Population-Level Representation of a Temporal Sequence Underlying Song Production in the Zebra Finch.

Animals

We used adult (>90 days post hatch) male and female zebra finches that were obtained from an outside breeder and maintained in a temperature-and humidity-controlled environment with a 12 hour:12 hour light:dark schedule. All animal maintenance and experimental procedures were performed according to the guidelines established by the Institutional Animal Care and Use Committee at the New York University Langone Medical Center. Song detection and reward We recorded singing behavior with an omni-directional lavalier condenser microphone (AT803, Audio-Technica) and amplified the signal with a solid-state preamplifier (Ultragain Pro MIC2200, Behringer). Song was detected using a digital signal processor (RX8, Tucker-Davis Technologies) and custom software written using the RPvdsEx interface (Tucker-Davis Technologies) and MATLAB. Because the zebra finch produces a song containing short gaps between syllables and motifs, singing behavior was defined as time periods in which the ratio of high frequency power to low frequency power (0–1 kHz and 1–7 kHz respectively) was greater than 3 for more than 50% of a one second sliding window. Singing behavior was only evaluated during 20 second trial periods when the female zebra finch was visible (e.g. Figure 1A ). Liquid rewards were administered using a gravity-fed line though a solenoid (NResearch) that was opened for 200 ms in order to dispense approximately 20 μL of water. To evaluate the detection algorithm, we manually noted false negatives and false positives, enabling us to calculate the sensitivity [True Positives/(True Positives + False Negatives)] and positive predictive value [True Positives/(True Positives + False Positives)].

Surgical procedures

All surgical procedures were performed under isoflurane anesthesia (1–3% in oxygen) following established guidelines. Prior to the onset of training, a small (3.9 mm by 4.55 mm, 1.3 mm) stainless steel headplate with two threaded holes was affixed with dental acrylic over the inner leaflet of the skull with the posterior edge approximately 5 mm rostral to the bifurcation of the sagittal sinus. Cranial window surgery and viral injection were performed once the zebra finch demonstrated the ability to sing reliably in the head-fixed condition (see Figure 1 and S1 ). HVC was identified electrophysiolgically during surgery using a bipolar stimulating electrode placed into RA ( Long et al., 2010 ). In most birds, we injected AAV9.Syn.GCaMP6s.WPRE.SV40 (Penn Vector Core) into HVC using a beveled pipette (opening diameter: 30 μm, length of bevel: 100μm). In two birds, we injected a 1:1 mix of AAV9.CamKII0.4.Cre.SV40 and either AAV9.CAG.Flex.GCaMP6f.WPRE.SV40 (Bird #192) or AAV9.CAG.Flex.GCaMP6s.WPRE.SV40 (Bird #193). We performed 3–6 injections (30–100 nL per site) separated by approximately 300 μm using an oil-based pressure injection system (Nanoject II, Drummond Scientific). At the end of the injection procedure, we sealed a 3 mm diameter circular cover glass (#1 thickness, Warner Instrument) with Kwik-Sil adhesive (WPI) and fixed the edge of the glass with cyanoacrylate. We also cemented a black plastic ring (inner diameter: 5 mm, outer diameter: 7.5 mm) around the cranial window to prevent light contamination during imaging.

Two-photon imaging

We used a customized Movable Objective Microscope (Sutter instrument Company) to scan our field of view (frame rate: 28.8 Hz unless otherwise noted) with a resonant system (Thorlabs) and ScanImage 4.2 software (Pologruto et al., 2003). All imaging was done using a 16× water immersion objective (Nikon) with a numerical aperture of 0.8 and a working distance of 3 mm. In order to protect the microscope from external light contamination, we wrapped it with a light-attenuating material. Additionally, we fit a black balloon to the tip of the objective on one end and the black ring surrounding the optical window on the other (Dombeck et al., 2010). The excitation source was a mode-locked Ti:sapphire laser (Chameleon, Coherent) tuned at 920nm and controlled by a Pockels cell (Conoptics 302RM).Fluorescent light was detected using a GaAsP photomultiplier tube (H10770PA-40 PMT Module, Hamamatsu) and a wide detection path (2″ collection lens).

Two-photon targeted electrophysiological recordings

In vivo electrophysiological recordings of HVC neurons expressing GCaMP6s were performed in isoflurane-anesthetized zebra finches. For these birds, we did not affix the cranial window to the brain with Kwik-Sil. Rather, the glass coverslip was sealed at the edges with cyanoacrylate and a small opening (~300 μm diameter) was produced near the center of the glass using a carbide burr. Borosilicate glass pipettes were fabricated on a horizontal puller (Sutter Instrument Company) with impedance values in the range of 4–5 MΩ and filled with an solution of K-gluconate (150 mM) and 5 μM of Red Alexa 594 (Molecular Probes, Invitrogen) to visualize the pipette under the microscope. Because HVC projection neurons produce only infrequent bursts outside of the context of singing ( Hahnloser et al., 2002 ; Long et al., 2010 ), we applied 0.1 mM of the GABA A receptor antagonist gabazine (Sigma) to the surface of the craniotomy in order to increase the rate of spontaneous bursting activity ( Mooney and Prather, 2005 ). Signals were recorded using a Neurodata IR183A single channel amplifier (Cygnus Technology) and custom MATLAB acquisition software. Data were low-pass filtered at 5 kHz and digitized with a National Instruments digital-to-analog converter (acquisition rate: 40 kHz). For these experiments, the soma was scanned at 52 Hz, and these data were aligned to spiking activity.

Intracellular recordings

Intracellular recordings during singing were carried out as described previously ( Vallentin and Long, 2015 ). Briefly, a motorized intracellular microdrive was installed on the head of the zebra finch. For antidromic identification of projection neurons, we implanted a bipolar stimulating electrode into RA and/or area ×. Sharp electrodes with an impedance of 70–130 MΩ were backfilled with 3 M of potassium acetate and inserted into HVC. Acceptable recordings were defined as having a spike height greater than 40 mV, a resting membrane potential more hyperpolarized than −50 mV, and a total recording duration greater than 3 minutes. Once stable recordings were achieved, a female bird zebra finch was presented to elicit directed singing. Neurons with fewer than two song motifs were excluded from our analysis.

Surgical procedures

All surgical procedures were performed under isoflurane anesthesia (1–3% in oxygen) following established guidelines. Prior to the onset of training, a small (3.9 mm by 4.55 mm, 1.3 mm) stainless steel headplate with two threaded holes was affixed with dental acrylic over the inner leaflet of the skull with the posterior edge approximately 5 mm rostral to the bifurcation of the sagittal sinus. Cranial window surgery and viral injection were performed once the zebra finch demonstrated the ability to sing reliably in the head-fixed condition (see Figure 1 and S1 ). HVC was identified electrophysiolgically during surgery using a bipolar stimulating electrode placed into RA ( Long et al., 2010 ). In most birds, we injected AAV9.Syn.GCaMP6s.WPRE.SV40 (Penn Vector Core) into HVC using a beveled pipette (opening diameter: 30 μm, length of bevel: 100μm). In two birds, we injected a 1:1 mix of AAV9.CamKII0.4.Cre.SV40 and either AAV9.CAG.Flex.GCaMP6f.WPRE.SV40 (Bird #192) or AAV9.CAG.Flex.GCaMP6s.WPRE.SV40 (Bird #193). We performed 3–6 injections (30–100 nL per site) separated by approximately 300 μm using an oil-based pressure injection system (Nanoject II, Drummond Scientific). At the end of the injection procedure, we sealed a 3 mm diameter circular cover glass (#1 thickness, Warner Instrument) with Kwik-Sil adhesive (WPI) and fixed the edge of the glass with cyanoacrylate. We also cemented a black plastic ring (inner diameter: 5 mm, outer diameter: 7.5 mm) around the cranial window to prevent light contamination during imaging.

Supplementary Material 1 2 3