1. Developmental Biology
  2. Neuroscience
Download icon

Spinal lumbar dI2 interneurons contribute to stability of bipedal stepping

  1. Baruch Haimson
  2. Yoav Hadas
  3. Nimrod Bernat
  4. Artur Kania
  5. Monica A Daley
  6. Yuval Cinnamon
  7. Aharon Lev-Tov  Is a corresponding author
  8. Avihu Klar  Is a corresponding author
  1. Department of Medical Neurobiology, IMRIC, Hebrew University – Hadassah Medical School, Israel
  2. Institut de recherches cliniques de Montréal (IRCM), Canada
  3. Ecology and Evolutionary Biology, University of California, Irvine, United States
  4. Institute of Animal Science Poultry and Aquaculture Sci. Dept. Agricultural Research Organization, The Volcani Center, Israel
Research Article
  • Cited 0
  • Views 671
  • Annotations
Cite this article as: eLife 2021;10:e62001 doi: 10.7554/eLife.62001

Abstract

Peripheral and intraspinal feedback is required to shape and update the output of spinal networks that execute motor behavior. We report that lumbar dI2 spinal interneurons in chicks receive synaptic input from afferents and premotor neurons. These interneurons innervate contralateral premotor networks in the lumbar and brachial spinal cord, and their ascending projections innervate the cerebellum. These findings suggest that dI2 neurons function as interneurons in local lumbar circuits, are involved in lumbo-brachial coupling, and that part of them deliver peripheral and intraspinal feedback to the cerebellum. Silencing of dI2 neurons leads to destabilized stepping in posthatching day 8 hatchlings, with occasional collapses, variable step profiles, and a wide-base walking gait, suggesting that dI2 neurons may contribute to the stabilization of the bipedal gait.

Introduction

The spinal cord integrates and relays the somatosensory inputs required for further execution of complex motor behaviors. Interneurons (INs) that differentiate at the ventral progenitor domain, V3-V0, are involved in the control of rhythmic motor activity, alternating between the left and right limbs, as well as between the flexor and extensor muscles (Lai et al., 2016; Osseward and Pfaff, 2019; Alaynick et al., 2011). Some of the dorsally born INs, dI1, dI3, and dI6, migrate ventrally and are also assembled within circuitries that control motor activity (Yuengert et al., 2015; Bui et al., 2013; Andersson et al., 2012), while other dorsal progenitor neurons, dI4 and dI5, give rise to INs that mediate somatosensation (Lai et al., 2016).

dI2 neurons originate in the dorsal spinal cord. The progenitor pdI2 cells, topographically positioned between the adjacent dorsally located dI1 and ventrally located dI3 neurons, express Ngn1, Ngn2, Olig3, and Pax3 transcription factors (TFs). Early postmitotic dI2 neurons undergo ventral migration and are defined by the combinatorial expression of Foxd3+/Lhx1+/Pou4f1+ TFs (Alaynick et al., 2011; Morikawa et al., 2009; Francius et al., 2013). Importantly, none of these TFs is specific to dI2 neurons; rather, their combinatorial expression defines dI2. The lack of dI2-specific cell fate markers and the dynamic expression of the above TFs in other INs causes ambiguity regarding the molecular profile and outcome of late postmitotic dI2 neurons. Using intersectional genetics, we have shown previously that dI2 neurons are commissurally projecting neurons (Avraham et al., 2009). The lack of dI2-specific TFs impeded their genetic targeting. Hence, little is known about the wiring and physiological function of dI2 neurons.

The maintenance of stability and the coordination, precision, and timing of movements are regulated and modulated by the cerebellum. Anatomical and electrophysiological studies of cats and rodents revealed two major pathways ascending from neurons in the lumbar spinal cord to the cerebellum: the dorsal spinocerebellar tract (DSCT) and ventral spinocerebellar tract (VSCT). DSCT neurons are considered to relay mainly proprioceptive information, while VSCT neurons are thought to relay internal spinal network information to the cerebellum along with proprioceptive data (Jankowska and Hammar, 2013; Spanne and Jörntell, 2013; Stecina et al., 2013; Jiang et al., 2015). While subpopulations of DSCT neurons are genetically accessible (Hantman and Jessell, 2010), the genetic inaccessibility of VSCT neurons hinders efforts to reveal their actual contribution to the regulatory functions of the cerebellum in locomotion and other motor behaviors.

In the present study, we investigated the possible functions of dI2 neurons in chick motor behavior. Employing intersectional genetics in the chick spinal cord, we targeted dI2 neurons and found evidence implicating them in the control of stability during locomotion. There are several advantages to performing these studies in chicks. The patterning of neurons within the spinal cord (Jessell, 2000) and the spinocerebellar tracts (Furue et al., 2010; Furue et al., 2011; Uehara et al., 2012) is conserved between mammals and birds. In addition, chicks use bipedal locomotion (evolved in humans and birds) that can be examined soon after hatching. To decode the circuitry and function of spinal INs, we developed a unique circuit-deciphering toolbox that enables neuron-specific targeting and tracing of circuits in the chick embryo (Hadas et al., 2014), and we utilized kinematic analysis of overground bipedal stepping by the hatched chicks following silencing of dI2 neurons.

Our studies revealed that lumbar dl2 neurons receive synaptic inputs from inhibitory and excitatory premotor neurons (pre-MNs) and relay output to the cerebellar granular layer, pre-MNs in the contralateral spinal cord and the contralateral dI2 cells. In a kinematic analysis of overground stepping by posthatching day (P) 8 hatchlings after inhibition of dI2 neuronal activity by expression of the tetanus toxin light chain gene, the genetically manipulated hatchlings showed an unstable gait, demonstrating that dI2 neurons play a role in shaping and stabilizing the bipedal gait.

Results

To define potential VSCT neurons within spinal INs, we defined the following criteria: (1) soma location in accordance with precerebellar neurons at the lumbar level, which were previously revealed by retrograde labeling experiments of the chick cerebellar lobes (Furue et al., 2010; Furue et al., 2011; Uehara et al., 2012), (2) commissural neurons, (3) excitatory neurons, and (4) non-pre-MNs (Lai et al., 2016; Osseward and Pfaff, 2019; Alaynick et al., 2011). Based on these criteria, dI1c and dI2 neurons are likely candidates (Bermingham et al., 2001; Yuengert et al., 2015; Figure 1—figure supplement 1). This is further supported by Sakai et al., 2012, who demonstrated that in the embryonic day (E) 12 chick, the dI1 and dI2 axons project to the hindbrain and toward the cerebellum. In the current study, we focused on deciphering the circuitry and function of dI2 neurons and their possible association with VSCT.

dI2 INs are mainly excitatory neurons with commissural axonal projections

To label dI2 neurons, axons, and terminals in the chick spinal cord, we used intersection between enhancers of two TFs expressed by dI2 – Ngn1 and Foxd3 – via the expression of two recombinases (Cre and FLPo) and double conditional reporters (Figure 1—figure supplement 1A). We have shown previously that the combination of these enhancers reliably labels dI2 neurons (Avraham et al., 2009; Hadas et al., 2014). The recombinases and the double conditional reporter plasmids were delivered via spatially restricted electroporation to the lumbar spinal cord at HH18. At E5, early postmitotic dI2 neurons migrate ventrally from the dorsolateral to the midlateral spinal cord (Figure 1A). As they migrate ventrally, at E6, dI2 neurons assume a midlateral position along the dorsoventral axis (Figure 1B). Subsequently, dI2 neurons migrate medially, and at E17, comparable to postnatal day 4 (P4) of mice, most of them (70.7 and 71.5% at the sciatic and the crural levels, respectively) occupy lamina VII (Figure 1C and F). At all rostrocaudal levels and embryonic stages, dI2 axons cross the floor plate (Figure 1A–D). After crossing, dI2 axons extend rostrally for a few segments in the ventral funiculus (VF) and subsequently turn into the lateral funiculus (LF) (Avraham et al., 2009; Figure 1C and D). Collaterals originating from the crossed VF and LF tracts invade the contralateral spinal cord (Figure 1C and D; white arrows).

Figure 1 with 3 supplements see all
Characterization and classification of dI2 neurons during embryonic development.

dI2 interneurons (INs) were labeled as cells that expressed both the Foxd3 and Ngn1 enhancers (Avraham et al., 2009; see Figure 1—figure supplement 1A). (A–D) dI2 axonal projection during development. At embryonic day (E) 5 (A), postmitotic dI2 neurons assume a dorsolateral position and start to migrate ventrally. At E6 (B), dI2 neurons occupy the midlateral domain. At E15-17, dI2 neurons are located at medial lamina VII at the lumbar level (LS3) (C) and thoracic level (T1) (D). dI2 axons cross the floor plate (yellow arrowheads), turn longitudinally at the ventral funiculus (white arrowheads) and eventually elongate at the lateral funiculus (white arrows). (E) Cross-section of an E17 embryo at the lumbar spinal cord (crural plexus level, LS2). Small-diameter dI2 neurons residing in lamina VII (E′) and ventromedial large-diameter dI2 neurons in lamina VIII (E″). (F) Density plots and laminar distribution (F′) of dI2 somata at the sciatic plexus level (cyan, N = 374 cells); large-diameter (magenta) and small-diameter dI2 (yellow) INs (N = 33 and N = 344 cells, respectively, from two embryos). (G, H) Neurotransmitter phenotype of dI2 neurons. dI2 neurons expressing GFP were subjected to in situ hybridization using the Vglut2 probe (G) or the VIAAT probe (H). (I) Distribution of excitatory (vGlut2, red) and inhibitory (VIAAT, blue) dI2 neurons at the sciatic and crural levels at E17 (N = 172 and 136 neurons, respectively, from two embryos). See Figure 1—source data 1 and 2.

Figure 1—source data 1

Localization of dI2 neurons at the sciatic level.

The X/Y coordinates of dI2, large-diameter dI2, and small-diameter dI2 neurons (Figure 1F).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig1-data1-v2.xlsx
Figure 1—source data 2

Localization of excitatory and inhibitory dI2 neurons.

The X/Y coordinates of dI2 neurons expressing either vGlut2 or VIATT (Figure 1I).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig1-data2-v2.xlsx

A recent study suggested that dI2 neurons at early stages of development in mice (E9.5–E13.5, comparable to chick E4–8) can be divided into several subclasses based on their genetic signature and degree of maturation (Delile et al., 2019). To assess the diversity of dI2 neurons in the chick, the expression of dI2 TFs in dI2::GFP cells was analyzed at E5 before and during ventral migration, at E6 and E14. The early postmitotic dI2::GFP cells at E5 were a homogenous population defined by Foxd3+/Lhx1+/Pou4f1+/Pax2- (Figure 1—figure supplement 2A, B, E). dI2 neurons that underwent ventral migration at E5, as well as at E6 and E14, express variable combinations of Lhx1, Pou4f1, and FoxP1/2/4 (Figure 1—figure supplement 2C–E). At E14, approximately 50% of dI2::GFP cells did not express any of the tested TFs (Figure 1—figure supplement 2E), suggesting that the early expression of TFs is required for cell fate acquisition, axon guidance, and target recognition, while their expression is not required after the establishment of the circuitry, as shown for other spinal INs (Bikoff et al., 2016). Interestingly, approximately 12% of ventrally migrating dI2 neurons (from E5 to E14) expressed Pax2 (Figure 1—figure supplement 2D and E). Pax2 is associated with a GABAergic inhibitory phenotype (Cheng et al., 2004), suggesting that a subpopulation of dI2 are inhibitory neurons. The distribution of excitatory and inhibitory dI2 neurons is also apparent at E17. In situ hybridization on cross-sections of the E17 dI2::GFP-labeled lumbar spinal cord using the vGlut2 probe revealed that 73% were vGlut2+, while the VIAAT probe, which labels GABAergic and glycinergic inhibitory neurons, measured 27% VIAAT+ dI2 neurons (Figure 1G–I; N = 308 neurons from two embryos). Similar percentages of Gad2- and Slc6a5-dI2-expressing cells were also found in mice (Delile et al., 2019). At E13–17 at the caudal lumbar level and at the level of the sciatic plexus, most dI2 neurons are located in the medial aspect of lamina VII. Approximately 91% of dI2 neurons are small-diameter neurons located in the lateral dorsal aspect of lamina VII, and 9% are large-diameter neurons. At the lumbar sciatic plexus level, large-diameter dI2 neurons are located mostly at the ventral aspect of lamina VII (Figure 1F) and at the level of the crural plexus in the ventral and dorsal aspect of lamina VII (Figure 1—figure supplement 3A). The location of the lumbar dI2 neurons, mainly lamina VII, is also apparent in posthatching chicks (P8, Figure 1—figure supplement 3C and D). Importantly, large-diameter dI2 neurons were apparent only at the lumbar level (Figure 1F, Figure 1—figure supplement 3A). The division of large- and small-diameter lumbar dI2 neurons was not reflected in the expression of the tested TFs or in a specific neurotransmitter phenotype; the inhibitory/excitatory ratios were 0.297 ± 0.13 and 0.343 ± 0.02, for large- and small-diameter dI2 neurons, respectively. Hence, dI2 neurons consist of several subpopulations, as has been shown in other spinal INs (Bikoff et al., 2016; Delile et al., 2019; Sweeney et al., 2018).

Subpopulation of dI2 neurons project to the cerebellum

To study the supraspinal targets of dI2 neurons, axonal and synaptic reporters were expressed in lumbar dI2 neurons (Figure 2A). At stage HH18 (E3), dI2 enhancers were co-electroporated with the double conditional axonal reporter membrane-tethered Cherry and the synaptic reporter SV2-GFP (Figure 1—figure supplement 1A). Expression in the lumbar spinal cord was attained by using thin electrodes positioned near the lumbar segments. At E17, the stage in which the internal granule layer is formed in the chick cerebellum, the axons and synapses of dI2 neurons were studied. dI2 axons cross the spinal cord at the floor plate at the segmental level, ascend to the cerebellum, enter through the superior cerebellar peduncle, and cross back to the ipsilateral side of the cerebellum (Figure 2B). Synaptic boutons were noticeable in the granule layer at the ipsilateral and contralateral sides of the anterior cerebellar lobules (Figure 2C). Synaptic boutons were also present in the central cerebellar nuclei (Figure 2—figure supplement 1A).

Figure 2 with 1 supplement see all
dI2 neurons project to the cerebellum.

(A) Experimental setup for labeling dI2 neurons that project to the cerebellum. dI2 neurons were genetically targeted at HH18, and precerebellar neurons were labeled using intracerebellar injection of replication-defective HSV-LacZ or PRV-Cherry at embryonic day (E) 15. The abbreviations in the coordinates: R: rostral; C: caudal. (B) A cross-section of E17 brainstem and cerebellum. The dashed polygon in (B′) is magnified in (B). dI2 axons reach the cerebellum, enter into it via the superior cerebellar peduncle, and cross the cerebellar midline. Calbindin (Purkinje neurons, magenta [B′] or red [B]). Abbreviations in the coordinates: D: dorsal; V: ventral. (C) A cross-section of E17 cerebellar cortex. Lumbar-originating dI2 synapses (cyan) in the granular layer of the anterior cerebellar cortex. Calbindin (Purkinje neurons, magenta), synaptotagmin (yellow). (D) A cross-section of an E15 embryo at the lumbar spinal cord level (sciatic plexus level). Precerebellar neurons were infected and labeled with HSV-LacZ (magenta), and dI2 neurons expressed GFP (cyan). A large-diameter dI2 neuron coexpressing LacZ and GFP is indicated by an arrow (magnification of the two channels in the insets). (E) Density plots of dI2 and precerebellar neurons (density values 10–90%) in the sciatic plexus segments (N = 374 and N = 289 cells, respectively). (F) PRV-Cherry-labeled precerebellar neurons (magenta) are in contact with dI2 axonal terminals (cyan). (G) Density plots of dI2 synapses and precerebellar neuron somata (density values 10–90%) in the sciatic plexus segments (N = 4735 synapses and N = 289 cells, respectively). (H) The laminar distribution of precerebellar neurons, dI2 neurons, and dI2 synapses at the sciatic level. See Figure 2—source data 1.

Figure 2—source data 1

Localization of precerebellar neurons and dI2 synapses at the sciatic level.

The X/Y coordinates of precerebellar neurons (Figure 2E) and dI2 synapses (Figure 2G) at the sciatic level.

https://cdn.elifesciences.org/articles/62001/elife-62001-fig2-data1-v2.xlsx

The difference in soma size between dorsally and ventrally located dI2 neurons prompted us to test which dI2 neurons project to the cerebellum. dI2 neurons and precerebellar neurons were colabeled by genetic targeting of dI2 at early stages of embryogenesis (HH18), coupled with intracerebellar injection of replication-defective HSV-LacZ at E15 or PRV-Cherry (Figure 2A). Cholera toxin subunit B (CTB) was coinjected with PRV-Cherry to verify primary infection of precerebellar neurons. Spinal neurons retrogradely labeled from the cerebellum consist of double-crossed VSCT neurons and ipsilaterally projecting DSCT neurons. However, dI2 neurons, double labeled by genetic targeting and retrograde labeling from the cerebellum, are all VSCT neurons since dI2 neurons are commissural neurons. The soma distributions of precerebellar neurons, dI2 neurons, and dI2 synapses overlapped at the sciatic level (Figure 2E, G and H) and to a lesser extent at the crural level (Figure 2—figure supplement 1B–D). We found that large-diameter dI2 neurons were mostly colabeled, most of them in the ventral aspect of lamina VII (Figure 2D). Interestingly, many of the cherry+ and LacZ+ neurons were contacted by dI2 axons (Figure 2F), suggesting that dI2 neurons innervate precerebellar neurons.

Segmental crossing at the lumbar level and recrossing back to the ipsilateral side at the cerebellum are characteristic of the VSCT projection pattern. To measure the proportion of precerebellar neurons in dI2, we used sparse labeling of lumbar dI2 neurons coupled with whole-mount imaging of the E13 spinal cord from the sacral level to the cerebellum utilizing light-sheet microscopy and iDISCO imaging (Belle et al., 2014; Renier et al., 2014; Figure 3; Video 1, Video 2, Video 3, Video 4). 85 neurons were labeled at the lumbar level, 17 of which were large-diameter neurons. The axons of all dI2 neurons cross the midline. Longitudinally projecting axons were apparent at the contralateral VF and LF (Figure 3A–F; Video 1, Video 2, Video 3). In the gray matter, axonal ramifications originating from collateral branching were apparent along the entire extent of the spinal cord (Figure 3A–C; Video 1, Video 2, Video 3). Bifurcation of the longitudinal axons was also apparent. The bifurcating branch elongated rostrally for a few segments and subsequently turned transversely into the spinal cord (Figure 3—figure supplement 1A). We counted the numbers of longitudinally projecting axons at different levels (Figure 3A and D-H): 22 axons at LS1 (Figure 3D), 30 at T1 (Figure 3E), 21 at C9 (Figure 3F), 8 at the rostral brain stem (Figure 3G), and 7 entering the cerebellum via the superior cerebellar peduncle (Figure 3H). The elevated number of axons in T1 levels likely reflects longitudinal bifurcations. Considering that the spinal cord was observed at E13, a relatively early stage of development, it is likely that additional axons enter the cerebellum at later stages of development.

Figure 3 with 1 supplement see all
3D reconstruction of dI2 neurons along the rostrocaudal axis.

(A) Spinal cord scheme describing dI2 axonal projection along the rostrocaudal axis (caudal is to the left, and rostral is to the right). The full lines represent the lumbar and brachial levels shown in (B) and (C). The broken lines represent the cross-sections shown in (D–G). The number of axons (and the funicular division) along the rostrocaudal axis is indicated adjacent to the corresponding letters (D–G). (B, C) Two representative dI2 neurons projecting their axons in the lateral funiculus (LF; green) and ventral funiculus (VF; yellow) at the lumbar (B) and brachial (C) levels. Numerous axonal collaterals are apparent. (D–F) Cross-sections at different levels of the spinal cord showing dI2 axons exiting the rostral end of the lumbar segments (D), entering the caudal brachial level (E), and exiting the rostral brachial level (F). Green: LF on the contralateral side (cLF); orange: VF on the contralateral side (cVF); cyan: LF on the ipsilateral side (iLF). (G–J) dI2 axons in the brainstem and cerebellum. (G) Axons entering the brainstem are indicated in green. (H) dI2 axons enter the cerebellum via the superior cerebellar peduncle (SCP). (I) Collaterals projecting into the brainstem. (J) The axons cross the cerebellar midline back to the ipsilateral side (two representative axons). A coordinate system is supplied in (B–G, I, J).

Video 1
dI2 interneurons (INs): transverse sections of lumbar segments.

Caudal to rostral transverse images of light-sheet microscopy images along the lumbar segments of an embryonic day (E) 13 spinal cord expressing Cherry in dI2 neurons (a reference to the location of the section is shown at the bottom left). dI2 cell bodies and axons are visible. Examples of large- and small-diameter dI2 neurons are indicated by arrows. The concentration of axons on the side contralateral to electroporation is clear.

Video 2
dI2 interneurons (INs): lumbar segments.

3D reconstruction of light-sheet microscopy images of the lumbar spinal cord. In the transparent mode, dI2 axons are apparent on the contralateral side. The trajectory of two representative axons (ventral and lateral projection axons in yellow and green, respectively) was reconstructed. Significant branching is apparent. All the axons exiting the lumbar segments are visible. A coordinate system is supplied in key frames.

Video 3
dI2 interneurons (INs): brachial segments.

3D reconstruction of light-sheet microscopy images of dI2 axons entering and exiting the brachial spinal cord. Two representative axons (ventral and lateral projection axons in yellow and green, respectively) were followed, and their collaterals along the spinal cord are demonstrated. A coordinate system is supplied in key frames.

Video 4
dI2 interneurons (INs): brainstem and cerebellum.

3D reconstruction of light-sheet microscopy images of dI2 axons projecting into the brain stem and the cerebellum (blue and red, respectively). Cerebellum midline crossing is demonstrated for two representative axons. The axonal collaterals to the brainstem are apparent. A coordinate system is supplied in key frames.

To evaluate the complexity of the branching pattern, we reconstructed the axonal projection and branching pattern of two large-diameter dI2 neurons at the lumbar and brachial levels (Figure 3B and C). Numerous collaterals that penetrate the spinal cord along its entire length were evident. Importantly, VSCT dI2 neurons projected to spinal targets at the lumbar, thoracic, and brachial spinal levels and to the brain stem and cerebellum (Figure 3).

To measure the proportion of dI2 in VSCT neurons, we labeled VSCT axons with GFP and dI2 axons with Cherry (for experimental design, see Figure 3—figure supplement 1B and C). The number of axons expressing the reporters at the contralateral superior cerebellar peduncle was scored. 10% of the VSCT axons belonged to dI2 neurons (Figure 3—figure supplement 1D). Thus, the large-diameter dI2 neurons constitute 10% of the VSCT neurons, consistent with the anatomical observation that the VSCT comprises a heterogeneous population of INs (Jankowska and Hammar, 2013; Stecina et al., 2013).

Mapping the synaptic input and output of dI2 neurons

To obtain the connectome of dI2 neurons, we employed enhancer-mediated synaptic labeling of presynaptic neurons coupled with soma labeling of postsynaptic neurons. We used three criteria for assessing synaptic contact: (1) the likelihood of connectivity was examined by spatial overlap of axonal terminals from the presumed presynaptic neurons and the somata of the postsynaptic neurons; (2) synaptic boutons were detected on the somatodendritic membrane of postsynaptic neuron; and (3) colabeling was observed between the presynaptic reporter and synaptotagmin (syn). We used confocal imaging and 3D reconstitution to score overlap (Figure 4—figure supplement 1A, B).

As a proof of concept, we tested the colabeling of dI2::SV2-GFP and syn in dI2 to contralateral pre-MN synapses. Of 144 genetically labeled boutons, 121 (84%) were syn+. The syn- boutons were significantly smaller (Figure 4—figure supplement 1C). We set a volume threshold (0.07 µm3, Figure 4—figure supplement 1C), and small-volume SV2-GFP boutons were not considered synapses in the study. Using these criteria, we mapped the putative pre-dI2 and post-dI2 neurons.

dI2 neurons receive synaptic input from pre-MNs and sensory neurons

To assess the synaptic input to dI2 neurons, we investigated their synaptic connectivity with the following: (1) dorsal root ganglion (DRG) neurons (Figure 4A). (2) Ipsilateral pre-MNs. General ipsilateral pre-MNs were labeled by injecting a PRV-Cherry virus into the ipsilateral hindlimb musculature (Hadas et al., 2014; Figure 4B). Two genetically defined classes of pre-MNs were examined: dI1i excitatory INs (Figure 4C, Figure 4—figure supplement 2) and the V1 inhibitory pre-MN population (Bikoff et al., 2016; Gosgnach et al., 2006; Figure 4D). (3) Reticulospinal tract neurons (Figure 4E). dI2, DRG, V1, and dI1 neurons were labeled using specific enhancers (Figure 1—figure supplement 1A).

Figure 4 with 3 supplements see all
Synaptic inputs to dI2 neurons.

Schematic representations of the experimental design for labeling dI2::GFP or dI2::Cherry interneurons (INs; cyan) and potential sources of synaptic inputs (magenta). The soma densities of dI2 INs and the synaptic densities are illustrated in (A–E). The density values presented are 10–80%, 20–80%, 25–80%, 30–50%, and 20–80%, respectively. The laminar distributions are illustrated on the right side of (A–E). Examples of dI2 neurons contacted by axons or synaptic boutons are shown in (A′–D′), and their 3D reconstruction is shown in (A″–D″). Genetic labeling was achieved using specific enhancers (Figure 1—figure supplement 1A) introduced by electroporation at HH18. (A) Dorsal root ganglion (DRG) neurons form contacts on dI2 neurons. Inset in (A): cross-section of embryonic day (E) 17 embryos at the crural plexus level of the lumbar cord. A dorsally located dI2 neuron contacted by numerous sensory afferents, magnified in (A′) and 3D-reconstructed in (A″) (N = 18 sections, the scheme was constructed based on one representative embryo). (B) Premotor neurons (pre-MNs) form contacts on dI2 neurons. dI2 neurons were labeled at HH18. At E13, PRV virus was injected into the leg musculature, and the embryo was incubated until the infection of the pre-MNs (39 hr) (N = 34 sections, the scheme was constructed based on two representative embryos). (C) dI1 neurons form synapses on dI2 neurons. (N = 8568 synapses, 2 embryos). (C′) A representative SV2::Cherry synapse on dI2 dendrites positive for synaptotagmin. Demonstrated by horizontal and vertical optical sections in the Z-axis (see cursors and color channels). (D) V1 neurons form synapses on dI2 neurons (N = 1923 synapses, 2 embryos). (E) dI2 neurons are not contacted by 5-HT synaptic terminals (N = 1718 synapses, 1 embryo). E17 cross-sections of dI2::GFP-labeled embryos were stained for 5-HT. See Figure 4—source data 1.

Figure 4—source data 1

Localizations of pre-dI2 terminals and synapses at the sciatic level.

The X/Y coordinates of dorsal root ganglion (DRG) axons and premotor neurons (pre-MNs) axons (Figure 4A and B). The X/Y coordinates of dI1, V1, and synapses and 5HT terminals at the sciatic level (Figure 4C–E).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig4-data1-v2.xlsx

A density profile of the axons of DRG neurons (Figure 4A, Figure 4—figure supplement 3A–D) was aligned with the density plots of the dl2 somata. Overlap between the axonal terminals of DRG neurons was evident (Figure 4A, Figure 4—figure supplement 3C and D). Contact between DRG axons and dI2 neurons was mainly apparent in the dorsal dI2 neurons, while the ventral dI2 neurons received little to no input from DRG neurons (2.8 ± 2.4 vs. 16.9 ± 11.3 contacts per neuron for ventral and dorsal dI2 neurons, respectively; p<1e-5; Figure 4A, Figure 4—figure supplement 3E). In contrast, large- and small-diameter dI2 neurons did not exhibit a significant difference in DRG axon contacts (10.4 ± 14.9 vs. 7.8 ± 5.7 contacts per neuron for large and small dI2 neurons, respectively; p=0.4) (Figure 4—figure supplement 3F).

The density plot of PRV-labeled pre-MNs overlapped with the density plots of the dl2 somata (Figure 4B), and the axonal terminals of pre-MNs were visible on dI2 somata and dendrites (Figure 4B′,B′′), suggesting that pre-MNs contacted dI2 neurons. To solidify the evidence for pre-MN/dI2 connectivity, we used synaptic reporters expressed in genetically identified dI1 and V1 pre-MNs. Excitatory dI1 synapses and inhibitory V1 synapses overlapped with the density plots of the dl2 somata (Figure 4C and D). Synaptic connections, evaluated by boutons found on dI2 dendrites and somata, were apparent from V1 and dI1i (Figure 4B–D, Figure 4—figure supplement 3G).

Serotonergic neurons are the main reticulospinal input to VSCT in cat (Hammar et al., 2004; Hammar and Maxwell, 2002). Serotonergic synapses were concentrated on motor neurons and were not observed on dI2 neurons (Figure 4E, Figure 4—figure supplement 3H). Double labeling of 5-HT and dI2 neurons did not reveal any synaptic input. The lack of synaptic serotonergic input may be related to the difference in species or may suggest that other, non-dI2 VSCT neurons located adjacent to motoneurons are contacted by the reticulospinal neurons. The analysis of synaptic inputs supports the concept that dI2 neurons constitute part of the VSCT. These cells receive input from sensory afferents and inhibitory and excitatory pre-MNs and project to the cerebellum.

dI2 neurons innervate contralateral lumbar and brachial pre-MNs and dI2 neurons

Axon collaterals of dI2 invade the gray matter along the entire length of the spinal cord, as revealed by whole-mount staining of spinal cords electroporated with an alkaline phosphatase reporter (dI2::AP) (Figure 5A), cross-sections of dI2 neurons expressing membrane-tethered EGFP (Figure 5B), and light-sheet microscopy analysis (Figure 3—figure supplement 1; Video 1, Video 2, Video 3). The region innervated by dI2 collaterals (arrow in Figure 5B) overlaps with that of the V0 and V1 pre-MNs (Lai et al., 2016; Griener et al., 2015) as well as with that of the contralateral dI2 neurons (Figures 1 and 5B). To assess the potential spinal targets of dI2 neurons, we inspected the degree of overlap between dI2 synapses and dI2 somata (Figure 5C), ipsilateral pre-MNs (Figure 5D), and contralateral pre-MNs (Figure 5E). The alignment revealed an overlap of dI2 synapses with ipsilateral/contralateral pre-MNs and dI2 neurons (Figure 5C–E), supporting their potential connectivity. Labeling of dI2 synapses coupled with labeling of the above neuronal population showed dI2 synaptic boutons on pre-MNs and dI2 neurons at the lumbar level (Figure 5C–E, Figure 5—figure supplement 1A–C).

Figure 5 with 1 supplement see all
Spinal synaptic targets of dI2 neurons.

(A) Whole-mount staining of the spinal cord (thoracic segments) expressing alkaline phosphatase (AP) in dI2 neurons. The lumbar dI2 neurons (not included in the image) were labeled with AP. dI2 axon collaterals project and into the spinal cord (arrows). Abbreviations in the coordinates: rostral: R: caudal: C. (B) Cross-section of an embryonic day (E) 17 embryo at the crural plexus level of the lumbar spinal cord. Axonal collaterals (white arrow) penetrating the gray matter of the contralateral side are evident. Schematic representations of the experimental design for labeling synapses (dI2::SV2-GFP, yellow) and potential targets (magenta) supplemented by cell soma density and dI2 synaptic densities are illustrated in (C–F). The laminar distribution of the somata and synapses is illustrated on the right side of (C–F). Examples of target neurons contacting synaptic boutons of dl2 neurons are shown in (C′–F′), and their 3D reconstruction is shown in (C″–F″). Genetic labeling was achieved using dI2 enhancers (Figure 1—figure supplement 1A) electroporated at HH18. Premotor neurons (pre-MNs) were labeled by injection of PRV-Cherry into the hindlimbs (D, E) or the forelimb (F) musculature at E13. The embryos were incubated until the pre-MNs were infected (39 hr). (C) dI2 neurons innervate contralateral dI2 neurons (N = 4735 synapses and N = 374 cells, respectively, two embryos). (D) dI2 neurons innervate ipsilateral projections of pre-MNs at the sciatic plexus level (N = 4735 synapses and N = 936 cells, respectively, scheme was done based on one representative embryo). (E) dI2 neurons innervate contralaterally projecting pre-MNs at the sciatic plexus level (N = 4735 synapses and N = 47 cells, respectively, scheme was done based on one representative embryo). (F) dI2 neurons innervate ipsilaterally projecting pre-MNs at the brachial level (N = 2215 synapses and N = 286 cells, respectively, three embryos). See Figure 5—source data 1.

Figure 5—source data 1

Localization of dI2 synapses on post-dI2 neurons at the sciatic and brachial levels.

The X/Y coordinates of dI2 synapses, ipsi premotor neurons (pre-MNs), commissural pre-MNs, and contralateral dI2 at the sciatic level; dI2 synapses and ipsi pre-MNs at the brachial level (Figure 5).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig5-data1-v2.xlsx

The pattern of dI2 collaterals along the entire rostrocaudal axis (Figure 3A–C, Figure 5A) suggests that dI2 neurons innervate contralateral pre-MNs and dI2 neurons at multiple levels. To test this hypothesis, labeling of lumbar dI2 neurons was coupled with labeling of brachial pre-MNs and dI2 somata by injecting PRV into the wing musculature or electroporating a reporter into brachial dI2 neurons, respectively (Figure 5F, Figure 5—figure supplement 1D). dI2 synapses overlapped with the putative targets, and synaptic boutons originating from lumbar-level dI2 neurons were apparent on dI2 neurons and on the contralateral and ipsilateral pre-MNs of the wings (Figure 5F, Figure 5—figure supplement 1D and E).

Neuronal and synaptic labeling experiments showed that lumbar dI2 neurons innervate the cerebellum, lumbar and brachial pre-MNs, and contralateral dI2 neurons. Hence, dI2 neurons may relay peripheral and intraspinal information to the cerebellum and to the contralateral lumbar and brachial motor control centers.

Silencing of dI2 neurons impairs the stability of bipedal stepping

The synaptic input to dI2 neurons and their putative targets implicates them as relaying information about motor activity to the contralateral spinal cord and the cerebellum. Thus, we hypothesized that manipulation of their neuronal activity may affect the dynamic profile of stepping. To study the physiological role of dI2 neurons, we silenced their activity by expressing the tetanus toxin (TeTX) light chain gene, which blocks synaptic transmission (Yamamoto et al., 2003), in the bilateral lumbar dI2 neurons. EGFP was cotargeted in a 2/1 TeTX/EGFP ratio for post hoc analysis of the efficacy of electroporation (Supplementary file 1). Chicks expressing EGFP in dI2 neurons and chicks that did not undergo electroporation were used as controls. To maximize the number of targeted dI2 neurons, we combined genetic targeting with the Foxd3 enhancer and spatial placement of the electrodes at the dorsal lumbar spinal cord (Figure 1—figure supplement 1A). Embryos were electroporated at HH18. Upon hatching, chicks were trained for targeted overground locomotion. The gait parameters of four controls and 5 TeTX-treated chicks were measured while chicks were walking toward their imprinting trainer along a horizontal track (6–20 walking sessions, 5–8 strides each, per chick).

To test whether silencing of dI2 neurons impairs posthatching development and muscle strength, the chicks were weighed at P8, and their foot grip strength was evaluated on the same day. All chicks were of comparable weight (average – 144.7 ± 12.1 g; Supplementary file 1). As a functional measure of foot grip, we tested the ability of the chicks to maintain balance on a tilted mesh surface. TeTX-manipulated chicks and control chicks maintained balance on the tilted surface up to 63–70°, with no apparent statistically significant differences (Supplementary file 1). Thus, manipulation of dI2 neuronal activity did not impair the development or balance or muscle strength.

Analysis of overground locomotion in the control and TeTX-treated chicks revealed no significant differences in swing velocity or striding pattern. A 180° out-of-phase pattern was found during stepping in all the manipulated and control chicks (Figure 6—figure supplement 1A, Table 1). However, substantial differences were scored in stability parameters: TeTX chicks exhibited whole-body collapses during stepping (Figures 6B and C and 7A), a wide-base gait (Table 2), and variable limb movements (Figure 6A, D and E; Figure 7B and C; Figure 7—figure supplement 1; Table 3).

Figure 6 with 1 supplement see all
Kinematic analysis of locomotion in posthatching chicks following the silencing of dI2 neurons.

(A) Schematic illustration of chick hindlimb joints (bold) and bones (regular). The knee joint connects the femur and the tibiotarsus, and the ankle connects the tibiotarsus and the tarsometatarsus, which connects to the phalanges at the TMP joint. During the swing phase of birds, ankle flexion leads to foot elevation, while the knee is relatively stable. (B, C) Stick diagrams of stepping in a control chicken d2::GFP (B) and in a d2::TeTX chicken (C). Arrows indicate collapses, and overshoots are denoted by arrowheads. (D) Overlays of knee height (demonstrated in insert) trajectories during the swing phase in all analyzed steps of each of the control and TeTX-treated posthatching day (P) 8 hatchlings are shown superimposed with the respective 20–80% color-coded density plots as a function of the percentage of swing (see text and Materials and methods). Arrows indicate collapses, and overshoots are indicated by arrowheads. (E) The angular trajectories of the TMP joint (shown in insert) during the swing phase in all analyzed strides of each of the control and TeTX-treated P8 hatchlings are shown superimposed on the respective 20–80% color coded density plots as a function of the percentage of swing (see text and Materials and methods). See Figure 6—source data 1, Figure 6—source data 2, Figure 6—source data 3.

Figure 6—source data 1

Analysis of knee height trajectories during the swing phase.

The knee height trajectories during the normalized swing in all the analyzed steps of all chicks (Figure 6D).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig6-data1-v2.xlsx
Figure 6—source data 2

Analysis of TMP angles during the swing phase.

The TMP angle trajectories during the normalized swing in all the analyzed steps of all chicks (Figure 6E).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig6-data2-v2.xlsx
Figure 6—source data 3

Statistical analysis of knee height trajectories and TMP angles.

Statistical analysis for the data presented in Figure 6.

https://cdn.elifesciences.org/articles/62001/elife-62001-fig6-data3-v2.docx
Figure 7 with 2 supplements see all
Parameters of reduced stability in bipedal stepping in TeTX-treated chicks.

(A) The percentage of steps with body collapses in the controls and TeTX-manipulated hatchlings (n = 4 and n = 5, respectively). p-value<0.0001 (Z-test). See Table 3 for the proportions of falls at the individual chick level. (B) Analysis of the mean range of knee height changes during the swing phase of control and TeTX-treated chicks (n = 4 and n = 5, respectively). p-value<0.0001 using a t-test allowing different variances. See Figure 7—figure supplement 1A and Table 3 for individual chick data and statistical analysis details. (C) Analysis of the mean range of TMP angular excursions during the swing phase of control- and TeTX-treated chicks (n = 4 and n = 5, respectively). p-value<0.0001, Watson–Williams test. See Figure 7—figure supplement 1B and Table 3 for individual chick data and statistical analysis details. (D, E) Schematic illustrations showing the connectome of lumbar dI2 neurons. The synaptic inputs (D) and outputs (E) of dI2 neurons are illustrated. dI2 neurons (magenta) receive synaptic input from sensory afferents (solid blue line indicates massive synaptic input, and dashed blue line indicates sparse innervation), from inhibitory and excitatory premotor neurons (pre-MNs; yellow), and from the contralateral lumbar dI2 neurons. dI2 neurons innervate the contralateral lumbar and brachial pre-MNs (both commissural and ipsilaterally projecting pre-MNs are innervated by dI2 cells), the lumbar and brachial contralateral dI2 cells, lumbar precerebellar neurons (green), and the cerebellar granule cells. See Figure 7—source data 1.

Figure 7—source data 1

Analysis of collapses.

The number of collapses of each chick (Figure 7A).

https://cdn.elifesciences.org/articles/62001/elife-62001-fig7-data1-v2.xlsx
Table 1
Stride velocity and left-right phase in control and tetanus toxin (TeTX-manipulated chicks).
ChickMean swing velocity (cm/s)Mean left-right phase (°)# of steps
TeTX146.78 ± 22.13184.679 ± 33.003113
TeTX262.24 ± 20.17182.293 ± 32.0163
TeTX348.06 ± 20.04180.784 ± 31.06469
TeTX457.24 ± 24.35180.502 ± 36.29159
TeTX536.66 ± 17.61181.97 ± 35.78793
Control 1 (GFP)79.65 ± 37.77182.369 ± 35.36647
Control 2 (GFP)41.91 ± 20.41182.384 ± 26.70819
Control 3 (not electroporated)41.09 ± 16.59N.D.121
Control 4 (not electroporated)42.3 ± 30.91N.D.51
Table 2
Maximum stride width in control and tetanus toxin (TeTX)-manipulated chicks.
ChickMaximum stride width (cm)# of steps
TeTX15.11 ± 1.8997
TeTX25.32 ± 1.3836
TeTX34.5 ± 1.0127
TeTX44.9 ± 1.1649
TeTX55.82 ± 1.71110
Control 84.15 ± 1.07137
Control 94.32 ± 1.32115
Table 3
Collapses, knee height, and TMP angle ranges in control and tetanus toxin (TeTX)-manipulated chicks.
Chick% of steps with collapseTMP angle: mean range (°)# of steps% of steps with collapse
All steps, meanCombined meanMinus collapses, meanMinus collapses, combined meanAll steps, meanCombined meanMinus collapsesMinus collapses, combined mean
TeTX14.43.05 ± 0.493.11 ± 0.742.87 ± 0.382.83 ± 0.5782.27 ± 2272.71 ± 20.5279.4 ± 24.568.97 ± 21.47113
TeTX220.603.5 ± 0.33.35 ± 0.4771.79 ± 2571.1 ± 25.6863
TeTX318.82.57 ± 0.312.27 ± 0.2664.17 ± 2162.79 ± 20.969
TeTX420.452.54 ± 0.552.477 ± 0.5672.48 ± 1766.22 ± 19.159
TeTX5293.86 ± 0.833.2 ± 0.272.86 ± 1265.85 ± 13.1393
Control 1 (GFP)2.121.91 ± 0.221.98 ± 0.331.91 ± 0.221.98 ± 0.3356.8 ± 16.449.34 ± 16.0356.95 ± 16.649.4 ± 16.1847
Control 2 (GFP)01.83 ± 0.251.83 ± 0.2541.42 ± 18.441.42 ± 18.419
Control 3 (not electroporated)02.42 ± 0.232.42 ± 0.2354.86 ± 9.6554.86 ± 9.65121
Control 4 (not electroporated)01.75 ± 0.11.75 ± 0.144.12 ± 12.5144.12 ± 12.5151

Whole-body collapses

A collapse was scored as a decline in knee height below 85% of the average knee height at the stance phase of the step (arrow in Figure 6C). We measured the number of collapses in 50–190 steps. In control chicks, collapses occurred in 0.53% ± 0.92% of the steps. In TeTX-manipulated chicks, we observed collapses in 19.46% ± 8.3% of the steps, which was significantly different from the rate in controls (Figure 7A). As some collapses were followed by an overextension (‘overshoot’ in leg elevation), also manifested in the profile of the knee height trajectory during the swing phase (Figure 6D), we further studied the relationship between the two phenomena. In general, there was high variability between chicks in this aspect.

Most collapses (64.9% ± 19.7%) were not preceded or followed by overextension. About 22% of the collapses (22.47% ± 21.5% of collapses, e.g., arrowhead in Figure 6C) were followed by overextension, suggesting a postcollapse compensation in the extensor drive. The rest of the collapses (12.63% ± 7.56%) were preceded by overextension.

Wide-base stepping

A wide-base stance is typical of an unbalanced ataxic gait. The stride width was measured between the two feet during the double stance phase of stepping. The mean stride in TeTX-manipulated chicks 1, 2, 4, and 5 was significantly wider than that in the control chicks, while the width in TeTX3 was similar to that in the controls (Table 2).

Variable limb movements

In stable gait, limb trajectories are consistent from stride to stride. Therefore, we compared the trajectories of knee height and angle of the TMP joint during the swing phase of stepping between control and TeTX-manipulated chicks. Plots of the knee height and TMP angle trajectories during the normalized swing in all the analyzed steps of each chick are shown superimposed in Figure 6D and E, respectively. These data demonstrate that the range of changes in TeTX-manipulated chicks was higher than that in control chicks.

Further analyses revealed that, overall, the control group showed lower knee height and TMP angle ranges than the TeTX-treated group, even though there were differences within groups (Figure 7—figure supplement 1). The average knee height range of the combined control chicks (1.981 ± 0.33) was significantly lower than the range of the combined TeTX-treated chicks (3.109 ± 0.74) (Figure 7B). A similar comparison of the combined ranges of angular excursions of the TMP joint during the normalized swing revealed that the average angle of the control group (49.34 ± 16.03) was significantly lower than the average of the TeTX-treated chicks (77 ± 22.15; Figure 7C).

Since the increased range of changes could be due to the effects of the substantial increase in body collapses during stepping (Figure 7A, see also Figure 6), we excluded steps featuring whole-body collapses and reanalyzed the data. The data summarized in Table 3 show that the significant difference between controls and the TeTX-treated chicks in the range of the knee height and the TMP angle excursions was maintained. Thus, the increase in irregularity in the TeTX-treated chicks is not caused exclusively by the body collapses of the TeTX-treated chicks.

Loss of balance can also arise from slipping, which can originate from a shallow landing angle (Clark and Higham, 2011). The landing angle was characterized as the angle between the ground and the imaginary line connecting the knee joint (which is located near the chicken’s center of mass; Daley and Biewener, 2006) to the TMP joint at the end of the swing phase (Figure 7—figure supplement 2A). Thus, we analyzed the landing angles of the manipulated and control chickens. No significant differences in landing angles were detected (Figure 7—figure supplement 2B). Additionally, we compared the landing angle before a collapse to landing angles not preceding a collapse. We found that the angles preceding a collapse were not smaller and even tended to be slightly larger than the landing angles that did not precede a collapse (Figure 7—figure supplement 2C, p=0.02). These results argue against the possibility that the increased frequency of collapses in the manipulated chickens stemmed from slipping and sliding events.

Overall, the kinematic parameters of the dI2-TeTX-treated chicks demonstrate a reduction in stability during locomotion, indicating a possible role of dI2 in the stabilization of bipedal stepping.

Discussion

The VSCT is thought to provide peripheral and intrinsic spinal information to the cerebellum to shape and update the output of spinal networks that execute motor behavior. The lack of genetic access to VSCT neurons hampers efforts to elucidate their role in locomotion. Using a genetic toolbox to dissect the circuitry and manipulate neuronal activity in the chick spinal cord, we studied spinal INs with VSCT characteristics. The main finding in our study is that dI2 neurons in the chick lumbar spinal cord are commissural neurons that innervate pre-MNs at the contralateral lumbar and brachial spinal levels and granule neurons in the ipsilateral cerebellum. Hence, a subpopulation of dI2 neurons form part of the avian VSCT. Targeted silencing of dI2 neurons leads to impaired stepping in P8 hatchlings. We described the spatial distribution of subpopulations of dI2 neurons, deciphered their connectomes, mapped the trajectory of their projections to the cerebellum, and suggested possible mechanisms for the gait perturbation resulting from their genetic silencing, as discussed below.

The connectome of dI2 neurons

Using the intersection between genetic drivers and spatially restricted delivery of reporters to define lumbar and brachial neurons, we identified several targets of dI2 lumbar neurons. Lumbar dI2 neurons innervate contralateral lumbar dI2 neurons as well as commissural and noncommissural lumbar pre-MNs. This connectivity may influence the bilateral spinal output circuitry at the lumbar cord (e.g., Bras et al., 1988; Jankowska and Hammar, 2013). Moreover, the ascending axons of lumbar dI2 neurons give off gray matter collaterals innervating contralateral dI2 neurons and commissural and noncommissural pre-MNs throughout the brachial spinal cord (Figure 7D and E). Therefore, lumbar dI2 neurons may also contribute to the inter-enlargement coupling described between the segments of the spinal cord that move the legs and the wings (e.g., Valenzuela et al., 1990; Ruder et al., 2016 for forelimb to hindlimb coupling connectivity in mice).

We demonstrated that lumbar dI2 cells receive sensory innervation, premotor inhibitory and excitatory innervation, and innervation from contralateral lumbar dI2 cells (Figure 7D and E). Thus, lumbar dI2 neurons can provide the cerebellum and the contralateral pre-MNs with proprioceptive information, copies of motor commands delivered from the ipsilateral pre-MNs, and integrated information from contralateral dI2 neurons (Figure 7D and E).

Our wiring-decoding studies are based on the availability of enhancer elements that direct expression in specific spinal INs. The lack of enhancer elements for known pre-MNs, such as V2, precluded their analysis. Future experiments using identified regulatory elements that direct expression in two other pre-MN populations, the dI3 and V0 neurons (Avraham et al., 2010b; Gard et al., 2017), will reveal the extent of premotor information relayed by dI2 neurons.

dI2 subpopulations

Our study reveals two anatomically distinct subpopulations of dI2 neurons: precerebellar projection dI2 cells, which also innervate spinal targets along the entire extent of the spinal cord, and propriospinal dI2 cells, which innervate targets within the lumbar level. The laminar and medial/lateral positions of the two dI2 populations are similar, and we did not find subtype-specific expression of the known dI2 TFs. Contrary to our findings, a recent study (Osseward et al., 2021) has shown that tract and propriospinal INs in the mouse spinal cord differ in the localization of their somata along the mediolateral axis and their transcription of TFs. Tract neurons reside in the lateral spinal cord and express group-N TFs, while propriospinal neurons settle at the medial spinal cord and express group-Z TFs. Several reasons may explain the discrepancy between our results and those of Osseward et al. The analysis in mice (Osseward et al., 2021) was applied to numerous cardinal populations of neurons but excluded dI2 neurons. Hence, dI2 may represent an exception. In addition, our study revealed that precerebellar dI2 neurons share two wiring patterns: tract and propriospinal neurons. Applying single-cell RNA sequencing to dI2 neurons will reveal whether short- and long-range targeting in dI2 neurons are characterized by distinct transcriptomes or by a shared N + Z transcriptome.

Physiological role of dI2 neurons

The unclear genetic origin of physiologically equivalent lumbar VSCT neurons has prevented a better understanding of their role in hindlimb locomotion. Our wiring and neuronal-silencing studies implicated dI2 as a significant contributor to the regularity and stability of locomotion in P8 hatchlings. The kinematic analysis of TeTX-treated hatchlings revealed imbalanced locomotion with occasional collapses, increased stride variability, a wide-base gait, and variable limb movements during stepping.

The mechanisms accounting for impaired stepping following dI2 neuron silencing are still unknown. One of the possible mechanisms is that silencing dI2 neurons perturbs the delivery of peripheral and intrinsic feedback to the cerebellum, leading to unreliable updating of the motor output produced by the locomotor networks, thereby impairing bipedal stepping. Another possible mechanism is based on the similarity of the gait instabilities of TeTX-treated hatchlings to ataxic motor disorders. Mammalian VSCT neurons receive descending input from reticulospinal, rubrospinal, and vestibulospinal pathways (Bras et al., 1988; Jankowska and Hammar, 2013). Neurons from the lateral vestibular nucleus have been reported to innervate extensor motoneurons at the lumbar level, as well as INs residing at medial lamina VII (Murray et al., 2018), the location where dI2 neurons were found to reside in our study. Thus, the vestibulospinal tract may convey input directly to the ipsilateral motor neurons and indirectly to contralateral motor neurons through dI2 neurons that innervate contralateral pre-MNs.

The local spinal connections between dI2 neurons and the contralateral pre-MNs and contralateral dI2 neurons may serve an important component of coordinated limb movements. dI2 synapses were found on both ipsilaterally and contralaterally projecting pre-MNs, both within their segmental level and at the brachial level, which regulates the movement of the wings. Thus, dI2 neurons may affect the motor output of the contralateral and ipsilateral sides of the cord by contacting commissural pre-MNs. Specific targeting of dI2 subpopulations – the precerebellar versus the propriospinal dI2 cells – is necessary to determine the relative contribution of dI2 subpopulations to the impaired stepping phenotype. However, there is no available genetic technique for differentially targeting the two subpopulations. In addition, the fact that the precerebellar dI2 neurons also innervate the lumbar spinal cord precludes the use of retrogradely target-derived neuronal activity modifiers.

In summary, our mapping studies of dI2 neurons and their connectomes, followed by characterization of the effects of their silencing on bipedal stepping, offer new insights into the function of dI2 neurons in vertebrates. We suggest that lumbar dI2 neurons not only relay sensory and intrinsic spinal network information to the cerebellum but also act as active mediators of motor functions at the lumbar segments and at the wing-controlling brachial segments of the spinal cord. Further circuit-deciphering studies of the constituents of subpopulations of dI2 cells, their targets, and their descending inputs are required to extend our understanding of the function of dI2 subpopulations in motor control.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Chicken)Gallus gallusGil-Guy Farm, IsraelNCBI Taxon: 9031
Strain, strain background (Pseudorabies virus)PRV152Enquist and Card, 2003NCBI Taxon: 10345
Strain, strain background (Pseudorabies virus)PRV614Enquist and Card, 2003NCBI Taxon: 10345
AntibodyRabbit anti-GFP(polyclonal)Molecular Probes, Eugene, Oregon, USAA-11122RRID:AB_221569Dilution(1:1000)
AntibodyMouse anti-GFP(monoclonal)AbcamAb1218AB_298911Dilution(1:100)
AntibodyGoat anti-GFP(polyclonal)AbcamAb6673RRID:AB_305643Dilution(1:300)
AntibodyRabbit anti-RFP(polyclonal)AcrisAP09229PU-NRRID:AB_2035909Dilution(1:1000)
AntibodyGoat anti-ChAT(polyclonal)Millipore, USAAB144PRRID:AB_2079751Dilution(1:300)
AntibodyMouse anti-synaptotagmin(monoclonal)Hybridoma Bank, University of Iowa, Iowa City, USAASV30RRID:AB_2295002Dilution(1:100)
AntibodyMouse anti-lhx1/5(monoclonal)Hybridoma Bank, University of Iowa, Iowa City, USA4F2RRID: AB_531784Dilution(1:100)
AntibodyMouse anti-FoxP4(monoclonal)Hybridoma Bank, University of Iowa, Iowa City, USAPCRP-FOXP4-1G7RRID:AB_2618641Dilution(1:50)
AntibodyRabbit anti-Pax2(polyclonal)Abcamab79389RRID:AB_1603338Dilution(1:50)
AntibodyChicken anti- lacZ(polyclonal)Abcamab79389RRID:AB_307210Dilution(1:300)
AntibodyRabbit anti-calbindin(polyclonal)SwantD-28kRRID:AB_2314070Dilution(1:200)
AntibodyGoat anti-FoxP2(polyclonal)Abcamab1307RRID:AB_1268914Dilution(1:1000)
AntibodyRabbit anti-5-HT(polyclonal)Abcamab140495Dilution(1:100)
Recombinant DNA reagentEdI1::CreAvraham et al., 2009N/A
Recombinant DNA reagentNgn1::CreAvraham et al., 2009N/A
Recombinant DNA reagentNgn1::FLPoHadas et al., 2014N/A
Recombinant DNA reagentFoxd3::FLPoHadas et al., 2014N/A
Recombinant DNA reagentFoxd3::CreAvraham et al., 2009N/A
Recombinant DNA reagentIsl1::CreAvraham et al., 2010aN/A
Recombinant DNA reagentCAG-LSL-GFPHadas et al., 2014N/A
Recombinant DNA reagentCAG-LSL-SV2-GFPHadas et al., 2014N/A
Recombinant DNA reagentCAG-FSF-LSL-GFPHadas et al., 2014N/A
Recombinant DNA reagentCAG-FSF-LSL-SV2-GFPThis paperN/AFigure 1—figure supplement 1;can be obtained from the Klar lab
Recombinant DNA reagentCAG-FSF-LSL-cherryThis paperN/AFigure 1—figure supplement 1;can be obtained from the Klar lab
Recombinant DNA reagentCAG-FSF-LSL-SV2-cherryThis paperN/AFigure 1—figure supplement 1;can be obtained from the Klar lab
Recombinant DNA reagentCAG-FSF-LSL-APThis paperN/AFigure 1—figure supplement 1;can be obtained from the Klar lab
Recombinant DNA reagentCAG-LSL-TeXTThis paperN/AFigure 1—figure supplement 1;can be obtained from the Klar lab
Recombinant DNA reagentCAG-LSL-F_SV2-cherry_F-GFPThis paperN/AFigure 1—figure supplement 1;can be obtained from the Klar lab
Recombinant DNA reagentpGEMTEZ-TeTxLCAddgene#32640
Sequence-based reagentFoxd3-FThis paperPCR primersTCATCACCATGGCCATCCTG
Sequence-based reagentFoxd3-RThis paperPCR primersGCTGGGCTCGGATTTCACGAT
Sequence-based reagentvGlut2-FThis paperPCR primersGGAAGATGGGAAGCCCATGG
Sequence-based reagentvGlut2-RThis paperPCR primersGAAGTCGGCAATTTGTCCCC
Sequence-based reagentVIAAT-FThis paperPCR primersCTGAACGTCACCAACGCCATCC
Sequence-based reagentVIAAT-RThis paperPCR primersGGGTAGGAGAGCAAGGCTTTG
Commercial assay or kitNucleoBond Xtra MidiMacherey-NagelCat # 740410.50
Chemical compound, drugCTB conjugated to Alexa Fluor 647Thermo FisherC347780.3 M
Software, algorithmJMPJMPhttps://www.jmp.com/en_gb/home.html
Software, algorithmAdobe PhotoshopAdobehttps://www.adobe.com/il_en/
Software, algorithmImageJImageJhttps://imagej.nih.gov/ij/
Software, algorithmIMARISOxford Instrumentshttps://imaris.oxinst.com/
Software, algorithmMacVectorMacVectorhttps://macvector.com/index.html
Other(electroporator)BTX ElectroporatorBTX Harvard ApparatusCat#45-0662
Other(confocal microscope)FV1000; OlympusOlympushttps://www.olympus-global.com/
Other(microscope)Eclipse NiNikonhttps://www.nikon.com/
Other(light-sheet microscope)LaVision Ultramicroscope II light-sheet microscopeLaVision BioTechttps://www.lavisionbiotec.com/

Animals

Fertilized white leghorn chicken eggs (Gil-Guy Farm, Israel) were incubated under standard conditions at 38°C. All experiments involving animals followed the designated policies of the Experiments in Animals Ethics Committee and were performed with its approval.

3D reconstruction and density plot analysis

Request a detailed protocol

The programs for both 3D reconstruction and the density plot analysis were written in MATLAB. The density plots were generated based on cross-sectional images converted to a standard form. The background was subtracted, and the cells were filtered automatically based on their soma size or using a manual approach. Subsequently, two-dimensional kernel density estimation was obtained using the MATLAB function ‘kde2d.’ Finally, unless indicated otherwise, a contour plot was drawn for density values between 20% and 80% of the estimated density range, with six contour lines.

In ovo electroporation

Request a detailed protocol

A DNA solution of 5 mg/mL was injected into the lumen of the neural tube at HH stage 17–18 (E2.75–E3). Electroporation was performed using 3 × 50 ms pulses at 25–30 V, applied across the embryo using a 0.5 mm tungsten wire and a BTX electroporator (ECM 830). Following electroporation, 150–300 μL of antibiotic solution containing 100 unit/mL penicillin in Hanks’ balanced salt solution (Biological Industry, Beit-Haemek) was added on top of the embryos. Embryos were incubated for 3–19 days prior to further treatment or analysis.

Immunohistochemistry and in situ hybridization

Request a detailed protocol

Embryos were fixed overnight at 4°C in 4% paraformaldehyde/0.1 M phosphate buffer, washed twice with phosphate buffered saline (PBS), incubated in 30% sucrose/PBS for 24 hr, and embedded in optimal cutting temperature (OCT) compound (Scigen, Grandad, USA). Sections with a thickness of 20 μm were cut on a cryostat. These sections were collected on Superfrost Plus slides and kept at −20°C. For 100 μm sections, spinal cords were isolated from the fixed embryos and subsequently embedded in warm 5% agar (in PBS), and 100 μm sections (E12–E17) were cut with a vibratome. Sections were collected in wells (free-floating technique) and processed for immunolabeling.

The following primary antibodies were used: rabbit polyclonal GFP antibody 1:1000 (Molecular Probes, Eugene, OR, USA), mouse anti-GFP 1:100, goat anti-GFP 1:300 (Abcam), rabbit anti-RFP 1:1000 (Acris), goat anti-ChAT antibody 1:300 (Cemicon, Temecula, CA, USA), mouse anti-synaptotagmin antibody 1:100 (ASV30), mouse anti-Lhx1/5 1:100 (4F2), mouse anti-FoxP4 1:50 (hybridoma bank, University of Iowa, Iowa City, USA), mouse anti-Brn3a 1:50 (Mercury), rabbit anti-Pax2 antibody 1:50 (Abcam), chicken anti-lacZ antibody 1:300 (Abcam), rabbit anti-Calbindin 1:200 (Swant), rabbit anti-VGLUT2 antibody (Synaptic Systems, Göttingen, Germany), goat anti-FoxP2:1000 (Abcam), anti-FoxP1:100 (ABR Synaptic), and rabbit anti-5-HT (Abcam). The following secondary antibodies were used: Alexa Fluor 488/647-conjugated AffiniPure donkey anti-mouse, anti-rabbit, and anti-goat (Jackson) and Rhodamine Red-X-conjugated donkey anti-mouse and anti-rabbit (Jackson). Images were taken under a microscope (Eclipse Ni; Nikon) with a digital camera (Zyla sCMOS; Andor) or captured using the integrated camera of a confocal microscope (FV1000; Olympus).

In situ hybridization was performed as previously described (Avraham et al., 2010a). The following probes were employed: Foxd3, vGlut2, and VIAAT probes were amplified from the cDNA of E6 chick embryos using the following primers. Foxd3: forward TCATCACCATGGCCATCCTG, reverse GCTGGGCTCGGATTTCACGAT. vGlut2: forward GGAAGATGGGAAGCCCATGG, and reverse GAAGTCGGCAATTTGTCCCC. VIAAT: forward CTGAACGTCACCAACGCCATCC, reverse GGGTAGGAGAGCAAGGCTTTG. The T7 RNA polymerase cis-binding sequence was added to the reverse primers.

Laminar division

Request a detailed protocol

The standard forms of the spinal cord (for the crural, sciatic, and brachial plexus levels) were computationally divided into polygons for the different laminae (Martin, 1979). The number of neurons or synapses inside each lamina border was scored using their coordinates.

Light-sheet microscopy dI2::mCherry was electroporated into the embryos at HH stage 17–18. Embryos were removed at E13, and the spinal cord and cerebellum were isolated. The tissue was cleared using the iDISCO technique as described (Renier et al., 2014). The mCherry-expressing neurons were stained by application of an anti-RFP antibody followed by Rhodamine Red-X-conjugated donkey secondary antibody. Each staining step included 3 days of incubation with the antibody and subsequent washing for 2 days. Then, the cleared tissue was divided into three segments: a lumbar spinal segment, a brachial spinal segment, and a segment including the brainstem and cerebellum. Each sample was placed in a quartz imaging chamber (LaVision BioTec) and scanned by a LaVision Ultramicroscope II light-sheet microscope operated by ImspectorPro software (LaVision BioTec). An Andor Neo sCMOS camera was used for 16-bit image acquisition. The imaging was performed at 2× magnification with a 0.5–1 µm step size and a green excitation filter (peak – 525 nm/width − 50 nm). Then, for 3D reconstruction and analysis of the samples, the resulting image z-stacks were converted to IMS format using Imaris File Converter (version 9.5). Because of the sample size, several z-stacks were required for full acquisition of each sample; they were stitched together into one z-stack by Imaris Stitcher. Then, the files were uploaded to Imaris (9.6 version) for advanced visualization and analysis. dI2 axons were tracked using the filament tracer feature in semiautomatic mode. After tracking, Imaris was used to generate videos and snapshots describing different features of the analyzed samples. Finally, text, arrows, and other symbols were added using Adobe AfterEffects software.

Synaptic marker validation

Request a detailed protocol

Validation of SV2-GFP reporter specificity was performed by using Imaris software. High-resolution confocal images of spinal cord sections, with clear SV2-GFP reporter expression and synaptotagmin (syn) immunolabeling, were used to quantify the degree of overlap of GFP+ terminals and syn+-labeled boutons. Both signals were three-dimensionally reconstructed, and we used the automatic quantification abilities of Imaris, further validated by additional manual counting, to quantify the number of GFP+ presynaptic terminals containing at least one syn+ bouton. In addition, the volume of GFP+ presynaptic terminals was documented to explore a possible dependence between terminal volume and syn+ bouton containment.

AP staining

Request a detailed protocol

The treated embryos were fixed with 4% paraformaldehyde–PBS for 24 hr at 4°C and washed twice with PBS for 30 min at 4°C. The fixed embryos were incubated at 65°C in PBS for 8–16 hr to inactivate endogenous AP activity. The treated embryos were washed with 100 mM Tris–Cl (pH 9.5) containing 100 mM NaCl and 50 mM MgCl2, and the residual placental alkaline phosphatase activity was visualized by incubating the embryos with NBT/BCIP (Roche) in the same buffer at 4°C for 24 hr. After extensively washing the embryos with PBS–5 mM EDTA, the spinal cord was imaged.

PRV infection and CTB retrograde labeling

Request a detailed protocol

From the attenuated PRV Bartha strain, we used two isogenic recombinants that express enhanced GFP (PRV152) and monomeric red fluorescent protein (PRV614). The viruses were harvested from Vero cell cultures at titers of 4 × 108, 7 × 108, and 1 × 109 plaque-forming units (PFU/mL). Viral stocks were stored at −80°C. Injections of 3 μL of PRV152 or PRV614 were made into the thigh, pectoralis, or distal wing musculature of E13 or E14 chick embryos using a Hamilton syringe (Hamilton; Reno, NV, USA) equipped with a 33-gauge needle. The embryos were incubated for 36–40 hr and sacrificed for analysis. For spinocerebellar projecting neuron labeling, we used a replication-defective HSV (TK-) that contains a lacZ reporter. The virus was injected into the cerebellum of E12–15 embryos in ovo, and the embryos were incubated for another 40–48 hr. Alternatively, CTB conjugated to Alexa Fluor 647 (Thermo Fisher) was injected into the cerebellum of E12–15 embryos together with the virus for visualization of both precerebellular neurons and the upstream neurons.

Force test

Request a detailed protocol

Muscle strength was evaluated using the measurement of the slope at which the chicks fell from a mesh surface as it was gradually tilted up from the horizontal. This test was repeated for each chick at least three times, and the average falling angle was calculated.

Analysis of left-right phase

Request a detailed protocol

Stride duration was measured as the time from right toe-off/foot-off to the next right toe-off (as a complete stride cycle for the right leg), and the ‘half-cycle’ duration was measured as the time from right toe-off to the time of left toe-off. The following formula was used to calculate the phase: ((LeftToeOff_1 - RightToeOff_1)/(RightToeOff_2 – RightToeOff_1))*360.

Behavioral tests and analysis

Request a detailed protocol

The embryos were bilaterally electroporated and then allowed to develop and hatch in a properly humidified and heated incubator. Afterwards, within 32 hr after hatching, the hatchling chicks were imprinted on the trainer. The P8 chicks were filmed in slow motion (240 fps) while freely walking (side and top views). The following parameters were scored: (1) weight; (2) foot grip strength; and (3) kinematic parameters during overground locomotion: (a) swing velocity, (b) swing and stance duration, (c) phase of footfalls, (d) heights of the knee and TMP joints, (e) angles of the TMP and ankle joints, (f) stride width (distance between feet during the double stance phase), and (g) landing angle.

Using semiautomated MATLAB-based tracking software (Hedrick, 2008), several points of interest were encoded. The leg joints as well as the eye and the tail were tracked. The position of these reference points was used for computational analysis using in-house MATLAB code for calculating different basic locomotion parameters (e.g., stick diagrams, velocity, joint trajectory, angles, range, and elevation), step patterns, and degrees of similarity. The landing angle was calculated as the angle between the imaginary line connecting the knee and the TMP joints and the ground, at the end of the swing phase. Dunnett’s test (Dunnett, 1955) was used to perform multiple comparisons of group means following one-way ANOVA. Circular statistics were used for analyses of angular data utilizing Oriana (KCS, version 4).

Data availability

All data generated or analysed during this study are included in the manuscript and the supporting files.

References

    1. Martin AH
    (1979)
    A cytoarchitectonic scheme for the spinal cord of the domestic fowl, Gallus gallus domesticus: Lumbar region
    Acta Morphologica Neerlando-Scandinavica 17:105–117.
    1. Yamamoto M
    2. Wada N
    3. Kitabatake Y
    4. Watanabe D
    5. Anzai M
    6. Yokoyama M
    7. Teranishi Y
    8. Nakanishi S
    (2003)
    Reversible suppression of glutamatergic neurotransmission of cerebellar granule cells in vivo by genetically manipulated expression of tetanus neurotoxin light chain
    The Journal of Neuroscience 23:6759–6767.

Decision letter

  1. Muriel Thoby-Brisson
    Reviewing Editor; CNRS Université de Bordeaux, France
  2. Ronald L Calabrese
    Senior Editor; Emory University, United States
  3. David SK Magnuson
    Reviewer; University of Louisville, United States
  4. Julien Bouvier
    Reviewer; Université Paris-Saclay, CNRS, France

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

The study by Haimson and colleagues investigates, in the chick model, the role of different sub-classes of developmentally-characterized dI2 interneurons in coordinating activity across different spinal regions, regulating stepping and reporting back activity to the brain. The work brings new insights in the so far under-investigated spinal dorsally-born neuronal types and about long-range projecting neurons that link the spinal cord with higher integrative centers.

Decision letter after peer review:

Thank you for submitting your article "Spinal dI2 interneurons regulate the stability of bipedal stepping" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Ronald Calabrese as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: David SK Magnuson (Reviewer #1); Julien Bouvier (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, when editors judge that a submitted work as a whole belongs in eLife but that some conclusions require a modest amount of additional new data, as they do with your paper, we are asking that the manuscript be revised to either limit claims to those supported by data in hand, or to explicitly state that the relevant conclusions require additional supporting data.

Our expectation is that the authors will eventually carry out the additional experiments and report on how they affect the relevant conclusions either in a preprint on bioRxiv or medRxiv, or if appropriate, as a Research Advance in eLife, either of which would be linked to the original paper.

Summary:

The three reviewers and I have carefully read your manuscript and we all have found a great interest in your results. However several concerns will have to be addressed in order to consider the paper further. In a first instance the paper requires careful revisions in the writing/editing, quantifications and clarity of the figures. Also the discussion on the main findings and their limitations should be revised in order to take into account reviewers' comments.

More specifically, the authors are asked to be more accurate/careful in describing:

1) The motor phenotype,

2) The identity of which sub-population of neurons is considered to be responsible for the locomotor behavior (di2 neurons vs vSCT neurons),

3) The anatomical data of the neurons of interest, possibly including their neurotransmitter phenotype,

4) The quantification and characterization of the synaptic contacts and their potential functional role. To address some of these issues additional experiments might be required, as more-specific silencing studies for example, but this might not be always necessary if the text is changed accordingly and if potential alternative explanations are given whenever possible.

Overall, we agreed that the paper has some important contributions to make if appropriately modified. Its suitability for publication will be re-evaluated after revisions.Reviewer #1:

This is a well-put-together manuscript describing carefully performed circuitry dissection and functional analysis of dl2 neurons in the chick. A genetic toolbox is used taking advantage of the electroporation technique applied to the embryos. The findings include a fairly convincing connectome for dl2 neurons and a functional phenotype that is, unfortunately, rather unsatisfying. The investigators conclude that dl2 interneurons regulate "stability" of bipedal stepping in the chick, which is fine, but the analysis misses an opportunity to more fully explore what the instability involves and thus to perhaps shed more light on the likely roles of this neuron population. The concerns/issues 3 and 4 below focus on this issue and the need for additional careful analysis of the behavior that will allow the phenotype to be more precisely described or ascribed to some aspect of stepping that might guide future studies in other models. For example, can the link between partial collapse and over-extensions be made more solid and thus argue that reduced extensor gain might be what results in the instability? What other analysis could be performed using the existing data/video to better describe the behavioral phenotype?

Concerns/issues:

1. The connectome part of the work appears solid and supports the concept that a sub population of the population are likely VSCT neurons, that the non VSCT neurons receive the bulk of the afferent input and that these neurons project to contralateral dl2 neurons (some which may be VSCT) and other premotor neurons. Anatomically, the only concern is that no distinctions were made between the lumbar and brachial populations, and if differences in these populations exist, it would be important and interesting to describe them.

2. Figure 2 Characterization of dl2/VSCT neurons as being primarily large dl2 neurons is quite convincing, and the observation that the dl2 neurons account for 10% of the VSCT axons is also of interest and quite compelling. A question arises, however, about the source, rostrocaudally, of the VSCT neurons and tract. Is the 10% for the total or for a specific level or levels? Can more be said/quantified about differences in these populations at different spinal levels?

3. Whole-body collapses and subsequent over-extensions are important and speak to changes in reflex arc and motor output. The statement "usually followed by" over-extension should be followed-up. Can this be further quantified? Are the two events linked or distinct, and did over-extensions happen in the absence of collapses?

4. These issues mesh with the lower knee height and angle of the TMP joint, even when collapses are excluded. It appears as though the control system to maintain muscle shortening (force output of extensors) is altered. I agree that stability is compromised, but could we go further to state that the compromise is due to extensor gain control?Reviewer #2:

This work addresses the possibility that developmentally-characterized di2 neurons contribute to the ventral spinocerebellar tract and regulate stepping in the chick. The work is sound considering that most information we have on spinal subtypes are for ventrally-born and local circuit interneurons (i.e. motor related), but less is known about the dorsally-born types and about long-range projecting neurons that link the spinal cord with higher integrative centers. Here, using a combination of cell-type specific manipulations, circuit tracing tools and kinematic analysis of gaits in the chick authors propose that spinal di2 interneurons contain multiple subgroups including a population that sends projection to the cerebellum. Silencing di2 neurons overal leads to impaired stepping.

Overall, the strategy is sound and there is potential novelty that make this paper a suitable candidate for eLife provided the weaknesses in the scientific demonstration listed below can be first addressed, experimentally and/or by additional analysis. Equally importantly also, the work suffers from a SEVERE lack of clarity (writing, figures, results). Both must be addressed for this paper to be considered further.

I start with the scientific weaknesses:

1. Synaptic connections rely mostly on the anatomical overlap between di2 cells and the synaptic field of their putative pre-synaptic partners. While this is indeed suggestive, it is not enough to ascertain actual synaptic connections, and even less so in a comparative manner between the different groups. Furthermore, some tracers (e.g PRVmCherry) do not seem to be under a synapse-specific promoter, so labelled elements might just as well be passing fibers. Clearer evidence of actual connections should be provided, functionally if possible or at the very least by showing clearer putative boutons onto neuronal somata/dendrites, quantifying them and quantifying differences between input cell types. Current figures (2F / 3B', C', D' / 4C, D', E', F') are not sufficiently convincing since we see only one cell and can barely detect boutons visually on some of them (not to mention that pseudo-colors keep changing, see other comment below). In addition, please consider using the term "putative" or "presumed" synapses, contacts and connections throughout the study.

2. The loss of function and gait analysis is stronger and convincingly presented. However, unless I missed it, the strategy silences all di2 neurons but cannot discriminate the contributions of the pre-cerebellar ones. This poses problems for the interpretation of the data. Since this paper is about either subpopulations of di2, or the vSCT (see other comment about general scope of the work), it would be more robust if more specific silencing was included. It is currently assumed that one likely mechanism for the disturbed gait owes to the function of di2 as precerebellar neurons (line 385, 389) but the phenotype could also, or even entirely, be due to their proprio-spinal connectivity. This is a major caveat.

On top of this, writing and data presentation MUST be substantially improved on multiple aspects:

3. Please have the manuscript deeply proofread. In addition to numerous English mistakes (missing "the", "or", plural and singulars, lots of unnecessary comas, etc…) examples of confused writing include (non-exhaustive list):

(a) Line 128: what does this phrase mean ("TF expression is redundant"…).

(b) Line 159: I don't understand here, the Di2 ascend to the cerebellum, cross the midline to the targeted di2?? To which Di2 do authors refer to here, it sounds like they are in the cerebellum, or that the ascending Di2 redescend to the spinal cord…

(c) The term targeted is in fact used alternatively and confusingly to refer to either "manipulated" cells, "synaptically-targeted" cells, there is also "targeted overground locomotion",….

(d) Stage HH18 is sometimes referred to as E3. Please be consistent throughout.

(e) When describing inputs onto di2, add "neurons" (i.e. "onto di2 neurons").

4. I would appreciate more background on di2 neurons in the introduction and why these have been investigated. Currently, most of this is given in the first paragraph of the results (lines 91-100 and also line 103). Also, it is stated first that "the role of di2 neurons is elusive due to the lack of genetic targeting means" (line 59). This contradicts the later statement that "the progenitor pdi2 expresses [various transcription factors]", and that the "post mitotic di2 are defined by…" (line 103). Please clarify what is known and not known about di2 already in the introduction.

5. Related to the above, it is not sufficiently clear what is investigated here. The genetic identity of ventral spinocerebellar neurons? Or the diversity of di2 neurons? In the way the introduction is written, it gives the impression that it is the former, but then functional investigations are not specific enough (since they are targeted to the overall di2 population, see dedicated comment later). Authors should revise to make clearer what is the scope of the work.

6. Histology Figures should be made more convincing, self-explanatory, and to a higher standard.

(a) Anatomical landmarks MUST be paced on ALL figures, e.g: the midline and minimal nuclei of the cerebellum, the deep cerebellar nuclei should be indicated in Figure S4,… Also, please give the orientation axis on ALL figures (especially the ones illustrating large territories, like 2B, 4A).

(b) Add the CTB or HSV tracer on Figure 2A and check coherence: I believe for instance that HSP is wrongly stated instead of HSV in Figure 2D and PRV is wrongly stated instead of CTB in Figure 2F (and there might be other confusions throughout).

(c) It is extremely confusing that histology pseudo-colors are sometimes changed from one related figure to the other, for unclear reasons (e.g. 2B, 2B', 2C, also 2C and S4A…). Consistency will help the reader go through all panels and figures comparatively.

(d) Figures must be addressed in proper order. This also applies to supplemental figures. Otherwise, it gives the impression we have missed something.

(e) What is the rationale for plotting the overlap in area versus volume (Figure 2H, I)? If overlap with area shows a higher percentage than with volume, does it mean that the overlap is only limited to a given A/P plane? I'm really confused about this representation and its meaning.

7. Authors should avoid relying on subjective formulations like "that reside at the lateral dorsal aspect of lamina VII". Instead, they MUST demonstrate the positioning of Di2 neurons into the different spinal laminae with some form of quantitative measurements. This is currently just an "impression" that large, precerebellar Di2 are more ventral, in lamina VII and possibly VIII but without the representation of lamina borders on figures, this information cannot be appreciated by the reader. It is all essential that these borders are depicted in Figures and neurons be quantitatively allocated to each laminae. In addition/alternatively, authors should report the average D/V position of the different subtypes and test for significant differences to make the case of different spatially-confined populations stronger.

8. FoxD3 expression on Supplemental Figure 2B is not convincing. It is also not reported in the statistics of Figure 1E. Do we have to assume that all di2 investigated here are FoxD3-positive? If so, one would need a better illustration and quantifications should be given. Otherwise, I would suggest to simply rely on literature and removing that Figure S1B which is not helping. On other panels of that supplemental Figure 2, please add arrow/arrowheads on all neurons that are or not co-labelled so we can appreciate co-labelling.

9. The demonstration that di2 are excitatory is essential. It is the title of a paragraph (line 102), thus I think that the corresponding data with the neurotransmitters (Vglut2, GAD) would deserve to be in the main Figures. Also, the chosen illustration only shows ONE double-labelled cell with Vglut2. Authors should be able to show a field of view that more convincingly conveys the message with more cells.Reviewer #3:

This study by Haimson et al., aims at examining the diversity of dI2 interneurons and their role in coordinating activity across difference region of the spinal cord and in reporting back activity to the brain. The results show that dI2 interneurons comprise different sub-classes based on their axonal projections, soma diameter and transmitter identity. They also show that some dI2 interneurons project rostrally from the lumbar spinal cord and make putative synaptic contacts with other dI2 interneurons in the brachial spinal cord on their way to the cerebellum. Finally, it is shown that some dI2 interneurons receive putative inputs from DRG neurons and may serve to transmit movement-related feedback. An indiscriminate silencing of dI2 interneurons results in instability of locomotion. Overall, this study reports some interesting observations by showing the heterogeneity of dI2 interneurons and their potential function. I have the following concerns:

1) 12% express Pax2 and are considered inhibitory. However, Gad is expressed in only 25% of dI2 interneurons while vGlut is expressed in 88%. These proportions suggest that there are dI2 neurons that co-express vGlut and Gad. Is this the case? Are there additional inhibitory dI2 neurons in addition to those expressing Pax2 which could explain the fact that Gad labels 25% of dI2 neurons. These points need some clarifications and discussion.

2) Of all dI2 interneurons, 91% are small diameter and 9% are large diameter neurons – large diameter neurons are mostly apparent in the lumbar spinal cord. The small and large diameter dI2 neurons cannot be differentiated by their expression of TFs, but can be distinguished by their transmitter identity? Is the proportion of small and large diameter neurons the same along the spinal cord?

3) Do all dI2 neurons receive putative synaptic contacts from DRG neurons? Unless I have missed it, it would be helpful to provide quantification of the number of small vs large diameter dI2 neurons with regard to the different putative synaptic contacts they receive from DRG neurons, dI2 and V1 interneurons.

4) Lines 218-220: It is stated that DRG putative contacts are mainly targeting dorsal dI2 neurons while ventral ones receive virtually no contacts. Since large diameter VSCT dI2 neurons are located ventrally, they do not seem to receive direct sensory information. However, the authors conclude that VSCT dI2 neurons receive sensory input (lines 227-228) and also in the Discussion. There seem to be a mismatch between the results and the conclusion drawn by the authors (lines 374-377). Unless I am missing something here, this is not consisting with the conclusions of this study. Please clarify.

5) The silencing experiments are interesting, however it is unclear which sub-class of dI2 neurons and at what level (lumbar vs brachial spinal cord or cerebellum) the observed behavioral perturbations take place. It is possible to selectively silence excitatory vs inhibitory or only VSCT neurons to provide some link between dI2 sub-classes and behavioral perturbations.

https://doi.org/10.7554/eLife.62001.sa1

Author response

Summary:

The three reviewers and I have carefully read your manuscript and we all have found a great interest in your results. However several concerns will have to be addressed in order to consider the paper further. In a first instance the paper requires careful revisions in the writing/editing, quantifications and clarity of the figures. Also the discussion on the main findings and their limitations should be revised in order to take into account reviewers' comments.

More specifically, the authors are asked to be more accurate/careful in describing

1) The motor phenotype,

In the original manuscript we described that there was a higher variability in the knee joint trajectory and TMP angle during the swing phase, a significantly higher number of collapses, and a wider base stepping in the manipulated compared to the control chickens. The swing velocity and the inter-limb phase relation during stepping in the manipulated chickens were similar to those of the un-manipulated controls. In the revised version of the manuscript, we analyzed two additional aspects of the motor phenotypes: (1) we examined whether slipping over the walking surface contributed to the observed instability (e.g. Clark and Higham, 2011, Daley and Biewener, 2006). (2) according to the request of Reviewer 1, we reanalyzed the occurrence of collapses during stepping and their relation to over-extensions.

1) Slipping and instability: Analyses of the landing angles during stepping revealed no significant differences between the manipulated and control chickens. Moreover, the angles preceding a collapse were not smaller but rather slightly higher than those that did not precede a collapse. Collectively these findings suggest that the instability does not involve over ground slipping. These analyses are shown in the new figure 7—figure supplement 2 and described on p. 34 lines 396-407, in the methods section lines 671-673.

2) Collapse and over-extension: Our analyses revealed that 65% of the collapses were not preceded by or followed by overextensions. Only 22.5% of the collapses, were followed by an overextension, and there was a high variability between chickens. In 12.5% of the cases the collapses were preceded by an overextension. These new analyses are summarized on p. 30-33 lines 358-369.

Together, the new analyses strengthen our original inferences outlined in the discussion, that the motor phenotype can be attributed to perturbed spinal information supply to the cerebellum. Furthermore, interference with the descending regulation of spinal motor output, or a lack of proper direct input to intraspinal premotor neurons. The few overextensions observed after collapses, may reflect a post-collapse compensation of the extensor drive (p. 30 lines 368-369).

2) the identity of which sub-population of neurons is considered to be responsible for the locomotor behavior (di2 neurons vs vSCT neurons),

Similar to other spinal cardinal populations of INs, dI2 neurons consistof several subpopulations. Among them, the VSCT dI2 and the propriospinal dI2. The division to these subclasses is based on the size of the neuron somata (larger versus small cell diameter), and the intra- and supra-spinal projection pattern. The lack of genetic means that enable targeting these dI2 sub-populations, hinders the analyses of their relative contribution to stable locomotion. An alternative experimental paradigm is to target specifically the VSCT dI2 by target-derived activation of neural-modulators. Namely, to activate neural modulator in dI2 neurons retrogradely from the cerebellum or from the contralateral lumbar spinal cord (injection of PRV-Cre into the cerebellum). In the original manuscript we demonstrated that dI2 axons send numerous collaterals along the entire extent of the spinal cord. The origin of these collaterals, either from the VSCT or the propriospinal dI2 neurons, could not be accurately resolved in the original manuscript. We suspected that the VSCT dI2 neurons also innervate the lumbar spinal cord, which may undermine the use of target-derived neural-modulators activation.

In order to resolve the connectivity of VSCT dI2 neurons, we employed lumbar-specific sparse labelling of dI2, and followed in a single-cell resolution, their entire lumbar to cerebellum trajectory patterns, utilizing light sheet microscopy. The new data are presented in the new figure 3, four supplementary videos, supplementary figure 3—figure supplement 1A (p. 15-16, lines 199-222). Our new data provides evidence that VSCT dI2 neurons send collaterals that innervate the contralateral targets along the entire path, from the lumbar spinal cord to the brainstem and the cerebellar nuclei.

This new analysis questions that validity of target-derived silencing of dI2 subpopulations, either via retrograde activation from the cerebellum or the contralateral lumbar spinal cord, since this approach will target both the propriospinal lumbar dI2 and the VSCT dI2 neurons. Future experiments, aimed to uncover the transcriptome of dI2, may provide genetic means to target dI2 subpopulations and subsequently to manipulate their neuronal activity. We discuss the relative potential contribution of these two subpopulations to stepping on pages 42-43 (lines 473-489).

In this regard, It should be noted that the physiological role of other spinal interneurons that control motor activity, like V1, V2, V3, V0, dI3 and dI1 (Yuengert et al., 2015, Gosgnach et al., 2006, Zhang et al., 2008, Talpalar et al., 2013, Bui et al., 2013, Crone et al., 2008), was previously studied via genetic targeting using regulatory elements of transcription factors expressed early in their development, similar to our targeting and silencing approach. Hence, our wiring and behavioral studies fall within the scope of the experimental strategies in the field.

3) the anatomical data of the neurons of interest, possibly including their neurotransmitter phenotype,

The new light sheet microscopy analyses (figures 3, figure 3—figure supplement 1A and the four supplementary videos) provide insights into this question. To complement these, we also preformed new in situ mRNA localization using vesicular inhibitory amino acid transporter (VIAAT) RNA probes to detect both GABAergic and glycinergic inhibitory interneurons. We also preformed new in situ hybridization experiments with a vGglut2 probe for detection of excitatory interneurons. The proportion of the excitatory/inhibitory dI2 neurons is 73%/27%. The new data are presented in figure 1G-I and in p.11 lines 150-153.

4) The quantification and characterization of the synaptic contacts and their potential functional role.

In the original manuscript we have used two criteria for detection of synaptic contacts to and from dI2: (1) The likelihood of connectivity was examined by spatial overlap of axonal terminals from the presumed presynaptic neurons and the somata of the post synaptic neurons. (2) Detection of synaptic boutons, labelled by a synaptic reporter, on the somatodendritic membrane of the post synaptic neurons. We also included an image showing an overlap between a synaptic reporter and a synaptic protein (former figure 3C, current figure 4C’).

In the revised manuscript we thoroughly tested the validity of the genetically-delivered synaptic reporter by quantifying the co-labelled staining of synaptic reporter with synaptotagmin. We used confocal imaging and 3D reconstruction using IMARIS software to score signal overlap. We tested the co-labelling of dI2::SV2-GFP and synaptotagmin in dI2 neurons synapses on contralateral pre-motoneurons. From 144 genetically labeled boutons, 121 (84%) were synaptotagmin+. The synaptotagmin- boutons were significantly smaller. We set a volume threshold, so that small volume SV2-GFP boutons were not considered as synapses throughout the study (Figure 4—figure supplement 1 and p. 20 lines 244-249).

Thus, we are confident that our synaptic reporter staining coupled with the labelling of the somata of presumed target neurons, reliably represent anatomical synapses. We also would like to note that similar anatomical-synaptic analysis, are the convention in the field of spinal circuitry (Baek et al., 2019, Dougherty et al., 2013, Goetz et al., 2015, Ruder et al., 2016).

Reviewer #1:

This is a well-put-together manuscript describing carefully performed circuitry dissection and functional analysis of dl2 neurons in the chick. A genetic toolbox is used taking advantage of the electroporation technique applied to the embryos. The findings include a fairly convincing connectome for dl2 neurons and a functional phenotype that is, unfortunately, rather unsatisfying. The investigators conclude that dl2 interneurons regulate "stability" of bipedal stepping in the chick, which is fine, but the analysis misses an opportunity to more fully explore what the instability involves and thus to perhaps shed more light on the likely roles of this neuron population. The concerns/issues 3 and 4 below focus on this issue and the need for additional careful analysis of the behavior that will allow the phenotype to be more precisely described or ascribed to some aspect of stepping that might guide future studies in other models. For example, can the link between partial collapse and over-extensions be made more solid and thus argue that reduced extensor gain might be what results in the instability? What other analysis could be performed using the existing data/video to better describe the behavioral phenotype?

Concerns/issues:

1. The connectome part of the work appears solid and supports the concept that a sub population of the population are likely VSCT neurons, that the non VSCT neurons receive the bulk of the afferent input and that these neurons project to contralateral dl2 neurons (some which may be VSCT) and other premotor neurons. Anatomically, the only concern is that no distinctions were made between the lumbar and brachial populations, and if differences in these populations exist, it would be important and interesting to describe them.

The present study was focused on the characterization of lumbar VSCT dI2 and their presumed contribution to hindlimb locomotion (see the modified title of the manuscript). We agree with the reviewer that the studies of the connectome and function of brachial populations of dI2 are important and interesting. We plan to study the brachial dI2 and the presumed role of lumbar to brachial dI2 connectivity in coordinating legs/wings movements in the near future.

2. Figure 2 Characterization of dl2/VSCT neurons as being primarily large dl2 neurons is quite convincing, and the observation that the dl2 neurons account for 10% of the VSCT axons is also of interest and quite compelling. A question arises, however, about the source, rostrocaudally, of the VSCT neurons and tract. Is the 10% for the total or for a specific level or levels? Can more be said/quantified about differences in these populations at different spinal levels?

See our response to the general comment #2 above, in pages 2-3 in this document. As stated in the manuscript “large-diameter dI2 neurons are only apparent at the lumbar level” (p. 12, lines 158).

3. Whole-body collapses and subsequent over-extensions are important and speak to changes in reflex arc and motor output. The statement "usually followed by" over-extension should be followed-up. Can this be further quantified? Are the two events linked or distinct, and did over-extensions happen in the absence of collapses?

4. These issues mesh with the lower knee height and angle of the TMP joint, even when collapses are excluded. It appears as though the control system to maintain muscle shortening (force output of extensors) is altered. I agree that stability is compromised, but could we go further to state that the compromise is due to extensor gain control?

See our response to the general comment #1 above. We omitted the term “usually followed” and modified the text accordingly (p. 30-33, lines 358-369).

Reviewer #2:

This work addresses the possibility that developmentally-characterized di2 neurons contribute to the ventral spinocerebellar tract and regulate stepping in the chick. The work is sound considering that most information we have on spinal subtypes are for ventrally-born and local circuit interneurons (i.e. motor related), but less is known about the dorsally-born types and about long-range projecting neurons that link the spinal cord with higher integrative centers. Here, using a combination of cell-type specific manipulations, circuit tracing tools and kinematic analysis of gaits in the chick authors propose that spinal di2 interneurons contain multiple subgroups including a population that sends projection to the cerebellum. Silencing di2 neurons overal leads to impaired stepping.

Overall, the strategy is sound and there is potential novelty that make this paper a suitable candidate for eLife provided the weaknesses in the scientific demonstration listed below can be first addressed, experimentally and/or by additional analysis. Equally importantly also, the work suffers from a SEVERE lack of clarity (writing, figures, results). Both must be addressed for this paper to be considered further.

I start with the scientific weaknesses:

1. Synaptic connections rely mostly on the anatomical overlap between di2 cells and the synaptic field of their putative pre-synaptic partners. While this is indeed suggestive, it is not enough to ascertain actual synaptic connections, and even less so in a comparative manner between the different groups. Furthermore, some tracers (e.g PRVmCherry) do not seem to be under a synapse-specific promoter, so labelled elements might just as well be passing fibers. Clearer evidence of actual connections should be provided, functionally if possible or at the very least by showing clearer putative boutons onto neuronal somata/dendrites, quantifying them and quantifying differences between input cell types. Current figures (2F / 3B', C', D' / 4C, D', E', F') are not sufficiently convincing since we see only one cell and can barely detect boutons visually on some of them (not to mention that pseudo-colors keep changing, see other comment below). In addition, please consider using the term "putative" or "presumed" synapses, contacts and connections throughout the study.

See our response to the general comment #4 above (the quantification and characterization of the synaptic contacts and their potential functional role).

We agree with the reviewer that demonstrating functional connections is the gold standard for synaptic connectivity. However, we would like to note that the chick is not a common model organism for studying neuronal circuitry and post embryonic motor behavior in genetically manipulated embryos. Hence, many of the circuit-deciphering tools are not yet applicable to avian. The tools that we employed for decoding circuitry in the chick spinal cord, were mostly developed and implemented by us (Hadas et al., 2014). The main caveat is the non-germline targeting of specific neurons via electroporation. The efficiency of transgenesis via electroporation varies from 10-60%. Hence, the number of labelled putative synapses between two genetically-labeled neurons is markedly lower from the expected number in germline targeted mice. In addition, performance of in vitro recording from spinal cord neurons in chick, for demonstrating functional connectivity, is limited to early stages of embryonic development and are not applicable post E12 (O'Donovan et al., 1994). At this stage the network has not yet matured. Our efforts to modify the experimental conditions has not yet materialized.

As to the use of PRV-cherry to label contacts between pre-MNs and dI2, we agree with reviewer that this type of contacts is suggestive. We noted this in line 268-270. However, the use of 3D confocal imaging enables us to distinguish between a terminal and a passing-by axon. Importantly, in the following experiment we also use synaptic reporters expressed in two type of pre-MNs (dI1 and V1) to provide supportive evidence to the connection between pre-MNs and dI2.

2. The loss of function and gait analysis is stronger and convincingly presented. However, unless I missed it, the strategy silences all di2 neurons but cannot discriminate the contributions of the pre-cerebellar ones. This poses problems for the interpretation of the data. Since this paper is about either subpopulations of di2, or the vSCT (see other comment about general scope of the work), it would be more robust if more specific silencing was included. It is currently assumed that one likely mechanism for the disturbed gait owes to the function of di2 as precerebellar neurons (line 385, 389) but the phenotype could also, or even entirely, be due to their proprio-spinal connectivity. This is a major caveat.

See our response to the general comment #2 above (The identity of which sub-population of neurons is considered to be responsible for the locomotor behavior).

On top of this, writing and data presentation MUST be substantially improved on multiple aspects:

3. Please have the manuscript deeply proofread. In addition to numerous English mistakes (missing "the", "or", plural and singulars, lots of unnecessary comas, etc…) examples of confused writing include (non-exhaustive list):

(a) Line 128: what does this phrase mean ("TF expression is redundant"…)

We have corrected this to: “…their expression is not required after the establishment of the circuitry”. The manuscript was deeply edited.

(b) Line 159: I don't understand here, the Di2 ascend to the cerebellum, cross the midline to the targeted di2?? To which Di2 do authors refer to here, it sounds like they are in the cerebellum, or that the ascending Di2 redescend to the spinal cord…

This was a typo. The corrected sentence is “…and cross back to the other side of the cerebellum ipsilaterally to the targeted granular neurons”.

(c) The term targeted is in fact used alternatively and confusingly to refer to either "manipulated" cells, "synaptically-targeted" cells, there is also "targeted overground locomotion",….

The word targeted in the sentence: "targeted overground locomotion" is indeed redundant and was omitted. We reduced the use of “target” to indicate the putative post-synaptic neurons of dI2, throughout the manuscript. However, we didn’t eliminate it completely, since the use of “target” in conjunction to both genetic-targeting and synaptic-target is acceptable and very common in the field of spinal circuitry.

(d) Stage HH18 is sometimes referred to as E3. Please be consistent throughout.

We use now the term HH18 throughout the manuscript.

(e) When describing inputs onto di2, add "neurons" (i.e. "onto di2 neurons").

The term ״neurons״ is now added.

4. I would appreciate more background on di2 neurons in the introduction and why these have been investigated. Currently, most of this is given in the first paragraph of the results (lines 91-100 and also line 103). Also, it is stated first that "the role of di2 neurons is elusive due to the lack of genetic targeting means" (line 59). This contradicts the later statement that "the progenitor pdi2 expresses [various transcription factors]", and that the "post mitotic di2 are defined by…" (line 103). Please clarify what is known and not known about di2 already in the introduction.

A new background paragraph was added to the introduction on p. 3, lines 59-70.

5. Related to the above, it is not sufficiently clear what is investigated here. The genetic identity of ventral spinocerebellar neurons? Or the diversity of di2 neurons? In the way the introduction is written, it gives the impression that it is the former, but then functional investigations are not specific enough (since they are targeted to the overall di2 population, see dedicated comment later). Authors should revise to make clearer what is the scope of the work.

We thank the reviewer for highlighting this issue. The focus of the study is indeed dI2. We modified the introduction section accordingly. As to dI2 subpopulations, see our response to comment #2 in the summary section of the eLife decision letter.

6. Histology Figures should be made more convincing, self-explanatory, and to a higher standard.

(a) Anatomical landmarks MUST be paced on ALL figures, e.g: the midline and minimal nuclei of the cerebellum, the deep cerebellar nuclei should be indicated in Figure S4,… Also, please give the orientation axis on ALL figures (especially the ones illustrating large territories, like 2B, 4A).

We replaced the image of the central cerebellar nuclei in figure 2—figure supplement 1A (the previous S3A). A low magnification image is now included and the midline and the margins of the central cerebellar nuclei are indicated. We are not aware to the term “minimal nuclei of the cerebellum”.

We inserted coordinates to indicate the rostral-caudal ipsi-contra and medial-lateral axes. However, we did not indicate the ventral-dorsal axis in the cross-section images, since it is acceptable that ventral is down in these images.

(b) Add the CTB or HSV tracer on Figure 2A and check coherence: I believe for instance that HSP is wrongly stated instead of HSV in Figure 2D and PRV is wrongly stated instead of CTB in Figure 2F (and there might be other confusions throughout).

In Figure 2D HSP is indeed HSV. In Figure 2F we have used PRV-cherry. We calibrated the post-injection time that is required for the infection of lumbar level pre-cerebellar neurons. We also co-injected PRV-cherry+CTB to verify that just the pre-cerebellar (not the pre-pre-cerebellar) neurons are labelled.

We amended the text accordingly on p. 15 lines 187-189.

(c) It is extremely confusing that histology pseudo-colors are sometimes changed from one related figure to the other, for unclear reasons (e.g. 2B, 2B', 2C, also 2C and S4A…). Consistency will help the reader go through all panels and figures comparatively.

We have now adjusted the colors: dI2 neurons are cyan, pre-dI2 are magenta and post-dI2 neurons are yellow.

(d) Figures must be addressed in proper order. This also applies to supplemental figures. Otherwise, it gives the impression we have missed something.

Done.

(e) What is the rationale for plotting the overlap in area versus volume (Figure 2H, I)? If overlap with area shows a higher percentage than with volume, does it mean that the overlap is only limited to a given A/P plane? I'm really confused about this representation and its meaning.

We agree that the added value in showing the overlapping histograms is marginal. Instead, we inserted a laminar distribution chart of each type of neuron and synapses, as suggested by the reviewer, in figures 1, 2, 4 and 5 and the corresponding supplemental figures. The overlap between the cell-to-cell and synapses-to-cells is also apparent in this presentation.

7. Authors should avoid relying on subjective formulations like "that reside at the lateral dorsal aspect of lamina VII". Instead, they MUST demonstrate the positioning of Di2 neurons into the different spinal laminae with some form of quantitative measurements. This is currently just an "impression" that large, precerebellar Di2 are more ventral, in lamina VII and possibly VIII but without the representation of lamina borders on figures, this information cannot be appreciated by the reader. It is all essential that these borders are depicted in Figures and neurons be quantitatively allocated to each laminae. In addition/alternatively, authors should report the average D/V position of the different subtypes and test for significant differences to make the case of different spatially-confined populations stronger.

The laminar distribution of all the neurons and synapses is now included. See figures 1,2,4 and 5 and the corresponding supp figures.

8. FoxD3 expression on Supplemental Figure 2B is not convincing. It is also not reported in the statistics of Figure 1E. Do we have to assume that all di2 investigated here are FoxD3-positive? If so, one would need a better illustration and quantifications should be given. Otherwise, I would suggest to simply rely on literature and removing that Figure S1B which is not helping. On other panels of that supplemental Figure 2, please add arrow/arrowheads on all neurons that are or not co-labelled so we can appreciate co-labelling.

As suggested by the reviewer we now show only the expression of FoxD3 in the post-mitotic pre-migratory dI2, and we provide the proper citation for Foxd3 expression. We preformed new in situ experiment and modified Figure 1—figure supplement 2B. We added arrow/arrowheads on all neurons in figure 1—figure supplement 2.

Figure 1—figure supplement 1B is informative for the general audience. Thus, we think that it is important to include it.

9. The demonstration that di2 are excitatory is essential. It is the title of a paragraph (line 102), thus I think that the corresponding data with the neurotransmitters (Vglut2, GAD) would deserve to be in the main Figures. Also, the chosen illustration only shows ONE double-labelled cell with Vglut2. Authors should be able to show a field of view that more convincingly conveys the message with more cells.

We preformed new in situ mRNA localization experiment, this time using the VIAAT probe for detecting the inhibitory dI2 neurons. The proportion of the excitatory/inhibitory dI2 is 73%/27% (see figure 1G-I).

Reviewer #3:

This study by Haimson et al., aims at examining the diversity of dI2 interneurons and their role in coordinating activity across difference region of the spinal cord and in reporting back activity to the brain. The results show that dI2 interneurons comprise different sub-classes based on their axonal projections, soma diameter and transmitter identity. They also show that some dI2 interneurons project rostrally from the lumbar spinal cord and make putative synaptic contacts with other dI2 interneurons in the brachial spinal cord on their way to the cerebellum. Finally, it is shown that some dI2 interneurons receive putative inputs from DRG neurons and may serve to transmit movement-related feedback. An indiscriminate silencing of dI2 interneurons results in instability of locomotion. Overall, this study reports some interesting observations by showing the heterogeneity of dI2 interneurons and their potential function. I have the following concerns:

1) 12% express Pax2 and are considered inhibitory. However, Gad is expressed in only 25% of dI2 interneurons while vGlut is expressed in 88%. These proportions suggest that there are dI2 neurons that co-express vGlut and Gad. Is this the case? Are there additional inhibitory dI2 neurons in addition to those expressing Pax2 which could explain the fact that Gad labels 25% of dI2 neurons. These points need some clarifications and discussion.

Pax2 labels GABAergic inhibitory neurons. In order the label all the inhibitory interneurons we have now employed VIAAT as a probe. See also our response to the general comment #3 above.

2) Of all dI2 interneurons, 91% are small diameter and 9% are large diameter neurons – large diameter neurons are mostly apparent in the lumbar spinal cord. The small and large diameter dI2 neurons cannot be differentiated by their expression of TFs, but can be distinguished by their transmitter identity? Is the proportion of small and large diameter neurons the same along the spinal cord?

We have now analyzed the number of inhibitory/excitatory neuron is the small and large diameter dI2 neurons. We found that the excitatory/inhibitory ratio in both subpopulation is similar. These new data are presented in Figure 1G-I and in line 150-153.

3) Do all dI2 neurons receive putative synaptic contacts from DRG neurons? Unless I have missed it, it would be helpful to provide quantification of the number of small vs large diameter dI2 neurons with regard to the different putative synaptic contacts they receive from DRG neurons, dI2 and V1 interneurons.

We have now analyzed the synaptic inputs of DRG neurons to dI2 neurons. This quantification is now presented in figure 4—figure supplement 3E,F (p. 21, lines 261-267).

4) Lines 218-220: It is stated that DRG putative contacts are mainly targeting dorsal dI2 neurons while ventral ones receive virtually no contacts. Since large diameter VSCT dI2 neurons are located ventrally, they do not seem to receive direct sensory information. However, the authors conclude that VSCT dI2 neurons receive sensory input (lines 227-228) and also in the Discussion. There seem to be a mismatch between the results and the conclusion drawn by the authors (lines 374-377). Unless I am missing something here, this is not consisting with the conclusions of this study. Please clarify.

We have now analyzed the number of contacts between DRG axons and dorsal/ventral dI2 and large/small dI2 neurons. The number of contacts with the dorsal dI2 neurons is significantly higher than with the ventral dI2. However, the number of contacts with the small and large dI2 neurons is similar. The decrease of contacts in the ventral dI2 is mainly due to fewer contacts with the small ventral dI2. These new data are presented in Figure 4—figure supplement 3E,F.

5) The silencing experiments are interesting, however it is unclear which sub-class of dI2 neurons and at what level (lumbar vs brachial spinal cord or cerebellum) the observed behavioral perturbations take place. It is possible to selectively silence excitatory vs inhibitory or only VSCT neurons to provide some link between dI2 sub-classes and behavioral perturbations.

See our response to the general comment #2 above.

References

Baek, M., Menon, V., Jessell, T. M., Hantman, A. W. And Dasen, J. S. 2019. Molecular Logic of Spinocerebellar Tract Neuron Diversity and Connectivity. Cell Rep, 27, 2620-2635 e4.

Bikoff, J. B., Gabitto, M. I., Rivard, A. F., Drobac, E., Machado, T. A., Miri, A., Brenner-Morton, S., Famojure, E., Diaz, C., Alvarez, F. J., Mentis, G. Z. and Jessell, T. M. 2016. Spinal Inhibitory Interneuron Diversity Delineates Variant Motor Microcircuits. Cell, 165, 207-219.

Bui, T. V., Akay, T., Loubani, O., Hnasko, T. S., Jessell, T. M. And Brownstone, R. M. 2013. Circuits for grasping: spinal dI3 interneurons mediate cutaneous control of motor behavior. NEURON, 78, 191-204.

Clark, A. J. And Higham, T. E. 2011. Slipping, sliding and stability: locomotor strategies for overcoming low-friction surfaces. J Exp Biol, 214, 1369-78.

Crone, S. A., Quinlan, K. A., Zagoraiou, L., Droho, S., Restrepo, C. E., Lundfald, L., Endo, T., Setlak, J., Jessell, T. M., Kiehn, O. And Sharma, K. 2008. Genetic ablation of V2a ipsilateral interneurons disrupts left-right locomotor coordination in mammalian spinal cord. Neuron, 60, 70-83.

Daley, M. A. And Biewener, A. A. 2006. Running over rough terrain reveals limb control for intrinsic stability. Proc Natl Acad Sci U S A, 103, 15681-6.

Dougherty, K. J., Zagoraiou, L., Satoh, D., Rozani, I., Doobar, S., Arber, S., Jessell, T. M. And Kiehn, O. 2013. Locomotor rhythm generation linked to the output of spinal shox2 excitatory interneurons. Neuron, 80, 920-33.

Goetz, C., Pivetta, C. And Arber, S. 2015. Distinct limb and trunk premotor circuits establish laterality in the spinal cord. Neuron, 85, 131-144.

Gosgnach, S., Lanuza, G. M., Butt, S. J., Saueressig, H., Zhang, Y., Velasquez, T., Riethmacher, D., Callaway, E. M., Kiehn, O. And Goulding, M. 2006. V1 spinal neurons regulate the speed of vertebrate locomotor outputs. Nature, 440, 215-9.

Hadas, Y., Etlin, A., Falk, H., Avraham, O., Kobiler, O., Panet, A., Lev-Tov, A. And Klar, A. 2014. A 'tool box' for deciphering neuronal circuits in the developing chick spinal cord. Nucleic Acids Res, 42, e148.

Hamburger, V. And Oppenheim, R. 1967. Prehatching motility and hatching behavior in the chick. J Exp Zool, 166, 171-203.

Hammar, I., Bannatyne, B. A., Maxwell, D. J., Edgley, S. A. And Jankowska, E. 2004. The actions of monoamines and distribution of noradrenergic and serotoninergic contacts on different subpopulations of commissural interneurons in the cat spinal cord. Eur J Neurosci, 19, 1305-16.

Hammar, I. And Maxwell, D. J. 2002. Serotoninergic and noradrenergic axons make contacts with neurons of the ventral spinocerebellar tract in the cat. J Comp Neurol, 443, 310-9.

O'donovan, M., Ho, S. And Yee, W. 1994. Calcium imaging of rhythmic network activity in the developing spinal cord of the chick embryo. The Journal of neuroscience : the official journal of the Society for Neuroscience, 14, 6354-69.

Provine, R. R., Sharma, S. C., Sandel, T. T. And Hamburger, V. 1970. Electrical activity in the spinal cord of the chick embryo, in situ. Proc Natl Acad Sci U S A, 65, 508-15.

Ruder, L., Takeoka, A. And Arber, S. 2016. Long-Distance Descending Spinal Neurons Ensure Quadrupedal Locomotor Stability. Neuron, 92, 1063-1078.

Talpalar, A. E., Bouvier, J., Borgius, L., Fortin, G., Pierani, A. And Kiehn, O. 2013. Dual-mode operation of neuronal networks involved in left-right alternation. Nature, 500, 85-8.

Yuengert, R., Hori, K., Kibodeaux, E. E., Mcclellan, J. X., Morales, J. E., Huang, T. P., Neul, J. L. And Lai, H. C. 2015. Origin of a Non-Clarke's Column Division of the Dorsal Spinocerebellar Tract and the Role of Caudal Proprioceptive Neurons in Motor Function. Cell Rep, 13, 1258-1271.

Zhang, Y., Narayan, S., Geiman, E., Lanuza, G. M., Velasquez, T., Shanks, B., Akay, T., Dyck, J., Pearson, K., Gosgnach, S., Fan, C.-M. And Goulding, M. 2008. V3 spinal neurons establish a robust and balanced locomotor rhythm during walking. NEURON, 60, 84-96.

https://doi.org/10.7554/eLife.62001.sa2

Article and author information

Author details

  1. Baruch Haimson

    Department of Medical Neurobiology, IMRIC, Hebrew University – Hadassah Medical School, Jerusalem, Israel
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Writing - original draft, Writing - review and editing
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0163-6196
  2. Yoav Hadas

    Department of Medical Neurobiology, IMRIC, Hebrew University – Hadassah Medical School, Jerusalem, Israel
    Contribution
    Data curation, Formal analysis, Investigation, Methodology
    Competing interests
    none
  3. Nimrod Bernat

    Department of Medical Neurobiology, IMRIC, Hebrew University – Hadassah Medical School, Jerusalem, Israel
    Contribution
    Methodology
    Competing interests
    none
  4. Artur Kania

    Institut de recherches cliniques de Montréal (IRCM), Montréal, Canada
    Contribution
    Conceptualization, Writing - review and editing
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5209-2520
  5. Monica A Daley

    Ecology and Evolutionary Biology, University of California, Irvine, Irvine, United States
    Contribution
    Conceptualization, Data curation, Writing - original draft
    Competing interests
    none
  6. Yuval Cinnamon

    Institute of Animal Science Poultry and Aquaculture Sci. Dept. Agricultural Research Organization, The Volcani Center, Rishon LeZion, Israel
    Contribution
    Conceptualization, Data curation, Methodology
    Competing interests
    none
  7. Aharon Lev-Tov

    Department of Medical Neurobiology, IMRIC, Hebrew University – Hadassah Medical School, Jerusalem, Israel
    Contribution
    Conceptualization, Formal analysis, Investigation, Methodology, Supervision, Validation, Writing - original draft, Writing - review and editing
    For correspondence
    aharonl@ekmd.huji.ac.il
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3906-0057
  8. Avihu Klar

    Department of Medical Neurobiology, IMRIC, Hebrew University – Hadassah Medical School, Jerusalem, Israel
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Resources, Supervision, Validation, Writing - original draft, Writing - review and editing
    For correspondence
    avihu@mail.huji.ac.il
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9248-2179

Funding

Israel Science Foundation (1400/16)

  • Avihu Klar

United States - Israel Binational Science Foundation (2017/172)

  • Avihu Klar

The Avraham and Ida Baruch Endowment Fund

  • Avihu Klar

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

The authors thank Haya Falk for PRV purification; Alona Katzir, Cole Bendor, Mevaseret Avital, Sapir Shevah, Eitan Yisraeli, Ruth Segal, Fedaa Bazan, and Eden Kimchi for technical assistance; Nadav Yayon for assistance with the light sheet microscopy; and Michael O’Donovan for comments on the manuscript. This work was supported by grants to AK from the Israel Science Foundation (grant no. 1400/16), the US–Israel Binational Science Foundation (grant no. 2017/172), and the Avraham and Ida Baruch endowment fund.

Ethics

All experiments involved with animals were conducted in accordance with the designated Experiments in Animals Ethic Committee policies and under its approval.

Senior Editor

  1. Ronald L Calabrese, Emory University, United States

Reviewing Editor

  1. Muriel Thoby-Brisson, CNRS Université de Bordeaux, France

Reviewers

  1. David SK Magnuson, University of Louisville, United States
  2. Julien Bouvier, Université Paris-Saclay, CNRS, France

Publication history

  1. Preprint posted: January 8, 2020 (view preprint)
  2. Received: August 11, 2020
  3. Accepted: August 11, 2021
  4. Accepted Manuscript published: August 16, 2021 (version 1)
  5. Version of Record published: September 17, 2021 (version 2)

Copyright

© 2021, Haimson et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

Metrics

  • 671
    Page views
  • 106
    Downloads
  • 0
    Citations

Article citation count generated by polling the highest count across the following sources: Crossref, PubMed Central, Scopus.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Download citations (links to download the citations from this article in formats compatible with various reference manager tools)

Open citations (links to open the citations from this article in various online reference manager services)

Further reading

    1. Developmental Biology
    2. Stem Cells and Regenerative Medicine
    Alessandro Bonfini et al.
    Research Article

    The gut is the primary interface between an animal and food, but how it adapts to qualitative dietary variation is poorly defined. We find that the Drosophila midgut plastically resizes following changes in dietary composition. A panel of nutrients collectively promote gut growth, which sugar opposes. Diet influences absolute and relative levels of enterocyte loss and stem cell proliferation, which together determine cell numbers. Diet also influences enterocyte size. A high sugar diet inhibits translation and uncouples ISC proliferation from expression of niche-derived signals but, surprisingly, rescuing these effects genetically was not sufficient to modify diet's impact on midgut size. However, when stem cell proliferation was deficient, diet's impact on enterocyte size was enhanced, and reducing enterocyte-autonomous TOR signaling was sufficient to attenuate diet-dependent midgut resizing. These data clarify the complex relationships between nutrition, epithelial dynamics, and cell size, and reveal a new mode of plastic, diet-dependent organ resizing.

    1. Developmental Biology
    2. Physics of Living Systems
    Yonghyun Song, Changbong Hyeon
    Research Article Updated

    Spatial boundaries formed during animal development originate from the pre-patterning of tissues by signaling molecules, called morphogens. The accuracy of boundary location is limited by the fluctuations of morphogen concentration that thresholds the expression level of target gene. Producing more morphogen molecules, which gives rise to smaller relative fluctuations, would better serve to shape more precise target boundaries; however, it incurs more thermodynamic cost. In the classical diffusion-depletion model of morphogen profile formation, the morphogen molecules synthesized from a local source display an exponentially decaying concentration profile with a characteristic length λ. Our theory suggests that in order to attain a precise profile with the minimal cost, λ should be roughly half the distance to the target boundary position from the source. Remarkably, we find that the profiles of morphogens that pattern the Drosophila embryo and wing imaginal disk are formed with nearly optimal λ. Our finding underscores the cost-effectiveness of precise morphogen profile formation in Drosophila development.