13 units from 6 mice). (h)
Firing rate averaged over the illumination period versus baseline firing rate for neurons recorded in awake mouse brain (n = 37 units from 11 mice for
200-µm-fiber illumination; n = 13 units from 6 mice for transcranial illumination; n = 14 units from 5 mice for transcranial illumination of
non-opsin-expressing neurons). (i) Representative extracellular recordings of neurons 1,800 µm (top) and 1,300 µm (bottom) below the surface of
the awake brain undergoing illumination via a 200-µm fiber (red bar, ~130 mW/mm
2
, 635 nm) 500 µm above the electrode tip (left) or transcranially
(right) through the intact skull (~10 mW/mm
2
; 635 nm), recorded 8 weeks after injection with AAV8-hSyn-Jaws virus. (j) Transcranial and
200-µm-fiber illumination were equally efficacious (left) in inhibiting neurons (n = 8 units from 6 mice; P = 0.5894; n.s., not significant) over
depths of up to 3 mm (right). Values are means ± s.e.m. Paired t-test in j, t = 0.5655.

© 2014 Nature America, Inc. All rights reserved.
6 ADVANCE ONLINE PUBLICATION nature neurOSCIenCe
t e C h n I C a l r e p O r t S
performance under different wavelengths (Fig. 4f ). As expected, both
Jaws variants robustly inhibited spontaneous neural activity when illu-
minated with either red (88.8 ± 15.9% inhibition for Jaws, 97.2 ± 3.9%
inhibition for Jaws-ER2) or yellow wavelengths (94.2 ± 9.3% inhibition
for Jaws, 99.0 ± 1.6% inhibition for Jaws-ER2), while eNpHR3.0’s red-
light inhibition dropped significantly to only 27.5 ± 30.3% relative to its
peak yellow-light performance of 76.1 ± 11.1% inhibition (Fig. 4f; n = 5
or 6 units, P = 0.0121), and we found these red-light results to hold over
a range of firing rates (Fig. 4g; n = 5–8 units for each opsin).
Noninvasive optogenetic inhibition
Having demonstrated Jaws’s efficacy as a red-light-drivable reagent
in a variety of contexts, we next sought to determine its potency as a
noninvasive and long-distance neural inhibitor. Using red illumina-
tion (Fig. 5a), we found we could transdurally inhibit neurons in
the medial prefrontal cortex of a mouse, with a 86 ± 13% decrease
(mean ± s.d.) in firing for neurons showing suppression (n = 66 units
across 2 anesthetized mice; Fig. 5d). Using a similar strategy, we suc-
cessfully inhibited neurons in the awake mouse cortex by delivering
red light transcranially through the intact skull (635 nm, 10 mW/mm
2
out of a 1,500 µm fiber tip; Fig. 5a,b) to regions 1–3 mm below the
brain’s surface (Fig. 5c,e,f). Transcranial inhibition in the motor, som-
atosensory, insular and piriform cortices was efficient, with a 92 ± 14%
decrease in firing for neurons showing suppression (mean ± s.d.;
from 7 mice, n = 13 units; n = 9 units showed no change and n = 3
units showed an increase in activity; Fig. 5f).
The efficacy of transcranial optogenetic inhibition was similar across
several millimeters of the awake mouse brain (Fig. 5g) and over a
variety of baseline firing rates (Fig. 5h). To directly address the pos-
sibility that opsin-expressing apical dendrites near the brain’s surface
were being hyperpolarized, resulting in somatic or downstream net-
work inhibition, we next conducted paired recordings using a standard
acutely inserted 200-µm fiber as a positive control for transcranial illu-
mination outside the mouse’s intact skull. Illumination from a 200-µm
fiber has been shown to fall off rapidly within a few hundred microns
of the fiber tip
34,35
, and the likelihood of optogenetic inhibition has
been similarly demonstrated to fall off over a similar distance
31
. Our
paired recordings revealed similar degrees of suppression over a range
of 1–3 mm below the brain’s surface (Fig. 5i,j). From this, we conclude
Jaws is capable of noninvasive optogenetic inhibition comparable to
the current standard of invasive optical delivery.
DISCUSSION
We here report Jaws, a red light sensitive opsin with the most
red-shifted spectrum of any optogenetic inhibitor known to us. Jaws
enabled efficacious transcranial inhibition of neural activity in awake
mouse brain in response to red light, with a similar efficacy to stand-
ard invasive light delivery with a 200-µm optical fiber. Additionally,
Jaws enabled significantly more inhibition of stimulus-evoked neu-
ral activity than possible with previous halorhodopsins. Finally, it
restored photosensory responses in retinitis pigmentosa retinas in a
fashion achieving greater spike rates than previously achievable and
with a more naturalistic frequency range, which may be important
for retinal neural coding. Thus, Jaws has demonstrated utility across
a wide variety of key neuroscience applications, both basic science
and preclinical. We also demonstrate that light pulse sculpting can
ameliorate the amplitude of the post-inhibition rebound that has been
widely reported for optogenetic inhibitors.
It has been previously well established that optical penetrance
of mammalian tissues in the red to near-infrared (600–1,000 nm)
wavelengths is better than that of bluer wavelengths, as a result of
substantially less absorption from hemoglobin, myoglobin and lipid.
Despite this better red light penetrance, however, myelination may still
affect optical propagation in neural tissue and should therefore be taken
into account given the experimental system in question
36,37
. The relative
inexpensiveness and stability of red light sources may additionally be of
use to experimental investigators, as well as the ability to avoid poten-
tially confounding experimental visual artifacts, since mouse visual pig-
ments peak at approximately 380 nm (ultraviolet cones) and 500 nm
(green cones), but mouse vision above 600 nm is poor
38,39
.
As others have reported, we observed a small number of neurons
across multiple in vivo experiments that increased firing during opti-
cal illumination delivered either transcranially or from a 200-µm
fiber, most likely as a result of network inhibition from upstream
neurons
3,40,41
. It is not possible to noninvasively illuminate only deep
volumes with our experimental setup without also targeting surface
and intermediate layers. We therefore cannot explicitly rule out the
possibility that the observed noninvasive results were the result of indi-
rect network activity or apical dendritic illumination: a caveat for any
in vivo investigation of noninvasive optogenetics. However, the robust
photocurrents we observed both in vitro and in acute slices over a
range of irradiances are consistent with and corroborate the functional
inhibition we observed in vivo in awake mice. We therefore believe
our paired recordings comparing transcranial illumination with local
200-µm fiber illumination to be highly suggestive of direct inhibition.
In scenarios where brain integrity is mandatory, noninvasive optoge-
netics may be invaluable in helping to prevent the many known issues
associated with implants
8–10
. The efficacy and simplicity of transcra-
nial inhibition could be helpful for chronic optogenetic experiments
involving long-term imaging
42,43
, longitudinal monitoring of disease
progression
44
, or developmental studies, during which developmental
changes in brain structures might preclude the chronic implantation
of a fiber at a given target
45
. In principle, the need for brain surgery
could be eliminated altogether by using transcranial illumination with
a Jaws-expressing transgenic mouse strain
27
. Neural inhibition in ani-
mals with large brains, such as the rhesus macaque, in which perturba-
tion of behavior with optogenetic stimulation or inhibition of neurons
has recently been demonstrated
46,47
, may particularly benefit from the
use of Jaws. Finally, with increasing interest in the potential for thera-
peutic optogenetics
24,48
, the opsin described here may prove useful in
the context of the many prototype therapies being explored.
METHODS
Methods and any associated references are available in the online
version of the paper.
Accession codes. GenBank: KM000925, KM000926, KM000927 and
KM000928.
Note: Any Supplementary Information and Source Data files are available in the
online version of the paper.
ACKNOWLEDGMENTS
We thank J. Juettner for help making AAV, and Y.K. Cho, D. Schmidt, F. Chen,
A. Beyeler, J.M. Zhuo and R.E. Kohman for advice and discussion. A.S.C.
acknowledges the Janet and Sheldon Razin
′
59 Fellowship of the Massachusetts
Institute of Technology (MIT) McGovern Institute. E.S.B. acknowledges Jerry
and Marge Burnett, the US Defense Advanced Research Projects Agency
Living Foundries Program HR0011-12-C-0068, Harvard/MIT Joint Grants
Program in Basic Neuroscience, Human Frontiers Science Program, Institution
of Engineering and Technology A F Harvey Prize, MIT McGovern Institute
and McGovern Institute Neurotechnology (MINT) Program, MIT Media Lab,
New York Stem Cell Foundation-Robertson Investigator Award, US National
Institutes of Health (NIH) Director’s New Innovator award 1DP2OD002002, NIH
EUREKA award 1R01NS075421, NIH grants 1R01DA029639, 1RC1MH088182

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe ADVANCE ONLINE PUBLICATION 7
t e C h n I C a l r e p O r t S
and 1R01NS067199, US National Science Foundation (NSF) CAREER award
CBET 1053233 and NSF grants EFRI0835878 and DMS0848804, the Skolkovo
Institute of Science and Technology, a Society for Neuroscience Research Award
for Innovation in Neuroscience (RAIN) and the Wallace H. Coulter Foundation.
M.L.M. acknowledges funding from NSF DGE 1122492. J.A.C. acknowledges
funding from the Whitehall Foundation, the Klingenstein Foundation, the
Swebelius Family Trust, the Simons Foundation, an Alfred P. Sloan Fellowship,
a NARSAD Young Investigator Award, a Smith Family Award for Excellence in
Biomedical Research, NIH R00 EY018407, NIH R01 EY022951 and NIH R01
MH102365. V.B. acknowledges Human Frontier Science Program, Swiss National
Science Foundation and Volkswagen Foundation fellowships. B.R. acknowledges
the Gebert-Ruf Foundation, SNSF, European Research Council, and European
Union SEEBETTER, TREATRUSH, OPTONEURO and 3X3D Imaging grants.
X.H. acknowledges funding from an NIH Director’s New Innovator Award
(1DP2NS082126), the NINDS (1R01NS087950, 1R21NS078660, 1R01NS081716),
NIMH (5R00MH085944), Pew Foundation, Alfred P. Sloan Foundation, Michael
J. Fox Foundation, and Brain and Research Foundation. Y.L. acknowledges
funding from NIH RO1 MH091220-01. B.Y.C. acknowledges funding from US
Defense Advanced Research Projects Agency Living Foundries, the US National
Science Foundation Biophotonics and the Brain Research Foundation. K.M.T.
acknowledges funding from the Whitehall Foundation, Klingenstein Foundation,
JPB Foundation, PIIF Funding, R01-MH102441-01 (NIMH) and DP2-OD-
017366-01. G.A.C.M. was supported by the Simons Center for the Social Brain.
AUTHOR CONTRIBUTIONS
A.S.C. and E.S.B. coordinated all experiments and data analysis. A.S.C. designed
and developed Jaws and cloned all constructs. A.S.C. performed in vivo glass
pipette extracellular recordings and in vitro electrophysiology. M.L.M. and
J.A.C. performed in vivo tetrode extracellular recordings. V.B. performed in vivo
multielectrode array recordings. A.S.C., G.A.C.M., A.T.S. and A.Y. performed
slice electrophysiology. A.S.C., M.L.M., G.A.C.M., A.T.S., J.A.C., V.B. and M.O.
performed in vivo viral injections. A.S.C., M.L.M., V.B., G.A.C.M., A.T.S., S.B.R.
and M.O. performed histological processing and fluorescence imaging. S.B.K. and
C.R.F. designed or performed autopatch experiments. A.S.C. and N.C.K. performed
transfections, cell culture and in vitro viral infections. M.A.H. conducted Monte
Carlo modeling. L.C.A. carried out light propagation measurements. R.C.B. and
B.D.A. carried out X-ray scans to measure mouse skull thicknesses for the Monte
Carlo model. A.S.C., M.L.M., V.B., G.A.C.M., A.T.S., B.Y.C., X.H., J.A.C., B.R. and
E.S.B. contributed to study design and data interpretation. J.A.C., B.R., K.M.T., Y.L.
and E.S.B. supervised all aspects of the work. A.S.C. and E.S.B. wrote the paper
with contributions from the other authors.
COMPETING FINANCIAL INTERESTS
The authors declare competing financial interests: details are available in the online
version of the paper.
Reprints and permissions information is available online at http://www.nature.com/
reprints/index.html.
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© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe
doi:10.1038/nn.3752
ONLINE METHODS
Animal procedures. All procedures were in accordance with the National
Institute for Laboratory Animal Research Guide for the Care and Use of
Laboratory Animals and approved by the Boston University Institute Institutional
Care and Use Committee (for extracellular red light experiments conducted
transcranially or with a 200 um fiber), the Massachusetts Institute of Technology
Committee on Animal Care (for all in vitro experiments, slice electrophysiology
and light propagation measurements) or the Yale University Institute Institutional
Care and Use Committee (for inhibition of evoked response and transdural
red light experiments). The Swiss Veterinary Office approved all animal experi-
ments and procedures for viral delivery of opto-genes and multi-electrode array
recordings (for the retina experiments). All mice in this study were housed
3–5 per cage, maintained with a 12-h light-dark cycle, and had no previous
experimental history.
Plasmid construction and site directed mutagenesis. Opsin genes were
mammalian codon–optimized and synthesized (Genscript). Point mutants
were generated using the QuikChange kit (Stratagene) on the opsin-GFP fusion
cassette in the pEGFP-N3 backbone (Invitrogen). Cultured neuron experi-
ments were carried out by subcloning all genes into a lentiviral backbone con-
taining the Camk2a promoter and with a C-terminal GFP fusion. AAV vectors
were constructed by subcloning opsin-GFP cassettes into AAV vectors behind
the Camk2a, CAG or hSyn truncated human synapsin (SYN1) promoters. For
cone photoreceptor delivery, opsin-GFP cassettes were subcloned into pAAV2-
mCAR-EGFP, replacing the EGFP gene
24
.
All constructs were verified by sequencing, and codon-optimized
sequences were submitted to GenBank (accession codes KM000925, KM000926,
KM000927 and KM000928) and made available at http://syntheticneurobiology.
org/protocols/.
In vitro culture, transfection and imaging. Hippocampal neuron cultures were
prepared from postnatal day 0 or 1 Swiss Webster mice (Taconic), plated at a den-
sity of 16,000–20,000 per glass coverslip coated with Matrigel (BD Biosciences)
51
,
and transfected at 3–5 d in vitro (DIV) using calcium phosphate (Invitrogen)
1,3
.
GFP fluorescence was used to identify transfected neurons for the cruxhalorho-
dopsin screen and HEK293FT characterizations; all other experiments were
conducted by cotransfecting with an independent tdTomato plasmid, in which
case neurons were picked based solely on tdTomato fluorescence.
Whole cell patch clamp recordings were made using a Multiclamp 700B,
a Digidata 1440 and a PC running pClamp (Molecular Devices). Neurons
were recorded 14–24 DIV, bathed in room temperature Tyrode’s solution
containing 125 mM NaCl, 2 mM KCl, 3 mM CaCl
2
, 1 mM MgCl
2
, 10 mM
HEPES, 30 mM glucose, 0.01 mM NBQX and 0.01 mM GABAzine (Sigma)
at pH 7.3 (NaOH-adjusted) and osmolarity 300 mOsm (sucrose-adjusted).
HEK cells were bathed in an identical Tyrode’s bath solution lacking GABAzine
and NBQX. No all-trans-retinal was supplemented for any recordings. 3–9 MΩ
borosilicate glass pipettes (Warner Instruments) were pulled with a P-97 micro-
pipette puller (Sutter Instruments) and filled with a solution containing (in mM)
125 potassium gluconate, 8 NaCl, 0.1 CaCl
2
, 0.6 MgCl
2
, 1 EGTA, 10 HEPES,
4 Mg-ATP and 0.4 Na-GTP at pH 7.3 (KOH-adjusted) and osmolarity 298 mOsm
(sucrose-adjusted).
Ion selectivity tests were carried out in chloride-free recording solution
containing (in mM) 125 sodium gluconate, 2 potassium gluconate, 3 CaSO
4
,
1 MgSO
4
, 10 HEPES, 30 glucose, at pH 7.3 (NaOH-adjusted) and 305–310
mOsm (sucrose-adjusted), and using intracellular solution containing (in mM)
125 potassium gluconate, 8 sodium gluconate, 0.1 CaSO
4
, 0.6 MgSO
4
, 1 EGTA,
10 HEPES, 4 Mg-ATP, 0.4 Na-GTP, pH 7.3 (KOH-adjusted), 295–300 mOsm
(sucrose-adjusted).
No action spectrum data were gathered from cells voltage-clamped at −65 mV
with a leak current greater than −200 pA or with a resting membrane potential
more positive than −45 mV in current-clamp mode. Access resistance was
5–30 MΩ. Resting membrane potential was ~−65 mV for neurons and ~−30 mV
for HEK 293FT cells in current-clamp recording. All parameters were monitored
throughout recording.
All in vitro action spectra, photocurrent and voltage data were taken with a
Leica DMI6000B microscope. Action spectra were taken with a monochromator
(Till Photonics Polychrome V, 15 nm bandwidth centered around each value).
Spectra for a given cell were taken by averaging red-to-UV (685 to 387 nm)
and UV-to-red (387 to 685 nm) spectra, to eliminate history dependence. Photon
fluxes for all wavelengths were ~3.0 × 10
21
photons/s/m
2
(0.756 mW/mm
2
at 670 nm). Normalized action spectra were obtained by dividing the averaged
photocurrent data by the highest observed data point.
Trafficking in vitro photocurrents were taken with a DG-4 optical switch with
300 W xenon lamp (Sutter Instruments), delivered with a 575 ± 25 nm bandpass
filter (Chroma). All other in vitro photocurrents were measured using 470 nm,
530 nm or 625 nm LEDs (Thorlabs). LED spectra were bandpass filtered with the
following (Semrock): 530 nm LED with 543 nm ± 11 nm filter or 625 nm LED
with 632 nm ± 11 nm. All light powers were measured out of the objective lens
with a PM200B photodetector (Thorlabs). Data were analyzed using Clampfit
(Molecular Devices) and Matlab (MathWorks, Inc.).
AAV preparation. The AAV particles used for cone photoreceptor targeting
were produced in the laboratory of B.R.
52
, with titers between 5.2 × 10
11
and
6.8 × 10
12
GC/ml. All other AAV constructs were produced by the University
of North Carolina Chapel Hill Vector Core at a titer of ~6 × 10
12
c.f.u./ml.
All viral dilutions were carried out in phosphate-buffered saline (PBS; Life
Technologies).
In vivo rodent electrophysiology. All light powers were measured and reported
at the fiber tip (rather than calculated some distance away); for example, we
measured 4 mW emitted from a 200-µm fiber, which we calculated and reported
as 127 mW/mm
2
density out of the fiber tip.
Injections for glass-pipette electrophysiology were made under isoflurane
anesthesia and buprenorphine analgesia, and 1 µl AAV was injected through a
craniotomy made in the mouse skull into the motor cortex (1.78 mm anterior,
1.5 mm lateral and 1.75 mm deep, relative to bregma) or the piriform, insular or
somatosensory/motor cortices (1.78 mm anterior, 2.0 mm lateral and, respec-
tively, 4.0, 3.0 and 2.0 mm deep, relative to bregma) of female C57BL6 mice
5–9 weeks old. Comparisons with eNpHR3.0 and Jaws were carried out by inject-
ing 2 × 10
9
viral particles in 1 µl of an AAV-PBS mixture into motor cortex.
All viruses were injected at a rate of 0.15 µl/min through a 34-gauge injec-
tion needle, after which the needle was allowed to rest at the injection site for
10 min to allow viral diffusion. The craniotomy was marked with Examix
NDS (GC America), headplates were affixed to the skull with skull screws
(JL Morris), and the craniotomy and headplate were covered with dental
cement (C&B Metabond).
Expression in Jaws-targeted populations of neurons in the prefrontal
cortex was achieved by intracranial injection of AAV8-hSyn-Jaws in P60–120
male C57BL/6 mice. Opsins were expressed in excitatory neurons in primary
visual cortex by injecting AAV5-CAG-FLEX viruses into Emx1-cre or Pvalb-cre
(ref. 53) mice. Mice were anesthetized with 1.5% isoflurane and virus was injected
at 0.1µl/min; mice were given 4–5 weeks for recovery.
Extracellular recordings to measure inhibition of spontaneous neural
activity were made in the cortex of headfixed awake mice 1–2 months after
virus injection, using 3–10 MΩ saline-filled glass microelectrodes containing
silver/silver chloride electrodes
1,3
. Signals were amplified with a Multiclamp 700B
and digitized with a Digidata 1440, using pClamp software (Molecular Devices).
A 635-nm, 200-mW laser (Shanghai Laser Optics and Century) was coupled to
a 200-µm-diameter optical fiber. An optical fiber was attached to the recording
glass electrode, with the tip of the fiber 600 µm laterally from and 500 ± 50 µm
above the tip of the electrode, and guided into the brain with a Sutter manipulator.
Transcranial recordings were carried out with a 635-nm red laser coupled to a
1,500-µm-diameter optical fiber, and the recording electrode was also attached to
a 200-µm-diameter fiber, coupled to a different 635-nm laser, as described above.
Comparisons between Jaws and eNpHR3.0 were carried out using a 593-nm
yellow laser (Shanghai Laser Optics and Century) and a 637-nm red laser
(Coherent Lasers), which were both coupled into the same 200-µm-diameter
optical fiber. All lasers were controlled via Digidata-generated TTL pulses. Light
powers were measured with an integrating sphere (S142C, Thorlabs) and PM200B
photodetector (Thorlabs).
In vivo whole cell patching was conducted in the cortex of anesthetized mice
with an autopatcher
54
. A 200-µm fiber was coupled to a 635-nm red laser, and

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe
doi:10.1038/nn.3752
attached to a patch electrode as described for extracellular glass pipette recordings.
Data were acquired and analyzed using pClamp software (Molecular Devices).
Extracellular recordings to measure suppression of evoked neural activity
were conducted in mice anesthetized with 0.2–0.4% isoflurane and 0.8 mg/kg
fentanyl. Extracellular recordings were performed with moveable arrays of
tetrodes (Thomas Recording) with a 250-µm interelectrode distance. Data
acquisition at 40 kHz and online spike identification by waveform analysis used
Cheetah data acquisition software (Neuralynx). An optical fiber (200 or 1,500 µm)
was placed on the dura next to the electrode array and connected to either a
593-nm or a 635-nm laser. To activate Jaws, 1-s pulses of light were given at 0.1 Hz
at varying light intensities (10, 25, 50 or 75 mW/mm
2
). Visually evoked responses
were driven by randomized sequences of drifting gratings at 16 orientations and a
spatial frequency of 0.05 cycles/degree on a background of mean luminance. The
size of the visual stimulus was optimized for the receptive fields of the recorded
population. Response measurements for each neuron were taken at the optimal
orientation as identified by post hoc analysis.
Data were analyzed using Matlab (MathWorks)
1,3
. Briefly, spikes were detected
and sorted offline using Offline Sorter (Plexon) or waveform template match-
ing and principal component analysis discrimination (KlustaKwik and custom-
written Igor software, J.A.C.
55
). Only clusters with clear refractory periods in the
autocorrelogram, indicating a single-unit source, were used for further analy-
sis. As a further precaution against over-representation of the same unit, cross-
correlograms were run for every pair of identified units in a set. Clusters that
were not easily separable were discarded. Light-modulated units were identified
by performing a paired t-test between the 1- or 5-s illumination period against
the baseline firing period of the same time duration immediately before illumi-
nation, thresholding at P < 0.05. The degree of suppression by Jaws activation
was calculated for each cell by dividing the mean firing rate during the
light stimulus by the mean baseline firing rate during the same time duration
before light stimulation onset. Rebound firing rates were calculated in a window
immediately after the cessation of the light stimulus. The width of the window
was determined as the duration for which firing rates were elevated >3 s.d. of
the mean spontaneous firing rate.
Light onset latency to inhibition was calculated by sweeping a 20-ms sliding
window to identify the earliest 20-ms period deviating from the baseline firing
rate, as assessed by performing a paired t-test for the firing rate during each
window versus during the baseline period, averaged across all trials for each
neuron. The time for after-light suppression to recover back to baseline was calcu-
lated by taking the median of all trials, owing to a nonparametric distribution.
Ex vivo slice electrophysiology. For whole-cell patch clamp recordings in motor
cortex, coronal 300-µm brain slices were prepared 4 or 6 weeks after viral injection
of AAV8-hSyn-Jaws. Male C57BL6 mice were deeply anesthetized with sodium
pentobarbital (200 mg/kg), then transcardially perfused with 15–20 ml of ice-
cold modified artificial cerebrospinal fluid (ACSF; composition in mM: NaCl 87,
KCl 2.5, NaH
2
PO
4
1.3, MgCl
2
7, NaHCO
3
25, sucrose 75, ascorbate 5, CaCl
2
0.5,
in ddH2O; osmolarity 320–330 mOsm, pH 7.30–7.40) saturated with carbogen
gas (95% oxygen, 5% carbon dioxide). The brain was rapidly removed from the
cranial cavity and then sectioned using a vibrating-blade microtome (VT1000S,
Leica). Slices were allowed to recover for at least 90 min in a holding chamber
containing ACSF (composition in mM: NaCl 126, KCl 2.5, NaH
2
PO
4
1.25, MgCl
2
1, NaHCO
3
26, glucose 10, CaCl
2
2.4, in ddH
2
O; osmolarity 299–301 mOsm; pH
7.35–7.45) saturated with carbogen gas at 32 °C before being transferred to the
recording chamber for electrophysiology. Once in the recording chamber, slices
were continuously perfused at a rate of 2 ml/min with fully oxygenated ACSF at
32 °C with added picrotoxin (100 µM), NBQX (20 µM; Sigma) and AP5 (50 µM;
Tocris) to block fast synaptic transmission in the slice.
For whole-cell patch clamp recordings of dentate granule cells, four male
C57BL6 mice 9–10 weeks old were injected with AAV8-Camk2a-Jaws under
deep isoflurane anesthesia. 0.5 µl AAV was delivered twice at two locations into
the hippocampus (3.2 mm posterior, 3.1 mm lateral and, respectively, 2.3 and
2.7 mm deep, relative to bregma) at 100 µl/min with a 34-gauge needle attached
to a Hamilton 5-µl syringe, and the needle was left in place for 3 min to allow
viral diffusion. Four weeks after injection, mice were killed by decapitation and
horizontal 300 µm brain sections were prepared on a Leica VT1200 S vibratome.
Sucrose-based ACSF (composition in mM: sucrose 75, NaCl 67, NaHCO
3
26,
glucose 25, KCl 2.5, NaH
2
PO
4
1.25, CaCl
2
0.5, MgCl
2
7; pH 7.4, osmolarity 305–
310 mOsm) was used for cutting (at 4 °C) and subsequent storage of slices (32 °C
for 20–30 min, then maintenance at room temperature, 23.0–23.5 °C). After >1 h
incubation, slices were transferred to a submerged recording chamber continu-
ously perfused at 2 ml/min with ACSF (in mM: NaCl 119, NaHCO
3
1.24, glucose
10, KCl 2.5, NaH
2
PO
4
1.24, CaCl
2
2.5, MgCl
2
1.3, pH 7.4, 295 mOsm) maintained
at room temperature. Jaws-infected cells were identified under 460–480 nm blue
light, whereas visual guidance of the patch electrode was assisted by infrared
differential interference contrast (DIC) microscopy (Olympus BX51).
Electrodes for cortical recordings were pulled from thin-walled borosilicate
glass capillary tubing using a P-97 puller (Sutter Instruments) and had resistances
of 4–6 MΩ when filled with internal solution (composition in mM: potassium
gluconate 125, NaCl 10, HEPES 20, Mg-ATP 3, Na-GTP 0.4 and 0.5% biocytin, in
ddH20; osmolarity 289 mOsm; pH 7.31). Capacitance, series resistance and input
resistance were frequently measured throughout recording to monitor cell health.
Cells were visualized through a 40× water-immersion objective on an upright
microscope (Scientifica) equipped with infrared DIC optics and a Q-imaging
Retiga Exi camera (Q Imaging).
Tip electrode resistance for dentate granule cell recordings was 4.6–7.4 MΩ in
ACSF. Patch electrode solution consisted of (in mM) potassium gluconate 122.5,
KCl 12.5, KOH-HEPES 10, KOH-EGTA 0.2, Mg-ATP 2, Na
3
-GTP 0.3, NaCl 8
(pH 7.35, mOsm 296), and 0.2–0.4 mg/ml biocytin was added immediately before
use. 10-pA step hyperpolarization/depolarization square current pulses were used
to determine the input-output relationship and 300 pA ramp depolarization was
used for AP generation. Uninfected dentate granule cells recorded from the con-
tralateral dentate gyrus of virus-injected mice or uninjected animals served as
controls. None of these cells responded to 625-nm red light. Uncompensated
series resistance was typically 14–25 MΩ, and cells with resting membrane poten-
tial more positive than −50mV were discarded (n = 3).
Optical activation was delivered to motor cortex using 500-ms pulses from
590-nm or 625-nm LEDs (Thorlabs), which were additionally filtered with band-
pass filters (Semrock): 590-nm LED with 590-nm ± 10-nm filter or 625 nm LED
with 632-nm ± 11-nm filter, and to dentate granule cells using 1 s 625-nm red
light (M625L3-C1, ThorLabs, 68 mW/mm
2
). All light powers were measured out
of the objective lens with a PM200B photometer (Thorlabs). Illumination spot
sizes were measured by photobleaching an in-focus microscope slide coated with
Alexa 488 dye for 10 min under full-intensity illumination, then imaging with a
micrometer calibration slide to determine the photobleached radius.
All recordings were made using a Multiclamp 700B amplifier and Clampex 10
or 10.4 software (Molecular Devices, CA, USA). Cortical recordings were low-
pass filtered at 1 Hz and digitized at 10 kHz using a Digidata 1550 (Molecular
Devices, CA, USA); dentate granule cell recordings were digitized with a Digidata
1440. All data were analyzed using Clampfit (Molecular Devices) and Matlab
(MathWorks).
Retinal multi-electrode array recordings. Rodent information is outlined in
Supplementary Table 1. Experiments were conducted as previously described
24
.
Briefly, for viral injections to the retina, animals were anesthetized using 3%
isoflurane. A small incision was made with a sharp 30-gauge needle in the sclera
near the lens and 2 µl AAV particles were injected slowly into the subretinal space
using a blunt 5-µl Hamilton syringe held in a micromanipulator.
To record spike trains from retinal ganglion cells, isolated mouse retinas were
placed on a flat MEA60 200 Pt GND array (30-µm-diameter microelectrodes spaced
200 µm apart) (Ayanda Biosystems or Multi Channel Systems). The retina was
continuously superfused in oxygenated Ringer’s solution (110 mM NaCl, 2.5 mM
KCl, 1.0 mM CaCl
2
, 1.6 mM MgCl
2
, 22 mM NaHCO
3
and 10 mM -glucose
(pH 7.4 with 95% O2/5% CO
2
)) at 36 °C during experiments. Signals were
recorded (MEA1060-2-BC, Multi-Channel Systems) and filtered between
500 Hz (low cut-off) and 3,500 Hz (high cut-off). Action potentials were
extracted with a threshold of greater than 4 times the s.d. of the recorded trace
(Matlab, MathWorks).
Histology. We performed histology on n = 6 mice injected with AAV8-hSyn-Jaws,
n = 2 mice injected with AAV8-hSyn-eNpHR3.0 and n = 2 mice injected with
AAV8-hSyn-Jaws-ER2, representative images from each of which are shown in
Figures 4 and 5. We examined n = 7 opsin-expressing retinas, and the images

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe
doi:10.1038/nn.3752
shown in Figure 2 and Supplementary Figure 5 are representative of the whole
population.
Mice were terminally anesthetized with isoflurane, then perfused through the
left cardiac ventricle with 4% paraformaldehyde in PBS. The brain was removed
and sectioned into 40 µm coronal sections on a cryostat and subsequently
mounted with Vectashield HardSet (Vector Labs). Acute brain slices were fixed
in 4% paraformaldehyde overnight at 4 °C after recording and then washed in
PBS. The slices were blocked for 1 h at room temperature in PBS containing 3%
normal donkey serum (NDS) and 0.3% Triton, followed by incubation with Alexa
405-conjugated streptavidin (1:1,000; Life Technologies) in PBS containing 3%
NDS and 0.3% Triton, for 2 h at room temperature, to reveal biocytin labeling.
Brain slices were subsequently washed in PBS, mounted on glass slides and
coverslipped with PVA-DABCO (Sigma).
Cryosectioned slides were visualized and imaged with a Zeiss LSM 510 confo-
cal microscope using 20× and 63× objective lenses, or a Nikon TI-E microscope
using a 20× objective lens. Acute slices were imaged using a Olympus FV1000
confocal laser scanning microscope through a 10×, 0.40 numerical aperture objec-
tive or a 40×, 1.30 numerical aperture oil-immersion objective using Fluoview
software (Olympus). Images were subsequently processed in Adobe Photoshop
CS6 (Adobe Systems).
Retinal immunostaining was conducted as previously described
24
. Briefly,
retinas were isolated and fixed in 4% paraformaldehyde in PBS for 30 min at
room temperature, and washed in PBS overnight at 4 °C. Retinal whole mounts
were incubated in 30% sucrose treated with three freeze-thaw cycles, after
which all steps were performed at room temperature. Retinas were incubated in
a blocking solution (10% normal donkey serum (NDS, Chemicon), 1% bovine
serum albumin (BSA) and 0.5% Triton X-100 in PBS, pH 7.4) for 1 h. Primary and
secondary antibody applications were done in 3% NDS, 1% BSA, 0.02% sodium
azide and 0.5% Triton X-100 in PBS.
Primary antibodies for GFP (rat-GFP, 1:500, Nacalai/Brunschwig 04404-84)
and mCAR (rabbit-cone-arrestin, 1:200, Millipore AB15282) were applied for
3 d. After washing the retina three times for 10 min in PBS, the retina was incub-
ated with cyanine dye-conjugated secondary antibodies (1:200; donkey anti-
rat–Cy5, Jackson Labs 712-175-153, or donkey anti-rabbit–Cy3, Jackson Labs
711-165-152) and 10 µg/ml DAPI (Roche Diagnostics) for 2 h. After another
three 10-min washes in PBS, the retina was mounted on a glass slide with ProLong
Gold antifade reagent (Invitrogen). Confocal images of antibody-stained retinas
were taken using a Axio Imager Z2 equipped with a LSM 700 scanning head.
In vivo light propagation measurements and Monte Carlo modeling. Isotropic
light measurement probes were constructed by gluing a 300-µm-diameter spheri-
cal ruby ball lens to the end of a 400-µm-diameter multimode optical fiber with
transparent, UV-cured adhesive. Emitted fluorescence was measured using an
HR2000 CCD spectrometer (Ocean Optics) and recorded using SpectraSuite
(Ocean Optics). Prior to testing, a dark spectrum measurement was taken and
a nonlinearity correction was applied using default coefficients. The probe was
calibrated in water (n = 1.33) with collimated light.
532- or 635-nm lasers (Shanghai Laser Optics Company) were connected to
an optical shutter via a 200-µm-diameter multimode fiber, and the shutter was
FC-coupled to a 1.5-mm-diameter plastic optical fiber. Five male C57BL/6J mice
were used for green light measurements and five different C57BL/6J mice were
used for red light measurements. All measurements were taken under pentobar-
bital analgesia (50 mg/kg, i.p.). Of the 10 mice × 3 depths × 5 powers, one mouse
was not analyzed because the 3-mm-depth files were inadvertently not saved
owing to manual save process clunkiness; 6 other files were also inadvertently
not saved, but at least 3 power levels were measured for every depth and mouse
other than the one noted above.
In Matlab, we performed Monte Carlo simulations of light scattering and
absorption in the brain from light emitted from the end of an optical fiber
by dividing a cube of gray matter into a 100 × 100 × 100 grid of voxels, each voxel
50 µm × 50 µm × 50 µm in dimension, using previously published algorithms
56,57
.
To achieve accurate simulations of light propagation close to the optical fiber,
before the orientation of photon trajectories is randomized by multiple scattering
events, we used an anisotropic scattering model with either Henyey-Greenstein
or Gegenbauer kernel phase functions, as indicated below. We interpolated data
from ref. 58 to obtain scattering coefficients for gray matter and white matter at
wavelengths of 532 and 635 nm (below). Because ref. 58 used samples washed
of blood, the effects of blood on optical properties of these tissues would not
be reflected in their measurements; to accurately simulate in vivo conditions,
the effects of blood scattering must be added. We took the brain’s blood volume
fraction to be 4% (ref. 59) and the red cell volume fraction of blood (hematocrit)
to be 45% (ref. 60). Thus, excluding large blood vessels, the brain contains dilute
blood that has about a 1.8% hematocrit. In this red blood cell concentration
regime, the anisotropy coefficient is constant and the scattering and absorbing
coefficients are directly proportional to hematocrit
61
. To calculate the total scat-
tering coefficient, we added the scattering coefficient due to brain tissue alone
to the scattering coefficient due to the blood in the tissue. To calculate the total
absorption coefficient, we likewise added the absorption coefficient due to brain
tissue alone to the absorption coefficient due to the blood in the tissue. To cal-
culate the anisotropy coefficient, we took the weighted average anisotropy coef-
ficients of the tissue and blood, weighted by the relative likelihood of each of the
two components of causing a scattering event. We chose the Gegenbauer kernel
for the phase function when blood was present, as blood is a large component
of the scattering and has been shown to be modeled well using that phase func-
tion
61
. With no blood present, as was assumed for the bone, we used the Henyey-
Greenstein phase function as in ref. 58. Optical properties of bone at 532 nm and
633 nm were interpolated from data in refs. 62,63.
We used simplified geometries of gray matter and skull bone to illustrate the
effects of tissue variations in several experimental locations. In each case, the
tissue properties were set on a per-voxel basis, with no corrections for specular
reflections or refraction at boundaries between tissue types.
We launched 2 × 10
6
packets of photons in a fiberlike radiation pattern through
fibers modeled on the experimentally used fiber—1.5 mm diameter, 0.5 NA—and
modeled their propagation into the brain on the basis of the algorithm of ref. 64.
In essence, whenever a photon packet entered a voxel, our program probabil-
istically calculated the forecasted traveling distance before the next scattering
event. If that traveling distance took the photon packet out of the starting voxel,
then the packet would be partially absorbed appropriately for the distance it
traveled within the voxel and the voxel’s absorption coefficient. The process would
then restart upon entry of the photon packet into the new voxel. If the distance
traveled before scattering was less than the distance to the edge of the voxel, then
the packet would be partially absorbed appropriately for the distance it traveled
within the starting voxel, and a new direction of packet propagation would be
randomly chosen according to the phase function. Using this model, we gener-
ated Supplementary Figure 1a,b, which shows the contours at which the light
fluence falls off to various percentages of the light intensity emitted by the fiber.
The coefficients we used were as follows:
Gray matter at 532 nm with 4% v/v blood: absorption coefficient (mm
−1
) 0.942,
scattering coefficient (mm
−1
) 23.3, anisotropy factor 0.949, using the Gegenbauer
kernel phase function.
Gray matter at 633 nm with 4% blood: absorption coefficient (mm
−1
) 0.071,
scattering coefficient (mm
−1
) 20, anisotropy factor 0.95, using the Gegenbauer
kernel phase function.
Bone at 532 nm with no blood: absorption coefficient (mm
−1
) 0.105, scatter-
ing coefficient (mm
−1
) 34, anisotropy factor 0.93, using the Henyey-Greenstein
phase function.
Bone at 633 nm with no blood: absorption coefficient (mm
−1
) 0.06, scatter-
ing coefficient (mm
−1
) 33, anisotropy factor 0.93, using the Henyey-Greenstein
phase function.
White matter at 532 nm with 4% blood: absorption coefficient (mm
−1
) 1.0,
scattering coefficient (mm
−1
) 55.3, anisotropy factor 0.862, using the Gegenbauer
kernel phase function.
White matter at 633 nm with 4% blood: absorption coefficient (mm
−1
) 0.134,
scattering coefficient (mm
−1
) 53.8, anisotropy factor 0.87, using the Gegenbauer
kernel phase function.
To measure skull thickness for Supplementary Figure 1a, male C57BL/6 mice
were transcardially perfused with 4% formaldehyde in PBS, and the skulls were
removed and imaged with a Nikon XTH160 X-ray micro-CT system. Images were
generated at an X-ray voltage of 45 kV and a current of 7.5 W. Sixteen frames per
projection were acquired with an integration time of 100 ms, with a total acquisi-
tion time of 42 min per skull. Acquired images were reconstructed with CT Pro
3D (Nikon Metrology) and visualized with VGStudio 2.0 (Volume Graphics).

© 2014 Nature America, Inc. All rights reserved.
nature neurOSCIenCe
doi:10.1038/nn.3752
Coordinates were stereotactically defined relative to bregma and skull thicknesses
determined by measuring z-direction differences at a given coordinate.
Statistical analysis. No statistical methods were used to predetermine sample
sizes, but sample sizes in this paper were similar to those in previous papers
from our group
1,3,65,66
and were chosen to reflect sample sizes that might be
experienced by end-users. No blinding was carried out, with the exception of
cotransfected opsin-GFP and tdTomato data sets, as the chief goal of the paper
was to demonstrate the capabilities of a new technology. No randomization was
carried out. All replicates were biological.
Statistical analyses were performed using JMP Pro 10 (SAS Software) and
Prism 5 (GraphPad). All data were tested for normality with a Shapiro-Wilk
test. Two-sample comparisons were characterized with a two-tailed t-test, and
multiway comparisons for a single variable were characterized with an ANOVA
followed by a Newman-Keuls multiple comparison post-test between pairs.
Paired tests were conducted with a two-tailed paired sample t-test. A
Kolmogorov-Smirnov test was used to assess differences in spiking frequency
distributions because of its sensitivity to both location and shape of cumulative
distribution functions.
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