Two-photon single-cell optogenetic control of neuronal activity by sculpted light

We took a different approach for addressing the shortcomings of both the one-photon and the two-photon method in a fundamentally unique way and have demonstrated reliable single neuron–specific activation with temporal resolution (≤1 ms) in hippocampal slices. This was done by effectively decoupling the lateral (i.e., the waist size) and the axial (i.e., the Rayleigh length) beam parameters for two-photon absorption, which are usually coupled for a Gaussian beam, and thereby sculpting the spatial light distribution at the focal plane. For a pulsed laser source, the decoupling can effectively be achieved by using the spectrum of the pulse to control its two-photon absorption probability in the axial direction. This has been demonstrated in the technique of temporal focusing (28, 29), which has also found applications in widefield two-photon imaging (30) and in 3D multilayer super-resolution microscopy (31). Two-photon temporal focusing (TEFO) can be experimentally realized by first spatially broadening a femtosecond optical pulse using a diffraction grating. The spot on the grating is then imaged onto the sample plane using a telescope that consists of the microscope objective and an additional lens. This configuration leads to an axial geometry in which the pulse is broadened everywhere in the sample except at the image plane, where the spatially separated paths of the different frequency components in the pulse meet again and the pulse reaches its minimum width. (Fig. S1 A and B). The two-photon excitation probability in TEFO is inversely proportional to the pulse width squared, which leads to a depth of field ≈2 orders of magnitude shorter (31) than the widefield one-photon epifluorescence technique. This allows a sectioning capability close to a confocal or two-photon microscope (27), while providing simultaneous excitation of an area that is ≈3 orders of magnitude larger than the diffraction-limited spot (Fig. S1C). Therefore, TEFO can be used for simultaneous excitation of a thin disk-like volume in a biological sample. In the present work we took advantage of this fact and demonstrated that TEFO can be used for targeted single-cell excitation of ChR2 with high spatiotemporal resolution in mouse and rat hippocampal slices.

After confirming the above parameters of the excitation volume for our temporally focused beam (Figs. S1B and S2), we investigated the activation of ChR2 expressed in cultured human embryonic kidney (HEK293) (Fig. 1A) cells and measured ChR2-mediated currents with whole-cell voltage clamp recordings. Fast (≤2 ms rise time) and large (≤1 nA) inward currents mediated by ChR2 were detected using pulses between 1 and 100 ms (Fig. 1 B and C). We aimed to systematically identify the optimal parameters for two-photon temporal focusing excitation and investigated the activation of ChR2 as a function of laser power, wavelength, and beam size (Fig. 1 D–I). Keeping a constant beam diameter of 3.8 μm and pulse duration of 100 ms for a range of wavelengths between 800 nm and 960 nm, we first measured the peak amplitude and rise time of the inward current as a function of laser power (Fig. 1 D–F). At each wavelength for lower powers we observed a nonlinear dependence of ChR2 activation on power, which is the signature of the two-photon effect (27). This quadratic dependence on power is not evident at first sight over the studied range of powers, because it is overshadowed by other parameters of the channel kinetics, such as saturation and desensitization. However, under our experimental conditions it could clearly be seen for the power range between 0 and 50 mW (Fig. 1D, Inset). At high powers the relationship showed near-asymptotic behavior, likely indicating the saturation of channel activation (26). These experiments revealed the maximum two-photon activation efficiency to be at 880 nm, which is slightly blue shifted compared with the double of the one-photon absorption peak (Fig. 1F). Once the optimum wavelength was identified, we examined the dependence of the response on the beam size. Using 100-ms-long pulses at 880 nm we varied the power at three different spot sizes (6 μm, 10 μm, and 14 μm). The peak inward current increased with power for all beam sizes (Fig. 1G); however, by further reducing the spot size to 4 μm we could show that the highest efficiency of activation (as shown by the larger peak inward current and faster rise time; Fig. 1H) could be achieved with the ≈6-μm spot (Fig. 1I). This fact reflects the intrinsic tradeoff between the quadratic dependence of the probability for opening a channel on the intensity, which increases with decreasing spot size, and the total number of channels on the illuminated surface area, which decreases with the spot size. The induced peak current (or the total charge influx) is proportional to the intensity squared and the area of illumination. Therefore, for a constant photon number (or a constant power) the peak current should be inversely proportional to the square of the pot size (Fig. 1G, Inset). All cells were tested for activation of ChR2 with blue light (488 nm), which revealed comparable biophysical parameters to TEFO excitation (Fig. S3 A–E).

These results motivated us to investigate the potential of this method as a tool with high spatiotemporal resolution for neuronal circuit mapping and other neuroscience studies. We turned to hippocampal slices and used different types of ChR2-expressing neurons: CA1 pyramidal cells (CA1PCs) in Thy1-ChR2-YFP transgenic mice (32) and parvalbumin-positive interneurons (PV-INs) in PV-Cre mice (33) that were infected with a Cre-dependent ChR2-GFP containing adeno-associated virus (ChR2-GFP-AAV) (34) (Materials and Methods and SI Materials and Methods). First we aimed to determine the efficacy and basic biophysical characteristics of our technique to explore the optimal parameters for different types of applications. Somatic current-clamp recordings were performed in ChR2-expressing neurons while the soma was illuminated by a temporally focused spot of ≈5-μm diameter with 1-, 2-, 5-, 10-, and 100-ms pulse duration at 880 nm with ≈460-mW power at the sample (Fig. 2A). Similar to HEK293 cells, in both types of neurons fast (≈1–5 ms) and large (≈5–15 mV) ChR2-mediated depolarizations could be evoked using TEFO excitation (Fig. 2A). At a holding potential of −80 mV (to prevent generation of APs), the ChR2-induced membrane depolarization continuously increased with pulse duration and reached a saturation value of ≈15–20 mV for pulses longer than ≈10 ms (Fig. 2B). In these experiments we used a static beam path and moved the specimen to position the temporally focused spot for targeting a certain area. However, to produce fast spatiotemporal activation patterns, it would be advantageous to be able to move the excitation spot on a submillisecond time scale to any desired number of target points. To achieve this we integrated the temporal focusing beam into a microscope equipped with a pair of laser scanning mirrors (Materials and Methods and SI Materials and Methods). We tested this technical advance by recording from CA1PCs and subiculum pyramidal cells (SubPCs) in Thy1-ChR2-YFP mice and scanning variable spots in the slice on and around the somata and dendrites of the patched cells. As expected, ChR2-mediated responses were detected only when the excitation spot was placed precisely on the soma or the dendrite, but no response was evoked when the beam was placed away from the recorded cell (Fig. 2C). Using very short pulsed illuminations (0.1 ms) with short interpulse intervals (0.1 ms) at multiple points positioned on the soma, we could effectively increase the activated surface area and decrease the necessary time to achieve the desired depolarization level to evoke APs within a submillisecond time scale (Fig. 2D). (Further advantages of the multiple-spot activation along single dendrites are discussed in Fig. S4).

Precise spatial resolution of excitation was indicated by the elimination of ChR2-mediated depolarization when the beam was moved a few microns away from the dendrite (Fig. 3 A and B). Given that dendrites provide a spatially confined and a much smaller excitable region (≈1–2 μm diameter dendritic compartments) than the soma, they were ideal for precise measurement of the spatial resolution of our TEFO technique. We used thin apical dendrites (50–400 μm from the soma, in 50–160 μm depth) and quantitatively compared the spatial resolution of our techniques with that acquired with 488-nm light. We displaced the temporally focused spot relative to the dendrite in 3D while recording voltage signals at the soma (Fig. 3 A–D). When the TEFO excitation spot was moved laterally (x-direction) or axially (z-direction), the peak depolarization dropped off sharply to 50% of the peak response after ≈10 μm and was almost eliminated at ≈20 μm distance [Fig. 3 B, C, and D (red trace)]. However, in comparison, responses induced by 488-nm light only dropped off by ≈50% at ≈50 μm away from the dendrite (x-direction) [Fig. 3B (blue trace)] and remained almost unaltered at 100 μm above or below the dendrite (z-direction) (Fig. 3 C and D). This result demonstrates a 5-fold improvement in the lateral excitation precession and at least a 10-fold improvement in the axial excitation precession. The axial resolution could be further improved by decreasing the applied power by 50% (Fig. 3C). This is a strong indication, as also discussed in a recent publication (26), that saturation of ChR2 in the center of the target plane leads to an additional contribution of the out-of-focus regions to the evoked response.

We next quantified how the efficacy of TEFO excitation depended on the depth in the tissue. The maximum depolarization evoked by single 1-ms pulses, placed on somata located at variable depths in the slice, showed a linear attenuation with depth (Fig. 4A). However, as demonstrated above, the total induced depolarization could be increased by using the multispot temporal focusing excitation approach. As a result, when a multispot temporal focusing excitation pattern with pulsed illumination durations of 0.1–0.2 ms and interpulse intervals of 0.1 ms was used, the efficiency of the excitation was increased. As shown in Fig. 4A, this led for the studied cells to a more than 2-fold increase of depth of excitation for the same total illumination duration. This dependence of the spatiotemporal patterning of excitation was used to induce depolarizations that were strong enough to evoke APs even at the apical tuft at >150 μm depth, where depolarization induced by a single short-pulse illumination was barely detectable (Fig. 4B).

In current clamp, at the resting membrane potential (−58 to −67 mV), we set out to determine the parameters necessary to induce the minimal depolarization that was sufficient to reach AP threshold in a given cell of all three neuron types included in our study (Fig. 5A). We used either one spot with variable pulse durations (PV-INs and two of six CA1PCs; total t = 1–5 ms), or five spots with submillisecond pulses (four of six CA1PC, and all SubPC, total t = 0.1–0.5 ms × 5). In all cells tested (n = 13), total pulse duration of ≤5 ms evoked suprathreshold depolarization and AP firing (PV-INs: 3.0 ± 1.2 ms, n = 3; CA1PCs: 2.5 ± 0.8 ms, n = 6; SubPCs: 1.3 ± 0.1 ms, n = 4; Fig. 5A). We next tested the ability of our method to induce high-frequency firing in PCs and PV-INs. We found that all cell types usually fired APs at the first few stimulations in the train. Subsequently, PCs responded to the train of stimuli only intermittently (Fig. 5 B–D). Increasing pulse duration partially overcame AP failures and improved firing fidelity (Fig. 5C). This reduced firing probability is the consequence of the spatial confinement of the TEFO excitation leading to an inactivation of ChR2 at higher frequencies, an effect previously also observed for blue light excitation (10, 32, 35, 36). Only recently a new genetic manipulation of ChR2 (12) could overcome the ChR2 inactivation problem at higher frequencies of stimulation.

Our early data showing the lack of effect of glutamate receptor blockers on ChR2-evoked depolarization in Thy1-ChR2-YFP mice (Materials and Methods and Fig. 2) suggested that excitatory axons of ChR2-expressing principal cells, innervating the recorded cell, are not activated by two-photon temporal focusing within our standard parameter range. Further investigations of excitatory axonal activation in the CA1 region confirmed this fact (Fig. S5). Although we could not evoke synaptic responses from Schaffer collaterals, we hypothesized that certain types of terminals/axons with more favorable properties could be excited by TEFO. Synaptic boutons of PV-INs are possible candidates, because they are relatively large and they form multiple contacts on target cells (37), thus these could provide more surface area to the temporally focused two-photon illumination. To investigate that, we recorded from (ChR2-negative) CA1PCs that were innervated by ChR2-expressing PV-INs (mostly basket, axo-axonic, and bistratified cells) in AAV-ChR2-GFP virus-infected PV-Cre mice (Materials and Methods and SI Materials and Methods) in the presence of ionotropic glutamate receptor antagonists. The temporally focused beam spot was first placed around the soma and proximal dendrites of the recorded CA1PC, overlapping with the location of ChR2-expressing boutons provided by PV-INs. Our experimental conditions were optimized to detect GABAergic postsynaptic hyperpolarization (holding potential, −60 to −65 mV; reversal potential of chloride, −80 mV). Strikingly, using pulse durations of 1–2 ms, we could evoke inhibitory synaptic responses of 0.5- to 4.0-mV amplitude that were completely eliminated by a specific GABAA receptor antagonist (SR-95531, 10 μM, n = 3) (Fig. 6A). Increasing the pulse duration did not lead to any further increase of the inhibitory postsynaptic potential (IPSP) amplitude, suggesting that local synapses making contact with the postsynaptic soma were activated in an all-or-none fashion (Fig. 6B). One-photon excitation at 488 nm resulted in ≈6–9-mV hyperpolarization (Fig. 6 A and B), presumably because the longer axial size of the beam and scattering lead to activation of significantly more synapses. To examine whether activation of boutons elicited APs in the axon, we tried to evoke postsynaptic responses at different spatial locations of the activation beam. We marked 5 spots around the recorded CA1PC soma (Fig. 6 C–E, Soma), 10 spots in stratum pyramidale (Fig. 6 C–E, Pyr), 10 spots in proximal stratum radiatum (Fig. 6 C–E, Rad), and finally 5 spots on a ChR2-expressing interneuron soma (Fig. 6 C–E, IN). In all of the locations, synchronous activation of the spots evoked IPSPs in the postsynaptic cell, and the responses were eliminated by application of tetrodotoxin (TTX, 0.5 μM; Fig. 6D). Notably, the latency of the responses varied depending on the distance and activation time of the excitation spot (Fig. 6E), further indicating that axonal APs were elicited and propagated to evoke the synaptic responses. These results altogether demonstrate that the TEFO technique is efficient to directly activate the large (≈2–3 μm diameter) synaptic boutons (or population of boutons) arising from PV-INs (37).

Please refer to SI Materials and Methods for a detailed description. Briefly, a tunable 140-fs pulsed laser was used as the source for temporal focusing. After passing through a variable zoom telescope the beam was diffracted by a diffraction grating. The spot on the grating was imaged via a telescope consisting of an achromatic lens and the microscope objective onto the specimen plane. For the multispot excitation experiments the temporally focused beam was integrated into the scan head of a commercial two-photon scanning microscope.

Cre recombinase–dependent virus was assembled, harvested, and purified as previously described (40) and injected into the hippocampal CA1 region of parvalbumin-Cre (33) mice and hippocampal CA3 region of Sprague-Dawley rats.

Transverse slices were prepared as previously described (41), according to methods approved by the Janelia Farm Institutional Animal Care and Use Committee.