DOI: 10.5061/dryad.1zcrjdfvs
title: Motor cortex analogue neurons in songbirds utilize Kv3 subunits to generate ultranarrow spikes

description: Complex motor skills in vertebrates require specialized upper motor
 neurons with precise action potential (AP) firing. To examine how diverse
 populations of upper motor neurons subserve distinct functions and the
 specific repertoire of ion channels involved, we conducted a thorough
 study of the excitability of upper motor neurons controlling somatic motor
 function in the zebra finch. We found that robustus arcopallialis
 projection neurons (RAPNs), key command neurons for song production,
 exhibit ultranarrow spikes and higher firing rates compared to neurons
 controlling non-vocal somatic motor functions (AId neurons).
 Pharmacological and molecular data indicate that this striking difference
 is associated with the higher expression in RAPNs of high
 threshold, fast-activating voltage-gated Kv3 channels likely
 containing Kv3.1 subunits. The spike waveform and Kv3.1 expression in
 RAPNs mirror properties of Betz cells, specialized upper motor neurons
 involved in fine digit control in humans and other primates but absent in
 rodents. Our study thus provides evidence that songbirds and primates have
 convergently evolved the use of Kv3.1 to ensure precise, rapid AP firing
 in upper motor neurons controlling fast and complex motor skills.
description: <strong>Animal subjects</strong>: All of the work
 described in this study was approved OHSU’s Institutional Animal Care and
 Use Committee (Protocol #: IP0000146) and is in accordance with NIH
 guidelines. Zebra finches (<em>Taeniopygia guttata</em>) were
 obtained from our own breeding colony. All birds used were male and
 &gt; 120 days post hatch. Birds were sacrificed by decapitation and
 their brains removed. For electrophysiology experiments brains were
 bisected along the midline, immersed in ice-cold cutting solution, and
 processed as described below. For <em>in situ</em>
 hybridization experiments brains were cut anterior to the tectum and
 placed in a plastic mold, covered with ice-cold Tissue-Tek OCT
 (Sakura-Finetek; Torrance, CA), and frozen in a dry ice/isopropanol slurry
 and processed as described below.
 <strong><em>In situ
 </em>hybridization</strong>: To compare mRNA expression levels
 for <em>KCNC1, KCNC2, KCNC3, KCNC4, KCNMA1, KCNQ2, KCNQ3, KCNA1,
 KCNA2</em>, and <em>KCNA6 </em>across RA and AId, brains
 sections (thickness = 10 μm) were cut coronally on a cryostat and mounted
 onto glass microscope slides (Superfrost plus; Fisher Scientific, Hampton,
 NH, USA), briefly fixed, and stored at -80°C. For each brain, every
 10<sup>th</sup> slide was fixed and stained for Nissl using an
 established cresyl violet protocol. Slides were examined under a
 bright-field microscope to identify sections containing the core region of
 RA and AId as previously defined (Nevue et al., 2020). <em>In
 situ</em> hybridization was conducted using an established protocol
 (Carleton et al., 2014). Briefly, slides were hybridized under
 pre-optimized conditions with DIG-labeled riboprobes synthesized from
 BSSHII-digested cDNA clones obtained from the ESTIMA: Songbird clone
 collection (Replogle et al., 2008). Specific clones corresponded to
 GenBank IDs CK302978 (KCNC1; Kv3.1), DV951094 (KCNC2; Kv3.2), DV953393
 (KCNC3; Kv3.3), CK308792 (KCNC4; Kv3.4), DV954467 (KCNMA1; BK), FE737967
 (KCNA1; Kv1.1), FE720882 (KCNA2; Kv1.2), FE733881 (KCNA6; Kv1.6), DV954380
 (KCNQ2; Kv7.2), and CK316820 (KCNQ3, Kv7.2). After overnight
 hybridization, slides were washed, blocked, incubated with alkaline
 phosphatase conjugated anti-DIG antibody (1:600; Roche, Basal,
 Switzerland) and developed overnight in BCIP/NBT chromogen (Perkin Elmer;
 Waltham, MA, USA). Slides were coverslipped with VectaMount (Vector,
 Newark, CA, USA) permanent mounting medium, and then digitally
 photographed at 10X under bright field illumination with a Lumina HR
 camera mounted on a Nikon E600 microscope using standardized filter and
 camera settings. Images were stored as TIFF files and analyzed further
 using the FIJI distribution of ImageJ (Schindelin et al., 2012). We note
 that high-resolution parasagittal images depicting expression of
 <em>KCNC1, KCNC2, KCNA1, KCNA6, KCNQ2</em>, and
 <em>KCNMA1</em> in RA of adult male zebra finches are
 available on the Zebra Finch Expression Brain Expression Atlas
 (ZEBrA; www.zebrafinchatlas.org). All probes
 were evaluated for specificity by examining their alignment to the zebra
 finch genome and avoiding probes with significant cross alignments to
 other loci (as detailed previously (Lovell et al., 2020)).
 For each gene, we quantified both expression levels based on
 labeling intensity (i.e. average pixel intensity) and the number of cells
 expressing mRNA per unit area. We measured the average pixel intensity
 (scale: 0-256) in a 200 x 200 µm window placed over each target area in
 the images of hybridized sections. To normalize signal from background we
 subtracted an average background level measured over an adjacent control
 area in the intermediate arcopallium that was deemed to have no mRNA
 expression. The expression ratio was calculated as
 RA<sub>OD</sub>/AId<sub>OD</sub> where values
 greater than 1 are more highly expressed in RA and values less than 1 are
 more highly expressed in AId. We also quantified the number of labeled
 cells in each arcopallial region by first establishing a threshold of
 expression 2.5X above the background level. Standard binary filters were
 applied and the FIJI ‘Analyze Particles’ algorithm was used to count the
 number of labeled cells per 200 µm<sup>2</sup>. 
 <strong>Slice preparation for electrophysiology
 experiments</strong>: Frontal (180 μm for current clamp and 150 μm
 for voltage clamp) slices were cut on a vibratome slicer (VT1000, Leica)
 in an ice-cold cutting solution containing (in mM): 119 NaCl, 2.5 KCl, 8
 MgSO<sub>4</sub>, 16.2 NaHCO<sub>3</sub>, 10
 HEPES, 1 NaH<sub>2</sub>PO<sub>4</sub>, 0.5
 CaCl<sub>2</sub>, 11 D-Glucose, 35 Sucrose pH 7.3-7.4 when
 bubbled with carbogen (95% O2, 5% CO<sub>2</sub>; osmolarity
 ~330-340 mOsm). Slices were then transferred to an incubation chamber
 containing artificial cerebral spinal fluid (aCSF) with (in mM): 119 NaCl,
 2.5 KCl, 1.3 MgSO<sub>4</sub>, 26.2
 NaHCO<sub>3</sub>, 1
 NaH<sub>2</sub>PO<sub>4</sub>, 1.5
 CaCl<sub>2</sub>, 11 D-Glucose, 35 Sucrose pH 7.3-7.4 when
 bubbled with carbogen (95% O2, 5% CO2; osmolarity ~330-340 mOsm) for 10
 min at 37°C, followed by a room temperature incubation for ~30 min prior
 to start of electrophysiology experiments. 
 <strong>Patch clamp electrophysiology</strong>: RA
 and AId could be readily visualized in via infra-red differential
 interference contrast microscopy (IR-DIC) (Fig. 2B). Whole-cell
 patch-clamp recordings were performed at room temperature (~24°C) unless
 otherwise indicated. For experiments performed at 40°C, the bath solution
 was warmed using an in-line heater (Warner Instruments, Hamden, CT). The
 temperature for these experiments varied up to ± 2°C.
 Slices were perfused with carbogen-bubbled aCSF (1-2ml/min) and
 neurons were visualized with an IR-DIC microscope (Zeiss Examiner.A1)
 under a 40x water immersion lens coupled to a CCD camera (Q-Click;
 Q-imaging, Surrey, BC, Canada). Whole-cell voltage- and current-clamp
 recordings were made using a HEKA EPC-10/2 amplifier controlled by
 Patchmaster software (HEKA, Ludwigshafen/Rhein, Germany). Data were
 acquired at 100 kHz and low-pass filtered at 2.9 kHz. Patch pipettes were
 pulled from standard borosilicate capillary glass (WPI, Sarasota, FL, USA)
 with a P97 puller (Sutter Instruments, Novato, CA). All recording pipettes
 had a 3.0 to 6.0 MΩ open-tip resistance in the bath solution.
 Electrophysiology data were analyzed offline using custom written routines
 in IGOR Pro (WaveMetrics, Lake Oswego, OR, USA).  For
 current clamp recordings, intracellular solutions contained (in mM): 142.5
 K-Gluconate, 21.9 KCl, 5.5 Na2-phosphocreatine, 10.9 HEPES, 5.5 EGTA, 4.2
 Mg-ATP and 0.545 GTP, pH adjusted to 7.3 with KOH, ~330-340 mOsm. Synaptic
 currents were blocked by bath applying Picrotoxin (100 μM), DL-APV (100
 μM), and CNQX (10 μM) (Tocris Bioscience) for ~3 min prior to all
 recordings. To initiate current clamp recordings, we first established a
 giga-ohm seal in the voltage clamp configuration, set the pipette
 capacitance compensation (C-fast), and then set the voltage command to -70
 mV. We then applied negative pressure to break into the cell. Once stable,
 we switched to the current clamp configuration. Experiments in current
 clamp were carried out within a 15-min period. AP half-width was defined
 as the width of the AP halfway between threshold (when the rate of
 depolarization reaches 10 V/s) and the AP peak. The maximum depolarization
 and repolarization rates were obtained from phase plane plots generated
 from averaged spontaneous APs. We noted that the resting membrane
 potential tended to hyperpolarize to the same degree (~10 mV) in both RA
 and AId after positive current injections during these current clamp
 recordings (Alexander, Mitry, Sareen, Khadra, & Bowie, 2019).
 Recordings in which the resting membrane potential deviated by &gt; 10
 mV were discarded. We note that recordings were not corrected for a
 calculated liquid junction potential of +9 mV. 
 Estimated current clamp measurements of membrane capacitance
 (C<sub>m</sub>) were calculated from the measured membrane
 time constant (t<sub>m</sub>; fit with a
 double exponential at the onset of a negative current injection) and input
 resistance (R<sub>in</sub>; calculated slope of V-I plot)
 using the equation: t<sub>m</sub>(<em>ms</em>) = R<sub>in</sub>(MΩ) •
 C<sub>m</sub>(pF) For voltage
 clamp recordings we attempted to limit the speed and space clamp error by
 1) Cutting thinner slices (~150 µm) to eliminate more processes, 2)
 decreased the intracellular K+ concentration to decrease the driving force
 and 3) compensated the series resistance to 1MΩ. The intracellular
 solutions contained the following (in mM): 75 K-Gluconate, 5.5
 Na2-phosphocreatine, 10.9 HEPES, 5.5 EGTA, 4.2 Mg-ATP, and 0.545 GTP, pH
 adjusted to 7.3 with KOH, adjusted to ~330-340 mOsm with Sucrose. Rs was
 compensated to 1 MΩ, the uncompensated Rs = 7.7 ± 0.4 MΩ (Mean ± SE; N=12
 RAPNs and AId neurons). We ensured exclusion of interneurons by briefly
 observing the AP waveform in current clamp prior to switching to voltage
 clamp (Supplementary Fig. 2; (Zemel et al., 2021)). In order to isolate K+
 currents, slices were exposed to bath applied CdCl2 (100 μM), TTX (1 μM),
 Picrotoxin (100 μM), CNQX (10 μM), and APV (100 μM) for ~5 min prior to
 running voltage clamp protocols. After protocols were applied, TEA (500
 μM) was bath applied and the same voltage-clamp protocols were repeated
 after K+ currents were eliminated. K+ currents were isolated by
 subtracting the TEA-insensitive current traces from the initial traces.
 Capacitive currents generated during voltage-clamp recordings were
 eliminated by P/4 subtraction. Recordings were not corrected for a
 measured liquid junction potential of +12 mV. 
 <strong>Morphology</strong>: For these experiments
 Biocytin (4mg/ml; Sigma, St. Louis, MO, USA) and Alexafluor 488 (Life
 Technologies, Carlsbad, CA, USA) were included in the intracellular
 solution used in current clamp experiments. Upon entering the whole cell
 current clamp configuration, the current was set to 0 pA and the cell was
 held in for 20 to 30 minutes at room temperature to allow for complete
 filling. The electrode was then slowly removed at a diagonal angle as the
 fluorescence was monitored to determine when the electrode detached. Upon
 complete separation of the electrode from the cell, the slice was placed
 in 4% paraformaldehyde overnight. The slice was then washed in PBS with
 0.25% Triton-100x (3x for 10 minutes), blocked in a PBS solution
 containing 1% skim milk (1 hr) and then stained in a PBS solution
 containing 1% skim milk and avidin conjugated to Alexafluor 594 (1:200,
 Life Technologies, Carlsbad, CA, USA) for 2 hrs. The slice was then washed
 in PBS with 0.25% Triton-100x (3x for 10 minutes) before a final wash in
 PBS followed by mounting on a cover slip. Images were captured with a
 Zeiss LSM 980 Airyscan 2 confocal microscope.  Maximum
 projections of the images were made and analyzed in FIJI-Image J
 (Schindelin et al., 2012) to calculate the 2-D soma area, dendritic
 complexity (D. A. Sholl, 1956), and estimates spine density (as measured
 from averaged 30 µm stretches from multiple tertiary branches in each
 cell) shown in Fig. 1. In a subset of these neurons (3 RAPNs and 2 AId
 neurons) we used the recently developed, open-source software, ShuTu (Jin
 et al., 2019), to reconstruct the morphologies in 3-D (see examples in the
 Supplementary Fig. 1 A-B; Video1-2). 3-D renderings were expanded 2-fold
 in the Z-direction to allow for better visualization of cellular
 processes. Despite the planar appearance of reconstructed neurons,
 previous imaging work in RAPNs and hippocampal neurons suggest these
 dendrites tile equally across planes (Spiro et al., 1999) and have been
 disproportionately compressed in the z-axis as a result of tissue
 processing (Pyapali, Sik, Penttonen, Buzsaki, & Turner, 1998)
 respectively.  We used ShuTu’s automatic reconstruction script to trace
 high-contrast neurites filled with biocytin and to estimate their
 diameters. For low-contrast, broken or occluded processes, we manually
 corrected the reconstruction in ShuTu’s GUI. The reconstructions were
 stored in SWC format. We used ShuTu’s GUI to manually trace all visible
 dendritic spines. We annotated spines with thin necks and mushroom-like
 shapes using single SWC points projected away from the dendrite and
 following the spine’s neck direction. For spines with curved necks,
 irregular shapes or filopodia-like spines, we used multiple SWC points
 tracing their entire extent. We estimated spine areas in two ways. We
 treated spines annotated by a single SWC point as spheres attached by
 cylindrical necks with an average radius of 0.5 pixels (0.066 microns).
 For spines with multiple SWC points, we used the trapezoidal cylinders
 defined by every pair of consecutive SWC point. We also added a dome cap
 to the last SWC point. To obtain a continuous profile of spine density
 along each dendritic branch (Supplementary Fig. 1E), we used a rolling
 window of width 20 SWC points (window length fluctuated between 5 and 39
 µm, with an average of 17 µm for all cells) and of step size 1 SWC point.
 From the continuous spine density profile, we computed the average spine
 density of each dendritic segment (Supplementary Fig. 1C). We excluded
 from the density estimation of Supplementary Fig. 1C segments branching
 off the soma with fewer than 10 spines and any segment shorter than 20
 microns. We estimated the total dendritic area and length of each cell
 (Supplementary Table 1) from the reconstructions by constructing
 trapezoidal cylinders for each pair of consecutive SWC point. Axonal area
 and length were estimated in the same way, although we were only able to
 connect a small fraction of axon cable to the soma for 3 neurons. For 2
 neurons we were not able to trace any portion of the axon (Supplementary
 Table 1). We estimated the surface area of the soma of
 each neuron by triangulating its 3-D structure. We first used ShuTu’s GUI
 (Jin et al., 2019) to manually trace binary masks of the somas in all
 slices. Next, we traced the contours of the masks with edge detecting
 filters. We then took a fixed fraction of 20 equally spaced points in each
 contour in order to generate polygonal contours.  All polygonal contours
 in the end had the same number of edges. Next, we applied a 3-point moving
 average filter along each sequence of vertex in the Z direction to smooth
 out rough edges. We then calculated the areas of the triangles formed by
 the edges and vertices of each adjacent pair of polygonal contours. We
 added to this number the surface areas of the first and last non-zero
 binary masks to represent the caps. We also corrected for the areas of the
 surface patches where the dendrites attach to the soma. Since these
 surface patches are small, we approximated them by the cross-sectional
 area of the first dendrite SWC point connected to the soma.
 We estimated the volumes of the dendrites using 3-D binary masks
 which we constructed from the SWC structure. First, we re-scaled the SWC
 along the Z-axis to match the scale in the XY plane. Next, we interpolated
 the space between each connected pair of SWC points by trapezoidal
 cylinders with radii equal to the SWC points’ radii. The binary mask
 assigns ones to voxels intersecting either a SWC point or a trapezoidal
 cylinder and zeros otherwise. The volume of the dendrite is the fraction
 of active voxels in the mask multiplied by the volume of the
 voxel. We estimated the volumes of the somas using an
 adaptation of our method for the surface areas.  Starting from the
 polygonal contours representing the Z-stack, we used the polygons’
 vertices and baricenters to split the soma’s volume into a collection of
 tetrahedrons. The vertices of an edge in a slice, the vertices of the
 closest edge in the next slice, and the baricenters of the two contours
 define 4 tetrahedrons for which the volume can be easily calculated. The
 total volume of the soma is the sum of the volumes of the individual
 tetrahedrons. We approximated the volumes of the spines
 directly from the SWC structure. Since the spines are small, we estimated
 their volumes to be approximately the sum of the volumes of the individual
 SWC points. <strong>Comparative genomics of the
 KCNC/Kv3 (Shaw-related) potassium channel genes</strong>: To
 identify the full set of genes comprising this gene family in zebra
 finches, we first retrieved all genes annotated as KCNCs (voltage gated
 channel subfamily C members) from the latest RefSeq database for zebra
 finch (Annotation Release 106; GCF_003957565.2 assembly). We next
 retrieved the similarly annotated genes in other selected songbird and
 non-songbird avian species, observing the immediate synteny and correct
 cross-species BLAST alignments. To verify the correct orthology of avian
 genes to corresponding members of this gene family in mammals, we
 conducted cross-species BLAST searches, noting the top-scoring reciprocal
 cross-species alignments as well as conserved synteny as orthology
 criteria.  To build cladistic trees for evolutionary inferences, we also
 included (as outgroups to birds and mammals) representative extant
 organisms from selected branches of major vertebrate groups, as
 appropriate, including non-avian sauropsids (crocodiles, turtles,
 lizards), amphibians, and bony fishes.
 <strong>Pharmacologic compounds</strong>: DL-APV,
 Picrotoxin, CNQX, XE991 and iberiotoxin were purchased from Tocris
 Biosciences (Bristol, United Kingdom). TEA and 4-AP were purchased from
 Sigma -Aldrich (St. Louis, Missouri, USA). α-DTX was purchased from
 Alomone labs (Jerusalem, Israel). AUT5 was provided as a gift from
 Autifony Therapeutics (Stevenage, United Kingdom).
description: Electrophysiology data were analyzed offline using IgorPro
 software (Wave-metrics). Statistical analyses were performed using Prism
 4.0 (GraphPad). <em>In situ</em> hybridization data was
 analyzed using FIJI Image J (NIH). Morphology analysis was done using
 ShuTu software (http://personal.psu.edu/dzj2/ShuTu/).