Download Chapter 15 Viral Vector-Based Techniques for Optogenetic

Chapter 15
Viral Vector-Based Techniques for Optogenetic
Modulation In Vivo
Mathias Mahn, Shiri Ron, and Ofer Yizhar
Abstract
Optogenetics is a technical methodology that allows direct light-based manipulation of genetically specified
cells. Optogenetic methods have provided novel insights into the role of defined neuronal populations in
brain function and animal behavior. An expanding palette of single-component optogenetic tools provides
powerful interventional strategies for modulating the function of targeted neurons in awake, behaving
mammals and for detailed interrogation of circuit physiology in vitro. Although several genetic methods
can be utilized for delivering these genes into target cell populations, the use of viral vectors for delivery of
optogenetic tools has several important advantages. In recent years, techniques for viral vector-mediated
delivery of optogenetic tools have improved and expanded significantly. These techniques now allow modular use of optogenetic tools in defined cell types and circuits and dovetail well with genetic mouse models
and recombinase-based driver lines. Here, we review the use of viral vectors for delivering genes encoding
optogenetic tools into the rodent brain and provide a detailed protocol for viral transduction of mouse
cortical neurons and chronic implantation of a fiberoptic connector for light delivery in vivo.
Key words Optogenetics, Lentivirus, Adeno-associated virus, Circuit tracing, Fiberoptic cannula
1
Introduction
Microbial rhodopsins are light-sensitive, retinal-containing proteins known for many years to be crucial for the survival and function of a wide range of microbial species [1]. Recently, with the
discovery of channelrhodopsin [2, 3] and the first application of
this microbial opsin for stimulating neurons [4, 5], neuroscientists
began to realize the immense potential of these light-activated proteins to serve as genetically encoded tools for manipulating the
activity of defined neural circuit elements with high spatiotemporal
resolution [6]. Optogenetics relies on the natural capacity of
microbial rhodopsins and other engineered light-responsive
Mahn and Ron contributed equally to this work.
Riccardo Brambilla (ed.), Viral Vector Approaches in Neurobiology and Brain Diseases, Neuromethods, vol. 82,
DOI 10.1007/978-1-62703-610-8_15, © Springer Science+Business Media, LLC 2014
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proteins to encode both light sensation and effector function [7].
A major advantage of microbial opsin-based optogenetic tools is
that they are essentially cofactor free. The endogenous presence of
the organic cofactor all-trans-retinal (ATR) in vertebrate tissues
enables the use of these genes as single-component tools, allowing
for spatially and temporally precise functional modulation of the
intact mammalian nervous system. Neurons transduced with
microbial opsin genes become sensitized to light with the physiological effect of illumination dictated by the type of optogenetic
tool being used. For example, expression of channelrhodopsin-2
(ChR2), a light-gated cation channel [3, 4], renders cells excitable
by light. Light-gated ion pumps such as halorhodopsin (e.g.,
NpHR) or archaerhodopsin (e.g., Arch) can be used to hyperpolarize and inhibit the electrical activity of neurons [8, 9]. In recent
years, the optogenetic toolbox has greatly expanded and now
includes multiple tools for excitation, inhibition, and modulation
of signal-transduction pathways in neuronal and non-neuronal
cells [6]. We and others have reviewed the criteria for selection of
optogenetic tools for particular experiment types [7, 10–12]. The
choice of opsin for a given experiment will depend on a multitude
of factors, including the desired physiological effect, the type of
neuron targeted, the spatiotemporal extent of modulation, and the
light-delivery system utilized [13]. The use of viral vectors, therefore, provides cost effective and flexible means of optimizing
experiment-specific tools for each application.
When applied in the adult brain, optogenetics requires genetic
modification of post-mitotic neurons to induce expression of the
light-gated channels or pumps. Genetically engineered viruses are
by far the most popular means of delivering optogenetic tools.
Lentiviral vectors (LV) [14] and adeno-associated viral vectors
(AAV) [15] have been widely used to introduce opsin genes into
mouse, rat, and primate neural tissues [11]. These vectors allow
high expression levels over long periods of time with little or no
reported adverse effects [15]. AAV-based expression vectors possess lower immunogenicity and offer the advantage of larger transduced tissue volumes compared with LV due to their high viral
titers and diffusion properties. Additionally, AAV is considered
safer than LV as the currently available strains do not broadly integrate into the host genome and are thus rated as biosafety level
(BSL) 2 agents. Both LV and AAV vectors can be used in conjunction with cell type-specific promoters (e.g., [14, 16, 17], see [10]
for more examples), and both vector families support pseudotyping techniques, which in principle enable a wide range of cell-type
tropisms and transduction mechanisms [18, 19]. Finally, Credependent vectors, in which Cre recombination can activate the
expression of transgenes to achieve cell type-specific control [20,
21], are typically made with AAV-based vectors.
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Typical AAV vectors can be produced at much higher titers
than LV (typical yields of AAV reach 1013 genome-containing
capsids per ml, whereas the same amount of starting material will
yield ~109 VSVG-pseudotyped lentivirus). In spite of its huge
potential in brain research, several properties of AAV make its production as a recombinant viral vector a technical challenge for most
neuroscience laboratories. One such property is the inherent instability of the inverse tandem repeat sequences (ITR’s) required for
correct demarcation of the genomic sequence inserted into the
capsid, as well as for effective reverse strand synthesis which is a rate
limiting step for efficient gene expression from AAV. In addition,
in the absence of the full complement of AAV genes and a helper
virus (such as adenovirus) in the laboratory setting, recombinant
AAV is not exuded from the cells, thus requiring the lysis of the
producer cell culture during the purification process. This requires
several technically challenging downstream purification steps such
as purification columns or ultracentrifugation [22]. Although both
LV and AAV have been successfully used in optogenetic experiments, most neuroscience laboratories that do not possess molecular biology and tissue culture capabilities choose to purchase
AAV-based vectors from core facilities offering virus production
services where commonly used viruses are often kept in-stock as
injection-ready aliquots.
In optogenetic experiments, several critical factors should be
taken into account in order to assure that the desired physiological
effect is evoked by light stimulation (1) expression of the optogenetic actuator should be robust and allow modulation with moderate light power in order to avoid phototoxicity; (2) expression
should be restricted to the desired neuronal population, with minimal “genetic leak” to nontargeted cell populations; (3) the method
used to express the selected tool should be well tolerated and nontoxic to cells over the entire duration of the experimental period
(and ideally well beyond this time); (4) in behavioral experiments,
the physiological effect of the manipulation performed should
always be validated in one of several electrophysiological recording
methods to assure that the desired effect is achieved and to correctly interpret behavioral results. For example, acute slice patchclamp recordings can be performed to verify expression and the
efficacy of light stimulation; extracellular recordings can further
provide important validation of the effect of the optogenetic
manipulation in the intact circuit.
1.1 Target Volume
Considerations
Depending on the scientific question, the required spatial distribution of transduced cells can vary dramatically. This necessitates
adjustment of the viral delivery procedure and viral vector type to
allow for efficient transduction in the target tissue. Furthermore,
the choice of optogenetic tool and illumination method is critical
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to assure that the desired effect is achieved in the targeted cells.
The extent of viral transduction depends both on the type of vector
used and the brain region targeted. Generally, a relatively restricted
expression pattern can be achieved by choosing the appropriate
viral vector and injection volume. For example, AAV2 injection
results in expression patterns that are more localized compared
with the pseudotyped AAV2/5, AAV2/8, or AAV2/9. AAV2 is
therefore well suited for local expression in volumes smaller than
1 mm3 [18]. Although viral titer can be reduced in order to
decrease the size of the transduced volume, lower titer injections
are also likely to influence the number of genome copies in transduced cells, leading to lower expression levels of the transgene in
individual cells within the region [18]. On par with AAV2 transduction, LV transduction is more spatially restricted in vivo and
can thus be used to target smaller structures. However, LV has
been reported to exhibit a bias towards excitatory neurons in cortex [23], an effect which is likely also region specific, since other
more specialized cell types have been successfully targeted with
lentiviral vectors [24, 25].
If the target area is large, there are several possible limitations
to the spatial extent of tissue that can be affected through optogenetic techniques. In the case of optogenetic inhibition, light-gated
ion pumps such as NpHR [8] and Arch [9] are commonly used.
These opsins have been optimized for neuroscience applications by
adding various targeting motifs and are efficiently expressed in neurons [13, 26]. Due to the need for continuous illumination for
activation of these ion pumps, the limiting factor in experiments
utilizing these tools is the light power density required to achieve
effective hyperpolarization of the expressing neurons. Since brain
tissue strongly absorbs visible light and is a highly scattering
medium, light power density drops to ~1 % within 1 mm from the
light source (although this effect is strongly wavelength dependent;
see references [9, 10, 24, 27] for more details). The maximal applicable light power for optogenetic modulation is further limited by
its cytotoxic effect above certain irradiance levels. A power density
of 100 mW/mm2 is regarded as the upper bound for direct light
stimulation in brain tissue, thereby restricting the volume of tissue
that can be modulated with optogenetic tools requiring intense
light for effective modulation (for example, the effective power
dose required for eNpHR3.0 is 5 mW/mm2, while most channelrhodopsin variants require less than 1 mW/mm2 [13]). A single
injection of 1 μl of AAV2/5 or AAV2/1 can lead to expression in a
region >1 mm3 (see Fig. 3 for a representative example). If the
opsin expressed is eArch3.0 or eNpHR3.0, the volume affected by
light stimulation through an implanted optic fiber in this region is
likely to be smaller than the transduced volume [28]. Therefore, in
this scenario the transduction volume is normally not restrictive.
In the case of neuronal excitation, new tools such as the stepfunction opsins (SFO [29, 30]) can be used to effectively
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depolarize neurons at very low light power densities (up to 3 orders
of magnitude lower power density values than required for
stimulation with wild-type ChR2). The slow closing kinetics of
these channels allows for the accumulation of open channels over
long periods of time. This allows for neurons in large volumes of
brain tissue to be depolarized with relatively low light power densities, especially if millisecond precise control of neuronal activity is
not needed, or even undesired [30]. Here, viral transduction volume can become a limiting factor. The transduction efficiency, titer
and binding affinity to the cell surface of the injected viral vector as
well as its spread radius can be tailored to the desired outcome, as
detailed below.
Recent advances in AAV pseudotyping techniques have resulted
in strongly enhanced transduction efficiency [31]. AAV2/9 has
been shown to possess far superior transduction efficiency compared to wild-type AAV2 ([32, 33]; for review see [34]) and hence
results in larger transduction volumes [35]. The distance of viral
spread is dependent on multiple factors, including particle size and
abundance of the cognate receptor for cell entry. One method to
overcome the small range of diffusion-based injection protocols is
rapid injection, resulting in convection as shown in 2006 by
Raghavan and colleges [36]. This method, referred to as
convection-enhanced delivery (CED) seems promising, but
requires special injection needles that can prevent viral backflow
along the needle insertion tract. Another option is the coadministration of agents aimed at increasing the diffusion of viral
particles. Intravenous mannitol injection [37] can be used to
decrease intracranial pressure, resulting in increased diffusion distance. A 2 μl injection of AAV2/9, delivered using CED combined
with systemic mannitol administration allows for the effective
transduction of complete brain regions such as the hippocampus,
including contralateral structures [38]. In the case of AAV2, which
binds to heparan sulfate proteoglycans on the cell surface, coinfusion of heparin was reported to increase viral spread [39].
Although these advanced methods have not been explored directly
for optogenetic experiments, we expect that they will be useful
when the targeting of larger volumes is desired.
1.2 Circuit-Based
Expression of
Optogenetic Tools
Functional dissection of intact neural circuits is one of the most
widely
used
applications
of
optogenetic
techniques.
Channelrhodopsin can itself serve as a tool for anterograde circuit
mapping [40, 41]. Introduction of fluorescently tagged channelrhodopsins into a specific neuronal population in a defined brain
region enables the visualization and subsequent photoactivation of
projecting axon terminals throughout the brain. Simultaneous
electrophysiological recording at the projection site allows the
identification of specific postsynaptic components of the circuit
both in vivo and in the acute brain slice preparation [40, 42–46].
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At the whole-brain level, optogenetic stimulation has also been
integrated with functional magnetic resonance imaging (ofMRI
[47–49]) for identification of global circuits recruited by defined
local activity patterns. This method enables unbiased mapping of
functionally connected areas without a priori knowledge of circuit
connectivity and allows identification of downstream loci.
Recently, several new viral techniques have been developed that
allow the expression of optogenetic tools in neuronal populations
that are defined by their pattern of synaptic connectivity. Integrating
these tracing methods with optogenetics allows combined anatomical and functional dissection of defined circuits. Several recent
studies have described and optimized an elegant technique for tracing monosynaptic retrograde connections with a modified rabies
virus; this approach is based on a glycoprotein-deleted variant of
the rabies SAD B19 strain, SADΔG [50, 51]. The rabies virus glycoprotein (G), which is embedded in the viral membrane, is
required for trans-synaptic spread. By introducing the glycoprotein
gene in neurons prior to infection of the G-deleted mutant virus,
the virus spreads to presynaptic neurons and is restricted from further spread due to the lack of this complementary glycoprotein in
newly infected neurons [52]. This enables the dissection of direct
connections originating from a population of defined neurons or
even from a single primary infected neuron [53].
Osakada et al. [54] incorporated ChR2 into the glycoproteindeleted rabies virus (yielding SADΔG-ChR2-mCherry) and successfully displayed optical activation of presynaptic ChR2 expressing
cells. This system makes it possible to outline the function of specific connections through combination with electrophysiological
recordings or behavioral paradigms. For an additional layer of
specificity, the primary viral transduction can be directed to genetically defined post-synaptic neuronal subtypes by using the avian
receptor TVA system [55]. To achieve specificity, the SADΔG virus
is pseudotyped with an envelope protein from the avian sarcoma
and leukosis virus (ASLV). The TVA receptor, which is required
for infection by the pseudotyped rabies virus and is only found in
birds, is then expressed in the cells to be targeted for infection by
SADΔG along with the rabies glycoprotein. This allows the virus
to spread trans-synaptically exclusively from the TVA-expressing
cells to their presynaptic partners. Specificity in retrograde transport can also be achieved by targeting the rabies G protein gene to
specific neurons using the double-floxed Cre-based expression system [56]. In this preparation, only Cre-expressing cells will harbor
the necessary machinery for retrograde transport of the recombinant rabies virus. To broaden the microbial opsin repertoire available for monosynaptic retrograde tracing, one could introduce a
Cre recombinase-expressing SADΔG virus into transgenic mice
engineered for Cre-dependent ChR2/Arch/NpHR expression
[57]. The advantages of using rabies-based circuit tracing
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techniques are the unidirectional spread, fast expression, and high
amplification capabilities of the SAD B19 virus [58]. On the other
hand, the time course of survival of SAD B19-infected neurons is
limited with cell health diminishing about 2 weeks following infection [51], suggesting that for experiments requiring long-term
survival of transduced neurons, other systems might be required.
An additional method for trans-synaptic circuit tracing is the
use of Cre-tagged trans-cellular tracer proteins. Trans-cellular
tracer proteins can be transported across synapses after binding to
specific glycoconjugates on the membrane (reviewed in [59]).
When using these proteins to confer trans-synaptic transport function, it is possible to fuse proteins of interest to these tracers,
thereby allowing circuit tracing (with fluorescent markers) as well
as delivery of a functional protein if desired (such as enzymes).
Taking advantage of the trans-cellular transport properties of wheat
germ agglutinin (WGA), ChR2 can be expressed in a subset of
synaptically connected neurons [26]. This system requires two
separate constructs and viral injection sites: the first, a WGA–Cre
fusion protein for combined recombinase and trans-synaptic functionality, which can be expressed using AAV and targeted stereotactically to a defined brain region. Cre-dependent AAV vector,
conditionally expressing a microbial opsin gene, is then targeted to
a predicted trans-synaptic target region. Although this method has
been successfully demonstrated in cortico-cortical and hippocampal circuits, the directionality of transport of WGA–Cre has not yet
been fully characterized and might well be circuit dependent. In
this type of experiment, prior knowledge of synaptic connections
between the targeted neuronal populations is necessary in order to
determine the secondary injection site. For a more unbiased means
of functional mapping, it is possible to introduce WGA–Cre into
transgenic mice that conditionally express fluorescent reporter
proteins [60] or microbial opsins [57].
Although several retrograde trans-synaptic tracing approaches
have been described, anterograde-specific tracer viruses are not yet
sufficiently optimized for optogenetic experiments. Currently, the
most promising viral tool for anterograde tracing is the H129
strain of herpes simplex virus (HSV) type 1 [61] which has recently
been developed for genetically defined, Cre-dependent, anterograde tracing [62]. A possible application of this technology would
be the insertion of Cre-dependent microbial opsins into the H129
genome enabling light-induced activation of downstream circuit
elements. A different strain of HSV1, which is taken up by axons
and transported retrogradely, has been used for expression of
ChR2 in neurons projecting to the site of injection. This allows for
photostimulation-assisted identification of neuronal populations
(PINP [63]). Importantly, not all HSV viruses spread anterogradely, and the directionality depends on the specific strain used.
As with many other neurotropic viral vectors, the main
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disadvantage of this viral system is its adverse effect on cell and
animal health over a course of days and weeks.
1.3
2
2.1
Summary
Viral vector-based expression is a versatile and widely used method
for delivering optogenetic tools in the brain. However, animal-toanimal variation in the efficiency of viral transduction is also a
potential major source of variability in optogenetic experiments.
Although electrophysiology and immunohistochemistry can allow
researchers to correlate the behavioral or physiological effect with
the efficiency of genetic expression of the optogenetic tool, optimization of the method for viral transduction and light delivery to
the targeted cells can reduce this inherent variability and allow
more robust, reproducible results. In the protocol below, we focus
on the procedures developed for direct intracerebral delivery of
high-titer virus for expression of optogenetic tools in defined cell
populations. We also describe the procedure for chronic implantation of fiberoptic connectors for robust, reproducible optical modulation of virally transduced neurons in vivo.
Protocol
Reagents
Iodine antiseptic (e.g., Betadine)
Ethanol, 70 %
Sterile saline
Sterile distilled, deionized water (DDW)
Anesthetics and analgesics
Lubricant eye ointment
Optional: hydrogen peroxide (H2O2)
2.2
Equipment
2.2.1 Virus Injection
Surgical tools (e.g., curved sharp tip tweezers; fine, spring scissors;
surgical scalpel; dull-tip forceps; wide tipped bulldog clamps)
Hot bead tool sterilizer (see Note 1)
Dissection stereomicroscope (Fig. 1-a1)
Small animal stereotaxic apparatus with cannula holder, nonrupture tip ear-bars, tooth bar, rodent face mask for isoflurane
circulation (Fig. 1-a2, a3)
Temperature-controlled heating pad (Fig. 1-a4, a5)
Isoflurane vaporizer (LEI Medical; Fig. 1-a6)
Isoflurane induction box (Fig. 1-a7)
Small heat lamp
Small hair trimmer
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Fig. 1 Surgery setup and procedure. (a) Overview of the surgery setup including (1) stereomicroscope, (2) stereotaxic instrument, (3) display of stereotaxic coordinates, (4) mouse stretcher with temperature controlled
heating pad, (5) temperature controller, (6) isoflurane vaporizer, (7) isoflurane induction box. (b) Major steps of
viral vector injection procedure: (I) head fixation with tooth bar, face mask, and ear bars, (II) Scalp incision and
skull exposure using wide-tipped bulldog clamps, (III) axis zeroing with needle tip at bregma, (IV) head orientation by horizontal and vertical lambda alignment, (V) marking of the intended hole position, (VI) needle insertion
into the brain and injection. (c) Major steps of optical fiber implantation: (I) shortening of the optical fiber to
required length using the ear bar scale, (II) axis zeroing with optic fiber tip at bregma, (III) placement of the optic
fiber above the injection site, (IV) adhesion enhancement by application of Metabond, (V) fixation of the optic
fiber to the skull using dental acrylic, (VI) closing of the incision around the dental acrylic using tissue adhesive
Micro motor drill with micro burrs (0.5–0.7 mm)
1 ml syringe with needles (30G) for intraperitoneal injections
Microsyringe pump (e.g., World Precision Instruments UMP-4)
Cotton swabs, optional: absorbent swabs (Sugi)
50 ml conical tubes
Vetbond tissue adhesive or surgical suture
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2.2.2 Fiber Implants
Implantable fiberoptic connector with metal or ceramic ferrule—
fiber width: 200 μm diameter; fiber length: according to injection site DV coordinate (Fig. 1-bI)
Cannula holder for stereotactic frame
C&B Metabond kit
Acrylic dental cement
2.2.3 Behavioral Testing
Open field chamber for mice: square arena 50 cm × 50 cm made of
gray plastic with 40 cm high walls.
Mouse behavioral tracking system (Viewer3, BIOBSERVE GmbH,
St. Augustin, Germany)
Master-8 pattern generator (A.M.P.I, Jerusalem, Israel)
447 nm, 100 mW diode laser (O.E.M Laser Systems Inc., Salt
Lake City, UT, USA)
Fiberoptic patch-cord (200 μm core, Thorlabs Inc., cat # M72L05)
Fiberoptic patch-cord (200 μm core, FC/PC to bare ferrule connector, Doric Lenses, Quebec, Canada)
Fiberoptic rotary joint (Doric Lenses; Cat. # FRJ 1X1 FC/FC)
Zirconia sleeve for ferrule–ferrule connection (Thorlabs Inc., cat #
ADAF1)
2.3
Setup
1. Viral vectors
●
●
Storage: viruses should be stored in aliquots of 5–10 μl at
−80 °C.
Thawing: on day of surgery, virus aliquots should be
thawed quickly in a gloved hand or a warm water bath and
then kept at 4 °C until use. AAV can be stored after thawing for several days at 4 °C with minimal loss of activity.
LV is less stable and should be used on the same day.
2. Preparation of surgical tools
●
Set heat bead sterilizer to 250 °C.
●
Sterilize surgical tools by placing them, for at least 45 s, in
sterilizer so that the tips are fully covered by the beads.
●
Arrange the sterilized surgical tools on a sterile surgical
pad next to the stereotaxic instrument and allow them to
cool before beginning the procedure.
3. Preparation of the viral injection syringe
●
The syringe should be washed thoroughly with DDW and
then sterile saline prior to virus uptake. This is a good
opportunity to make sure that the needle is not clogged by
visually confirming that liquid is withdrawn and expelled
by slowly moving the syringe plunger up and down.
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4. Coordinates
●
Coordinates of the different brain regions can be determined using brain atlases [64] (see Note 2). Coordinates
of the intended injection area are given in three axes: X:
medial–lateral (ML), Y: anterior–posterior (AP), and Z:
dorsal–ventral (DV).
5. Anesthetics and analgesics: in this protocol, we describe the
use of isoflurane for anesthesia, but other methods are also
possible (see Note 3).
2.4 Surgical
Procedure
●
Induction: We suggest using a ketamine (80 mg/kg) xylazine (10 mg/kg) mix for anesthesia induction (see Note 4).
To prevent dehydration of the mouse during surgery, it is
recommended to administer 500 μl saline subcutaneously
during the procedure.
●
At least 30 min before the end of surgery 0.05 mg/kg
buprenorphine (or any other approved analgesic) should
be injected for postsurgical analgesia.
Surgeries must be performed in accordance with regulations and
should be conducted in clean conditions. For proper handling of
animals and recommended materials please refer to the NIH guidelines (http://www.oacu.od.nih.gov/ARAC/documents/Rodent_
Surgery.pdf). It is recommended to use a stereomicroscope
throughout the procedure for accuracy and reproducibility. It is
crucial to verify the depth of anesthesia by monitoring breathing
rate and toe-pinch reflex throughout the surgery at intervals of
10–15 min. Isoflurane clearance should be managed according to
local regulations, either through passive or active removal from
the mask.
1. Weigh the mouse and calculate the appropriate dose of preanesthesia (optional) and analgesia. Administer pre-anesthesia
or place the mouse in the isoflurane induction chamber.
2. Shave fur on mouse head using scissors or an electric hair trimmer. Trim from back of skull to slightly posterior to eyes.
3. Apply a layer of eye ointment over the whole eye area, to avoid
corneal drying during the surgery.
4. Place mouse in the animal stereotaxic apparatus:
●
Pull back skin on head and pull down bottom jaw.
●
Position head so that the upper teeth go through the slot
of the tooth bar. Pull the mouse gently backward to make
sure mouth is fastened correctly in tooth bar.
●
Position the isoflurane mask and tighten.
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5. Activate the isoflurane vaporizer. First adjust to 2 %, then
maintain at a working concentration of 1–1.5 % throughout
the procedure.
6. Monitor breathing and check toe-pinch reflex:
●
Monitoring of breathing and toe-pinch reflex should be
done throughout the surgery. Toe-pinch reflex should be
checked every 10 min but should be omitted when the
injection needle is in the brain (see below).
7. Head-fix mouse:
●
Position ear bars: Bars should be positioned either inside
the ear canal or slightly ventral and anterior to it, where
jaw meets skull. For ideal positioning, both bars should be
pushed against the skull simultaneously and from the same
starting position (see Note 5; Fig. 1-bI)
●
Adjust the height of the tooth bar and/or ear bars in order
to achieve correct placement of the skull with respect to
bars (see Note 6).
●
Once correct position has been obtained, tighten ear bars
and make sure the head is fixed tightly by gently applying
pressure to the head in a downward direction.
8. Heating pad:
●
Heating pad should be placed under the mouse and set
to 37 °C.
●
Body temperature should be monitored using a rectal probe.
!
Make sure heating pad does not overheat.
9. Sterilize scalp:
●
Use sterile cotton swabs to wash with iodine antiseptic,
then with 70 % ethanol. Repeat this step twice.
●
Stroke swab from center of the skull towards the sides.
10. Make incision:
●
Using fine-tipped tweezers pull a bit of skin upwards from
skull between the ears of mouse and make a small horizontal incision with the scissors, making sure not to damage other tissues.
●
Insert scissor tips through the cut so scissors are perpendicular to the skull and facing forward. Cut along the
midline until incision reaches between the eyes.
●
Use the bulldog clamps to clamp the skin on each side of
the incision and move laterally to reveal the entire skull
(Fig. 1-bII)
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●
Using a cotton swab, gently swab the entire skull to
remove conjunctive tissue.
●
Place syringe in stereotactic device:
–
Insert the injection syringe into its holder, making
sure that the plunger is straight and fastened.
–
If needle is beveled, rotate syringe so that the opening
faces the axis that will allow the virus to spread to target area most effectively. Example: bevel could be facing anterior if injection coordinate is relatively at the
posterior end of the intended area.
–
Lower the syringe plunger until base of plunger is
close to the bottom of the syringe.
11. Identify bregma (Fig. 1-bIII):
●
Identify the anterior part of the skull where the coronal
and sagittal sutures intersect (see Note 7).
●
Place syringe needle over bregma, touch lightly so as not
to bend the needle.
●
Zero coordinates on stereotactic device.
12. Identify lambda (Fig. 1-bIV):
●
Identify lambda as the imaginary point where the sagittal
suture would meet the lambdoid suture if it were a straight
line across (typically about 1 mm posterior to the actual
intersection point; see [64]).
●
Position syringe over lambda, touch lightly.
●
Lambda should fall on the same medial–lateral and dorsal–
ventral position as bregma.
●
Deviations in height between bregma and lambda should
not exceed 0.1 mm (see Note 8).
13. Place syringe over injection target AP and ML coordinates and
lower until lightly touching skull with needle (Fig. 1-bV).
Mark target with surgical marker if desired in order to identify
the area for drilling.
14. Craniotomy:
●
Using the micro-motor drill, make a craniotomy over the
target area by applying light pressure until the dura is visible. Manage bleeding using a cotton swab or Sugi absorbent spear.
●
Using fine tweezers or a needle with a bent tip, carefully
remove the dura from underneath the craniotomy without damaging the underlying cortex.
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15. Injection: virus withdrawal:
!
●
Precaution: Any object that has contacted the virus
suspension is considered biohazardous material and
must be treated according to the existing regulations.
Such material can be placed in a designated biohazards vial containing ethanol (70 %) or H2O2.
Using the syringe, withdraw the virus suspension at a rate
of 20–50 nl/s. It is recommended to withdraw at least 1.5
times the exact volume needed in order to ensure sufficient suspension.
16. Injection: preparation step:
!
This stage is important to perform in order to assure that
no air remains at the needle tip after virus withdrawal.
●
While syringe needle is still in the air, infuse until a drop
forms at the needle tip.
●
Use Sugi tip to absorb the drop and discard into biohazard vial.
17. Injection: Infusion:
!
At this stage, movement of the mouse, stereotax and
table should be minimized since any excessive vibration could cause movement of the needle inside the
brain tissue and leakage of virus through the gap
formed by this movement. Do not perform toe-pinch
reflex tests until the syringe is retracted from the
brain and minimize any movements of the table.
●
Recommended: Before injecting, verify the position of
bregma and reset the stereotactic coordinates to this location
since the skull or needle may have shifted in previous steps.
●
Insert syringe through the drilled hole (while confirming
correct AP and ML coordinates) and into the brain until
reaching the desired DV coordinate. This should be done
slowly once the brain is entered (at the rate of about
2 mm/min) for minimal tissue damage (Fig. 1-bVI).
●
Wait about 5 min for brain tissue to adjust to needle at
coordinate.
●
Set infusion speed to 100 nl/min and begin infusion.
●
Monitor infusion: If fluid is observed to accumulate at the
surface, stop injection, wait several minutes, and/or lower
the needle by several tens of microns before continuing
with infusion.
●
After infusion, wait 10 min for virus to diffuse away from
injection site.
●
Withdraw syringe slowly (~1 mm/min).
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18. Fiber optic implant:
●
The implantable fiberoptic lightguide (IFL) can either be
inserted into the injection area or into a predicted projection area of that region [10]. The IFL we implant has a
metal or ceramic ferrule adapter which allows for optimal
optical coupling between the fiberoptic patch-cord and
the implanted fiber.
●
The length of the fiber below the ferrule should be about
equal to the dorsoventral position of the target. It is recommended to cut the fiber using a diamond knife: score
the optic fiber with the diamond knife at the desired
length, lay the fiber on a straight surface, and tape down
its tip. Holding the ferrule, pull lightly away from taped
tip. The fiber should break at the scoreline (Fig. 1-cI).
●
Take into consideration that the tip of the fiber should
be placed about 0.5 mm above the injection site in
order to be able to illuminate maximum volume of the
virally-transduced area.
●
Replace the syringe holder arm with a cannula holder.
●
Place the IFL in the cannula holder so that the fiber tip is
pointing down towards the mouse head. Place the IFL
over bregma and reset the coordinates on the stereotax
(Fig. 1-cII).
●
Move the IFL to the target AP and ML coordinates. If the fiber
implant area is the same as the injection site, the fiber should be
above the previously drilled craniotomy (Fig. 1-cIII). If the
fiber implant area is different, proceed with drilling as mentioned above.
●
Lower the IFL slowly to the DV coordinate. Make sure
that the dura mater has been fully removed from the
area before lowering the fiber.
●
Once the IFL has been inserted into the correct coordinates, proceed with cementing. It is important not to
move the cannula guide yet.
19. Fix optic fiber implant to skull.
●
Apply a layer of C&B metabond over the entire surface of
the skull, avoiding the craniotomy region (see Note 9;
Fig. 1-cIV).
●
Allow metabond to dry (5–10 min).
●
Apply a few thick layers of dental cement on the metabond
and around the fiber and ferrule while leaving at least
5 mm from the top of the ferrule exposed (Fig. 1-cV).
This is important in order to ensure sufficient space for the
zirconia sleeve that will be attached to this area during the
experiments.
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Mathias Mahn et al.
●
Allow cement to dry (5–10 min).
●
Slowly raise the cannula holder while pressing down on
the dried cement with tweezers.
●
The secured ferrule should keep in place, while the cannula holder is being raised.
20. Closing incision:
●
With tweezers, press skin together and apply a drop of
Vetbond™ tissue glue onto the subcutaneous tissue. Start
posterior and progress anteriorly along the incision
(Fig. 1-cVI).
●
Alternatively, use surgical suture. Typically one suture is
done anterior to the implant and another one or two
posterior.
21. Remove mouse from stereotactic device and place in a separate
cage with fresh bedding and heated by a heat lamp.
●
If using isoflurane anesthesia: Before removing mouse,
remove ear bars, turn off isoflurane vaporizer, wait
1–2 min, turn off O2, then turn off active isoflurane clearance system (if used).
●
Following long surgical procedures, it is recommended to
leave mouse with O2 ventilation through the anesthesia
mask until detecting a toe pinch reflex.
22. Wash syringe:
2.5 Optogenetic
Behavioral Testing
●
Withdraw a small volume of saline into the virus-containing syringe.
●
Eject virus and saline from syringe into biohazard vial. Do
not touch syringe tip to ethanol.
●
Fill and empty the syringe three times with DDW. Repeat
this step several times with clean DDW to assure effective
removal of virus from needle. Dispose of DDW according
to biohazard regulations.
Behavioral testing can be initiated 2–4 weeks following virus injection, depending on the type of virus and its titer. If light stimulation is targeted at distal axons of the expressing cells, additional
time might be required for trafficking of the opsin to the axons.
Histological examination can be performed to evaluate the time
required for expression and trafficking of the optogenetic proteins
in the chosen paradigm. Here we briefly describe a behavioral paradigm used for evaluating the effects of optogenetic activation of
the primary mouse motor cortex following AAV-based expression
of ChR2(H134R)-mCherry in this region. The mouse used for
Viral Vector-Based Techniques for Optogenetic Modulation In Vivo
305
this behavioral experiment was treated as described above and
injected with 1 μl AAV2/5-CaMKIIα-ChR2(H134R)-mCherry in
M1 cortex [AP 1 mm; ML 1 mm (left); DV 1.5 mm]. The procedure detailed below was performed 4 weeks after injection.
2.6
Procedure
1. Position the rotary joint above the center of the open field
chamber and next to the video tracking camera.
2. Connect the long (5 m) patch cord to the laser fiber coupler
on one end and to the rotary joint at the other end.
3. Connect the second fiberoptic patch-cord from the rotary
joint so that the bare ferrule side terminates at the level of the
open field chamber.
4. Remove the mouse from the home cage and attach the fiberoptic patch-cord ferrule to the IFL.
5. Place the mouse in the center of the open field and initiate the
behavioral recording session.
6. At defined times, light can be delivered through the fiberoptic
cables by triggering the laser through TTL pulses from the
pattern generator. Depending on the laser type, digital or analog modulation can be applied to control the light power output (see Note 10).
7. At the end of the experimental session, remove the mouse
from the arena, disconnect the fiberoptic patch-cord from the
IFL, and return the mouse to the home cage.
2.7 Summary
and Expected Results
In the protocol section, we describe a procedure for viral transduction of primary motor cortex pyramidal neurons with
ChR2(H134R)-mCherry, followed by an optical fiber implantation for in vivo modulation of neuronal activity. To test the effect
of optogenetic activation of primary motor cortex pyramidal neurons in a freely moving mouse, we placed the implanted mouse in
an open field chamber and recorded the position of the mouse
during a behavioral session that lasted 30 min. At defined times, we
applied 447 nm light at different frequencies and analyzed the
changes in the direction of locomotion compared with baseline
(light-off) periods. The raw tracking data were imported into
Matlab and analyzed offline for the direction of locomotion. The
position of the mouse was sampled in 0.5 s intervals. We measured
the angle of the third point relative to the line defined by the first
and second points. The resulting vector showed an average angle
of 28° towards the contralateral side during episodes of optical
stimulation (with respect to the site of optical stimulation). While
the angle of this trajectory is clearly dependent on the sampling
frequency, the resulting estimated average radius of 6.3 cm is stable
over a wide range of sampling intervals. We demonstrate that
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Mathias Mahn et al.
Fig. 2 Optogenetic stimulation of virally transduced motor cortex. (a) Mouse implanted with fiberoptic connector, after recovery from surgery. Fiberoptic patch-cable is attached to the implant, allowing for light delivery
into the targeted region. (b) Overlay image indicating the position of the mouse before light stimulation (1),
during stimulation (2–5) and after stimulation (6). Stimulation was performed using a 447 nm diode laser at a
frequency of 13 Hz and pulse width of 5 ms for 6 s intervals. (c) Radial plot depicting the average speed and
direction of movement of the mouse during no-stimulation trials (black) compared with stimulation trials (blue).
The mouse’s body position was tracked and the average direction of motion was calculated at 2 Hz sampling
intervals. The vector length, representing mouse velocity, was normalized to control periods
activation of the left primary motor cortex leads to a bias in the
average direction of locomotion toward the right, resulting in circular walking trajectories (Fig. 2). Following behavioral testing we
verified the site of injection and observed substantial transduction
in the targeted region (Fig. 3).
3
Notes
1. Alternatively, tools can be sterilized by autoclaving or soaking
in disinfectant.
2. Brain atlases are usually of adult mice (8 weeks or older) and
variations can arise from using different strains and ages. If
mice are younger, appropriate adjustments to the target coordinates should be made. The average lambda–bregma distance
in C57BL/6J mice, reported in [64], is 4.21 mm. A scaling
factor for the stereotaxic coordinates can be obtained by dividing the actual distance in the given mouse by this value.
3. This protocol uses isoflurane for anesthesia and buprenorphine for analgesia. Alternatively, ketamine and xylazine or any
other suitable and approved anesthetic can be used.
Viral Vector-Based Techniques for Optogenetic Modulation In Vivo
307
Fig. 3 Expression pattern following 1 μl injection of AAV5-CaMKIIα-ChR2(H134R)mCherry into the mouse motor cortex. Virally transduced cells exhibit red fluorescence due to mCherry expression. (a) Coronal brain section showing injection
center (identified by injection tract) approximately 1 mm anterior to bregma. Brain
injury by the optic fiber is visible at the upper edge. Scale bar: 2 mm. (b) Dorsal
view of the brain prior to sectioning. Scale bar: 2 mm. (c) Close up of cells [box
in (a)]. Somata of transduced cells are clearly visible. Scale bar: 200 μm
4. Anesthesia induction can be performed by placing the mouse
in an induction box with 4 % isoflurane. However, light ketamine/xylazine anesthesia will allow for easier handling until
the mouse is placed on the stereotactic device and deeply anesthetized by isoflurane application.
5. The medial–lateral position normally does not require any
readjustment if the ear bars were positioned evenly. Use the
ear bar scales to verify even positioning.
6. It is advisable to first align height of lambda and bregma visually by looking at the skull from the side. If this is done correctly, little adjustment will be needed afterwards.
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Mathias Mahn et al.
7. If difficult to detect, it is possible to apply a bit of pressure with
tweezers on one of the skull bones in order to better emphasize the sutures under the stereomicroscope. Another possibility is to apply a bit of sterile 4 % H2O2 with a cotton swab.
After several seconds, wash the skull with sterile saline to stop
the reaction. Suture lines should become more visible after
~10 s of treatment.
8. If necessary, adjust skull position to align lambda and bregma.
The medial-lateral position can be adjusted by moving the ear
bars simultaneously without releasing the pressure on the
skull. If lambda is too high or too low relative to bregma,
adjust the mouse’s snout by raising/lowering the tooth bar.
9. Metabond acts as a bonding material between the scalp and
the dental cement. A thick layer should be spread on the scalp
around the implanted fiber and around the bottom part of the
ferrule. Contact between metabond and the exposed craniotomy can be eliminated by applying a layer of aqueous gel or
petroleum jelly (Vaseline) into the craniotomy prior to insertion of the IFL.
10. It is advisable to measure the light power output of the optical
system by using a portable light power meter (e.g., Thorlabs
P-1000D). This can be done using an IFL similar to the
implanted one, in order to account for light power losses at all
points of optical coupling in the path.
Acknowledgments
We thank the entire Yizhar lab for helpful comments and discussions on the manuscript and protocol. This work was supported
by grants from the Israeli Science Foundation (ISF grant
1351/12) and by the Israeli Center of Research Excellence
(I-CORE) in Cognition (I-CORE Program 51/11). Ofer Yizhar
is the incumbent of the Gertrude and Philip Nollman Career
Development Chair.
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