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From: Katja S. Grossmann, Aurore Giraudin, Olivier Britz, Jingming Zhang
and Martyn Goulding, Genetic dissection of rhythmic motor networks
in mice. In Jean-Pierre Gossard, Réjean Dubuc and Arlette Kolta,
editors: Progress in Brain Reasearch, Vol. 187,
Burlington: Academic Press, 2010, pp. 19-37.
ISBN: 978-0-444-53613-6© Copyright 2010 Elsevier BV.
Academic Press.

Author's personal copy
Jean-Pierre Gossard, Réjean Dubuc and Arlette Kolta (Eds.)
Progress in Brain Research, Vol. 187
ISSN: 0079-6123
Copyright Ó 2010 Elsevier B.V. All rights reserved.

CHAPTER 2

Genetic dissection of rhythmic motor networks
in mice
Katja S. Grossmann1, Aurore Giraudin1, Olivier Britz, Jingming Zhang
and Martyn Goulding*
Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, California, USA

Abstract: Simple motor behaviors such as locomotion and respiration involve rhythmic and coordinatedmuscle movements that are generated by central pattern generator (CPG) networks in the spinal cordand hindbrain. These CPG networks produce measurable behavioral outputs and thus represent ideal
model systems for studying the operational principles that the nervous system uses to produce specificbehaviors. Recent advances in our understanding of the transcriptional code that controls neuronaldevelopment have provided an entry point into identifying and targeting distinct neuronal populations
that make up locomotor CPG networks in the spinal cord. This has spurred the development of new
genetic approaches to dissect and manipulate neuronal networks both in the spinal cord and hindbrain.
Here we discuss how the advent of molecular genetics together with anatomical and physiologicalmethods has begun to revolutionize studies of the neuronal networks controlling rhythmic motor
behaviors in mice.
Keywords: locomotion; respiration; spinal cord; CPG; interneuron; mouse genetics.

Rhythmic motor outputs such as locomotion, respiration, and mastication are, in their simplest
forms, highly stereotyped motor behaviors. In fish
and tadpoles, locomotion primarily consists ofrepetitive lateral bending movements of the axis,
which are produced by waves of contraction andrelaxation that propagate rostrocaudally along
the body axis. In contrast, terrestrial vertebrates

propel themselves by flexing and extending
their limbs. This more complex mode of locomotion requires extensive coordination between theforelimbs and hindlimbs on each side of the animaland between individual limb joints. The additional
demands of land-based locomotion appear to have
driven the evolutionary elaboration of the spinal
motor circuitry in terrestrial vertebrates, both interms of numbers and types of neurons, as well as
sensory and supraspinal connectivity. In higher
vertebrates, the spinal locomotor system has thus

*Corresponding author, 1equal contribution.
Tel.: þ1-858-453-4100-1558; Fax: þ1-858-450-2172.
DOI: 10.1016/S0079-6123(10)87002-5

19

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20

evolved into a highly dynamic network that
produces a varied and flexible array of motor outputs in response to sensory feedback pathways
and descending influences from rubrospinal,
reticulospinal, vestibulospinal, and corticospinal
pathways.

Rhythmic motor circuits in the hindbrain
and spinal cord
The core neuronal networks that control rhythmic
respiratory and locomotor motor behaviors residein the hindbrain and spinal cord, respectively.
These CPG networks generate simple organizedmotor rhythms in an autonomous manner. Initialefforts to decipher the general organization ofthese simple motor CPGs in vertebrates reliedheavily on electrophysiological and pharmacological approaches. Such efforts were greatly aided
by the development of in vitro hindbrain–spinal
cord preparations in neonatal rodents (Kudoand Yamada, 1987; Smith and Feldman, 1987).
In addition to enabling investigators to localize
two rhythmic CPGs in the medulla that drive
respiratory movements, the pre-Bötzinger complex and parafacial respiratory group (Onimaru
and Homma, 1987; Smith et al., 1991; this book),
this in vitro preparation has been used to map
the hindlimb locomotor CPG. For instance, serially sectioning the lumbar spinal cord, bothrostrocaudally and dorsoventrally, has defined
the minimal area needed to generate a locomotor
rhythm. The area that is circumscribed encompasses laminae VII, VIII, and X of the lumbar
cord, with lumbar segments L1–L2 being particularly important for rhythm generation (Bracci
et al., 1996; Cazalets et al., 1995; Kjaerulff andKiehn, 1996). Forelimb movements are likewisecontrolled by networks distributed in the cervicalspinal cord (Ballion et al., 2001). Further insights
into the organization of the spinal locomotor
CPG network have come from in vitro pharmacological studies. From these studies, it appears that
locomotor rhythmogenesis is primarily reliant on

glutamatergic ipsilateral neurons (Bracci et al.,
1996; Kjaerulff and Kiehn, 1997), whereas thealternating patterns of flexor–extensor andleft–right activities are secured by GABAergic
and glycinergic pathways (Bracci et al., 1996;
Kjaerulff and Kiehn, 1996, 1997).
Over the past few years, efforts to delineate thegenetic programs that pattern the caudal neuroaxis
have provided new avenues for probing the composition of these CPG networks. An importantthematic that has emerged has been the role thatgenetically driven developmental programs playin directing neuronal specification and differentiation in the hindbrain and spinal cord. Significant
progress has been made in understanding how suchdevelopmental programs determine neuronal
identity and how neurons are wired together intofunctional motor circuits. These studies suggest
commonalities in the cellular composition of CPG
networks in the hindbrain and spinal cord, in so
far that these circuits are built from interneuron
cell types that share a similar embryonic heritage.
In this review, we will focus on genetic approaches
used in mice to identify cells that are components
of motor networks and assess their contribution
to rhythmic motor behaviors, with an emphasis
on locomotion.

The developmental program of the caudal
neuroaxis
The developmental events that pattern the caudal
neural tube play a central role in assembling sensorimotor circuits in the hindbrain and spinal cord
(Goulding, 2009; Jessell, 2000). The position thata progenitor cell occupies along the dorsoventral(DV) axis confers a specific genetic code to thesecells and thus serves as a major determinant ofcell identity (Fig. 1a). This DV patterning program segregates newborn neurons into generic
populations that are arrayed as longitudinalcolumns along the antero-posterior axis of thehindbrain and spinal cord. As development proceeds, neurons within these columns migrate

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21
(a)

(b)

Embryonic

dI2

Isl1

dI3

Pax2

dI4

pd5

Lmx1b

dI5

pd6

WT1

dI6

pV0

Evx1

V0

En1

V1

pd3
pd4

pV2
Nkx2-2

Nkx6-1

pV1

Chx10/Gata3

V2

pMN
Hb9
pV3
Sim1

Motor

Pax3/7
Pax6

Foxd3

pd2

Sensory

dI1

Lhx2
pd1

MN
V3

Progenitors

Postmitotic

Neurotransmitter

Axonalprojection

Dbx2
Neurog1/2

dI6 WT

GABA/glycine

Commissural

Dbx1
Dbx2

V0D ?
V0V Evx1
V0C Pitx2

GABA/glycine
Glutamate
Acetylcholine

Commissural
Commissural
Ipsilateral

Dbx2
Nkx6-2V1

En1

GABA/glycine

Ipsilateral

Nkx6-1
Foxn4
Ascl1

V2a Chx10
V2b Gata3

Glutamate
GABA / glycine

Ipsilateral

Olig2
Nkx6-1
Neurog2

MN Hb9

Acetylcholine

Peripheral

Nkx2-2
Nkx6-1
Neurog3

V3

GlutamateCommissuralfew Ipsilateral

Sim1

(c)
Postnatal

X
V0C
V1
?

V2b
V1
IaIN

VII

V2a
V0
dI6

?
VIII

V1
IaIN
IX

V3
MN

V1
RC

V1
RC

MN

Fig. 1. Genetic classification, properties, and organization of spinal neurons. (a) Schematic cross-section through the early
embryonic mouse spinal cord. Eleven progenitor domains (pd1-pV3) have been identified in the ventricular zone that give rise todistinct postmitotic neuronal populations (dI1-V3). Selected transcription factors expressed within the progenitor domains areshown on the left and those specific to postmitotic populations of neurons born around embryonic day E11 are depicted on theright. Postmitotic neurons that contribute to the locomotor CPG machinery are shown in color, whereas neurons implicated insensory pathways are shown in gray. (b) Summary of molecular factors as well as cellular and physiological properties thatcharacterize each locomotor neuronal class. (c) Schematic of the motor circuitry in the postnatal ventral spinal cord. The color
code used is the same as in (a). The locations of neuron classes and their known subsets, as well as some known connections
between these classes, are illustrated. p, progenitor domain. (For interpretation of the references to color in this figure, thereader is referred to the Web version of this chapter.)Author's personal copy
22

extensively to form the different laminae seen in
the adult cord. Many premotor commissural interneuron cell types, for example, migrate from theintermediate zone to lamina VIII, while ventral
interneurons that project ipsilaterally migrate laterally to lamina VII (Fig. 1c). Interestingly, theearly DV organization of postmitotic neurons
appears to be conserved across widely divergent
vertebrate phyla from fish to mammals and it
represents the developmental ground plan fromwhich different CPG modules in the spinal cordand hindbrain emerge (Goulding, 2009; Grillner
and Jessell, 2009).
The molecular programs that control the acquisition of DV positional identity have been exploredin some detail. In short, the graded activity ofseveral signaling molecules, sonic hedgehog (Shh)and TGF-like bone morphogenic proteins (BMPs),
retinoic acid (RA), and Wnts, initiate and orchestrate the DV patterning of dividing progenitors(Chiang et al., 1996; Liem et al., 1997; Timmer
et al., 2002 and reviewed in Jessell, 2000) in partby establishing dorsoventrally restricted domainsof patterning factors, including the Pax3/7, Pax6,
Nkx6-1, and Nkx2-2 transcription factors, within
the ventricular zone of the neural tube. These
patterning factors function instructively in a combinatorial manner to subdivide the neural tube into
11 progenitor zones (Goulding, 2009). As noted
previously, this developmental program operates
in all vertebrate embryos to activate the expressionof unique sets of homeodomain (HD) and basic
helix-loop-helix (bHLH) transcription factors
in the postmitotic neurons that arise from these
11 progenitor domains (Goulding, 2009; Fig. 1a).
Six classes of early born interneurons, thedI1–dI6 interneurons, are generated in the dorsal
alar plate. Two additional late born populationsof dorsal neurons have been identified that areprevalent in birds and mammals, the dILA anddILB (Gross et al., 2002; Mizuguchi et al., 2006;
Muller et al., 2002; Wildner et al., 2006). Thesecells are similar in molecular composition to dI4
and dI5 interneurons, and it is suspected thatthey represent an evolutionary expansion of

the early born dI4 and dI5 populations in terrestrial vertebrates (Gross et al., 2002; Mizuguchi
et al., 2006; Muller et al., 2002; Wildner et al.,2006). Although the majority of dorsal cell types
differentiate as sensory interneurons and sensory-relay neurons, some appear to make cellular
contributions to the locomotor CPG machinery. However, most of the cell types within thelocomotor CPG are derived from six generic neuronal classes that develop from basal plate progenitors: motoneurons and V0, V1, V2a, V2b,
and V3 interneurons (Goulding, 2009; Jessell,
2000). Derivatives of these ventral classes ofinterneurons settle in regions of the spinal
cord that are known to contain locomotor CPG
networks in quadrupedal mammals (Goulding,2009; Kiehn, 2006). Postmitotic populations of
V0, V1, V2a, V2b, and V3 interneurons aremarked by the expression of the transcription
factors Evx1/2, En1, Chx10, Gata2/3, and Sim1,
respectively (Al-Mosawie et al., 2007; Croneet al., 2008; Karunaratne et al., 2002; Kimura
et al., 2006; Lundfald et al., 2007; Moran-Rivard
et al., 2001; Peng et al., 2007; Saueressig et al.,
1999; Zhang et al., 2008; reviewed in Goulding,2009; Jessell, 2000; Fig. 1a and b). dI6 neurons
that arise more dorsally express Lbx1, Pax2, andLhx1 and settle in lamina VIII (Gross et al.,
2002; Muller et al., 2002). These cells are also
thought to contribute to the premotor circuitry(Fig. 1c). The hindbrain displays a similar DV
organization of early neuronal cell populations.
Of the 14 progenitor domains described in thehindbrain, 10 are equivalent to the spinal progenitor domains and they generate cell types that sharemany features with their spinal cord counterparts(Gray, 2008). Antero-posterior differentiation
events that are orchestrated by the homeobox classof HD transcription factors are of primary importance for the development of hindbrain nuclei thatarise from the same embryonic population.
Embedded in these nuclei are circuits that control
vital autonomic functions such as respiration, as
well as mastication, audition, and the supraspinalcontrol of locomotion (Dubreuil et al., 2009;

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23

Fujiyama et al., 2009; Pagliardini et al., 2008;
Storm et al., 2009). In the next section, we willsummarize what is currently known about theearly development of the spinal motor circuitry.

Generating ventral interneurons: instructiverole of progenitor domain-specifictranscription factors
As noted previously, each of the early DV progenitor domains expresses a unique combinationof transcription factors that play an instructiverole in generating V0, V1, V2a, V2b, and V3
interneurons (Fig. 1). Changes in the expressionof these factors within each progenitor domain
lead to major specification defects in their neuronal progeny. For instance, the HD transcription
factor Dbx1 is an essential factor for the specification of V0 interneurons. In Dbx1 mutant mice,neural progenitors that normally give rise to V0
interneurons are respecified to produce V1 anddI6 interneurons (Lanuza et al., 2004; Pierani
et al., 2001). Ablation of Nkx6-2, a homeobox
transcription factor specific to the V1 progenitor
domain, reportedly increases V0 interneuron
numbers at the expense of V1 interneurons(Vallstedt et al., 2001). Further ventrally, Nkx22 regulates V3 progenitor identity. In Nkx22 mutants, these cells undergo a ventral-to-dorsaltransformation in fate, thereby generating motoneurons at the expense of V3 interneurons(Briscoe et al., 1999). These findings support theidea that transcription factors present in adjacentprogenitor domains suppress the differentiation
programs that operate in the progenitor domains
they abut. They do so by cross-repressive interactions that position the boundaries between
adjacent domains. Consequently, loss of one factor often results in the expansion of an adjacentprogenitor domain and the cell types that arise
from that domain. Other transcription factors
are expressed in more than one progenitor
domain, where they are required for the generation of several neuron classes. Nkx6-1 has beenshown to be important for the generation of bothV2 interneurons and motoneurons (Sander et al.,2000). Inactivation of Pax6, in addition to causing
a loss of V1 interneurons, also alters the generation of branchiomotoneurons in the hindbrain(Burrill et al., 1997; Ericson et al., 1997; Sapir
et al., 2004).
The presence of several transcription factors
within each progenitor domain allows further subspecification of domains characterized by theexpression of Nkx6-1 or Pax6. The V2 progenitor
domain, for example, is defined by coexpressionof the forkhead transcription factor Foxn4 andthe bHLH transcription factor Ascl1. Interestingly, the V2 progenitor domain, unlike othersin the spinal cord, gives rise to two intermingled
but molecularly distinct classes, termed V2a andV2b interneurons. Foxn4 and Ascl1 are criticalfor establishing V2a and V2b cell fate (Del Barrio
et al., 2007; Kimura et al., 2008; Li et al., 2005;
Peng et al., 2007). In V2 progenitors, Foxn4
induces expression of Ascl1 and the Notch ligand
Dll4, which in turn activates Notch signaling inadjacent progenitors. Active Notch signaling inprogenitors undergoing a final cell division leads
to the generation of V2b interneurons, whereas
an absence of Notch signaling in progenitors
results in differentiation into V2a interneurons(Del Barrio et al., 2007; Kimura et al., 2008; Liet al., 2005; Peng et al., 2007). A similar mechanism involving Notch signaling has been implicated in the generation of dILA and dILBinterneurons in the dorsal spinal cord (Mizuguchi
et al., 2006; Wildner et al., 2006).

Generating diversity: postmitotic transcription
factors in interneuron differentiation
Neurons upon exiting the cell cycle migrate into
the mantle zone. During this transition, their transcription factor profile changes and they begin to
express factors exclusive to postmitotic neurons.
These changes in gene expression mark the acquisition of cell type-specific properties such as cell

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24

body position in the cord, neurotransmitter phenotype, axonal projection pattern, and synaptic
target selection. Many V0 interneurons are commissural neurons of mixed neurotransmitter
phenotypes that extend axons rostrally in theembryonic cord (Moran-Rivard et al., 2001;
Pierani et al., 2001). The postmitotic transcription
factor Evx1 is specific to V0V neurons that are predominantly glutamatergic but is not expressedin dorsally derived GABA/glycinergic V0D
interneurons (G. Lanuza and M. Goulding, unpublished). In Evx1 mutants, a large fraction of
V0V interneurons acquire the fate of V1 interneurons and extend axons into the ventrolateralfuniculus instead of across the ventral midline.
Consequently, Evx1 functions as a postmitotic
determinant of V0V interneuron identity (Lanuza
et al., 2004; Moran-Rivard et al., 2001). Other postmitotic neuron populations are also known todiverge into smaller subpopulations of neurons.
This diversification probably represents the essential developmental and evolutionary changes
needed to meet the increased requirements higher
vertebrates have with respect to complex locomotor movements and respiration.
The further specialization that occurs withinthese generic populations of interneurons is perhaps best exemplified by the V1 interneuron class.
V1 interneurons are marked by their transient
expression of the HD protein En1, although En1
has no discernable impact on V1 generic identity
or diversification (Sapir et al., 2004; Saueressig
et al., 1999). Recently, it has been shown that En1
expressing V1 cells differentiate into Renshaw cells
and Ia inhibitory interneurons (Alvarez et al.,2005; Sapir et al., 2004). Renshaw cells receive
strong excitatory innervation from motoneuron
axon collaterals and in turn inhibit motoneurons,
as well as Ia inhibitory interneurons, via synapses
that use both glycine and GABA. Remarkably,
Renshaw cells and Ia inhibitory interneurons
represent less than 25% of all V1-derived cells,
demonstrating that there are additional uncharacterized V1-derived cell types. Current efforts tomolecularly map these V1 subpopulations haverelied on a combination of expression screens andcandidate gene approaches. We have used genetic
marking of En1 expressing V1 interneurons topurify these cells from E12.5 spinal cords. Microarray analysis of mRNA isolated from thesecells has led to the identification of a number ofgenes that are enriched in this population(T. Hendricks and M. Goulding, unpublished).
Many of them appear to be expressed in subsets
of V1 interneurons and may therefore subdivide
the V1 population into functional units (Table 1).
Several transcription factors exhibit more selective patterns of expression. Pitx2, for example, isdifferentially expressed by a small subpopulationof ipsilaterally projecting V0 interneurons (V0C
interneurons) that form cholinergic synaptic connections on motoneurons (Zagoraiou et al., 2009).
In V1 interneurons, MafB is selectively expressed
in Renshaw cells but is absent from other V1 cells(T. Hendricks, F. Stam, M. Goulding, unpublished). Glutamatergic V2a and GABA/glycinergic
V2b interneuron populations are marked by theirdifferential expression of Chx10 and Gata2/3,
respectively (Al-Mosawie et al., 2007; Lundfald
et al., 2007). Chx10 and Gata2/3 appear not to govern the neurotransmitter identity or axonal morphology of the V2a and V2b interneurons;
however, inactivation of Gata2 in mice suggests
this transcription factor has a role in maintaining
postmitotic V2 cell identity (Zhou et al., 2000).
V3 interneurons that express the PAS-bHLH factor Sim1 can be subdivided into dorsal and ventral
populations (Zhang et al., 2008). However, none of
Table 1. Genes enriched in V1 and V3 interneuron
populations at E12.5

V1 enrichedgenes
V3 enriched
genes

Transcription
factors

Other

En1, Foxd3, MafB
Otx-like, Hoxa3,
Cux2
Neurog3, Olig3,
Sim1
Nkx2-2, Uncx

Nrxn3, Chmp1b,
Nrip3
Kcnk13, Scn2a1
Gtpbp6, Plekhh1,
Adam11
Sema5a, Slc5a6

Author's personal copy
25

the V3-enriched genes that have been identified
to date delineate these two populations (Table 1),
even though both populations do exhibit differences in their axonal morphology and electrophysiological properties (E. Geiman, Y. Zhang,and M. Goulding, unpublished). Although efforts
to map the genetic pathways that establish interneuron diversity in the developing spinal cord arestill in their infancy, they are likely to provide aclearer picture of the interneuronal compositionof the spinal motor circuitry. This will in turn facilitate further anatomical, physiological, and functional studies of specialized interneuron cell types
so as to determine their contributions to motor
control.

Genetically defined interneuronal populations
that shape the locomotor rhythm
The identification of genetic markers for different
spinal interneuronal populations has laid the foundation for functional studies aimed at assessing
the contribution that genetically defined cell types
make to the CPG networks controlling locomotor or respiratory rhythmogenesis (Chapter 3
in this book). These efforts have been centeredon broad interneuron classes including the V0,
V1, V2a, and V3 interneurons (Stepien and Arber,2008); however, attention is now turning tosubtypes within these larger populations, such as
the V0C interneurons (Zagoraiou et al., 2009).
In characterizing and determining the function ofgenetically defined interneuron populations inthe spinal cord, the following themes have emerged.

Modular nature of the mammalian
locomotor CPG
Interneurons that make up each of the early
generic populations in the embryonic cord typicallyshare a number of common characteristics, forexample, morphology of the primary axonal process or neurotransmitter phenotype. Moreover,this subdivision of embryonic interneurons into
discrete anatomical groupings has led to the suggestion that the spinal motor circuitry comprises
functional modules that are made up of geneticallydefined populations of interneurons. This proposition has been confirmed by genetic studies in mice
showing that particular interneuron cell types
control discrete aspects of the locomotor output.
Loss-of-function manipulations that target aparticular interneuron population modify specificaspects of the locomotor rhythm and pattern, andleave others unchanged. Some interneuron cell
types regulate left–right alternation, while others
determine the speed of stepping movements orcontrol the balance between motor outputs on
the left and right sides of the cord.
In Dbx1 null mutants, where the V0 population
is markedly depleted (Lanuza et al., 2004; Pierani
et al., 2001), intermittent episodes of left–rightsynchronous activity are seen in the locomoting
cord (Fig. 2c). This deficit is selective, with ipsilateral flexor–extensor alternating activity being
maintained and no major changes occurring in thestep cycle period. In addition to identifying a neuronal population that is required for proper
left–right coordination during locomotion, thisstudy provided evidence that the locomotor CPG
has a modular composition with respect to functional control by genetically defined populationsof interneurons. More recently, deletion experiments targeting V2a excitatory neurons haverevealed an interesting twist to the story, byidentifying V2a interneurons as an ipsilateral component of the commissural pathways that secureleft–right alternation (Crone et al., 2008; Figs. 1cand 2c). Once again the effects of deleting theV2a interneurons are primarily restricted toleft–right stepping behaviors. Moreover, left–right
coordination is only altered when mice lacking
V2a interneurons step quickly or trot, whichsuggests that the V2a interneurons are primarily
active at higher stepping speeds (Crone et al.,2009). The observation that V2a ipsilateral
neurons project onto V0 commissural interneurons
confirms that the V2a cells are part of the CPG

Author's personal copy
26(a) iFlexor

iExtensor

(b)
WT

F-F

cF
F-E

iF
iE

cFlexor

(c)

V1

F-F

V3

F-F

cF
F-E

iF

F-E

iE
V0

F-F

V2a

F-F

cF
iF

F-E

F-E

iE
Fig. 2. Fictive locomotor activities in mice defective in ventral interneurons. (a) Schematic of an isolated spinal cord from a neonate
mouse. When bathed with a cocktail of serotonin and N-methyl-D-aspartate, a fictive locomotor rhythm with left–right andflexor–extensor coordinations can be recorded from flexor L2 and extensor L5 lumbar ventral roots. (b) Schematic of locomotorlike activity observed in wild-type spinal cords. Bursts of activity are illustrated by rectangles, polar plots represent coordination
between two antagonistic roots. Note the strict left–right and flexor–extensor alternations. (c) Schematic of locomotor activity
recorded from spinal cords of different mutant mice. Defects in V1 or V3 interneurons disturb the locomotor step cycle resulting
in a slower step cycle period or an unbalanced locomotor rhythm without affecting bilateral and flexor–extensor coordination.
The absence, or ablation, of V0 and V2a interneurons results in a partial impairment of bilateral alternation, which is manifested
by episodes of left–right cocontraction. c, controlateral; i, ipsilateral; F, flexor; E, extensor; WT, wild type.

module that controls left–right coordination. They
most likely provide rhythmic excitatory drive tocommissural inhibitory neurons that generate
reciprocating left–right flexor and left–right extensor activity.
When a second population of excitatory interneurons, the commissural Sim1þ V3 interneurons,
were targeted, near-normal patterns of left–rightand flexor–extensor alternation were retained,even though there was a marked reduction in thecoherency of the locomotor rhythm. Cords inwhich Sim1þ V3 interneurons were silenced did,
however, exhibit a pronounced imbalance in the
duration of motor bursts on each side of the cord.
This typically involved bursts from one L2 ventral
root being prolonged, while those in the otherroot were truncated (Zhang et al., 2008 and Fig. 2c).
V3 excitatory pathways are therefore predisposed
to balancing the locomotor output between
both sides of the cord, as opposed to establishingalternating patterns of left–right or flexor–extensor
motor activity. Further evidence for the modularorganization of the locomotor CPG network hascome from experiments analyzing the role of theV1 class of interneurons. When V1 interneurons
are depleted in the embryonic spinal cord, marked
changes occur in the speed of locomotor rhythm(Gosgnach et al., 2006; Fig. 2). In these mice,
left–right stepping and flexor–extensor reciprocation is normal, suggesting the primary role ofthese cells in the walking CPG is to regulate burst
termination and initiation during each step cycle.

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27

Functional redundancy and complexity
in the locomotor CPG
Experiments demonstrating that the deletionof V0 interneurons produces only partial deficits
in left–right coordination reveal a degree of
redundancy in the commissural pathways that
secure left–right alternation. While it seems likelythat additional interneuron cell types help
coordinating left–right alternation, their identity
is not known. The recent demonstration that
Netrin-1 mutant cords display a switch fromalternating left–right to synchronous left–right
locomotor activity supports the idea that multiplecommissural interneurons control left–right stepping (Rabe et al., 2009). Moreover, Rabe et al.
(2009) also noted that the axons of glutamatergic
V3 commissural neurons still cross the ventral
midline when netrin-1 is absent, demonstratingthat this excitatory commissural pathway cannot
secure left–right alternation. We also know thatwhen V3 interneurons are inactivated, left–rightalternation is not compromised (Zhang et al.,2008), arguing the contribution this pathway
makes to left–right stepping is at best minor.
The observation that misspecification of a secondclass of putative excitatory commissural neurons,
the Evx1þ V0V cells, also leaves left–right stepping behaviors intact (Moran-Rivard et al., 2001;
Lanuza et al., 2004) leads us to believe that excitatory commissural pathways in the cord do nothave essential roles in controlling left–right alternation. In support of this idea, when V3 transmission is abrogated in the Dbx1/ mutant cord, we
saw no further degradation of left–right alternation over and above that previously described
for the Dbx1/ spinal cord (Lanuza et al., 2004;
Y. Zhang and M. Goulding, unpublished).
A second example of potential redundancy inthe interneuron pathways that control the locomotor output has emerged from efforts to understand how the locomotor CPG produces analternating pattern of flexor–extensor activity.
Multiple manipulations that abrogate ipsilateral
inhibitory V1 interneurons function leaveflexor–extensor coordination intact in the isolated
CPG or in adult mice (Gosgnach et al., 2006).
This was somewhat surprising in so far as an
earlier anatomical study had shown that V1 interneurons give rise to cells with the features of Ia
disynaptic inhibitory interneurons, a key component of the reciprocal inhibition in the stretch
reflex involving flexor–extensor alternation(Alvarez et al., 2005). Experiments undertakenin the lab of Eric Frank examined the integrityof disynaptic inhibitory pathways in the spinal
cord of mice lacking V1 interneurons. These
analyses revealed that the disynaptic inhibitory
pathway between quadriceps sensory afferents
and posterior biceps and semitendinosus motor
neurons is still operational in Pax6 mutant
animals (Wang et al., 2008) and in mice thatlack V1 neurotransmission (Z. Wang, J. Zhang,
M. Goulding, and E. Frank, unpublished). It now
seems that an additional population of inhibitory interneurons act in concert with the V1
population to control this aspect of the motor output (J. Zhang, G. Lanuza, and M. Goulding,unpublished).
With the discovery of additional interneuron
markers such as those described in Table 1, it is
now becoming apparent that the interneuron
composition of the spinal cord is quite complex.
As noted previously, three-quarters of the V1
interneurons remain unidentified (Alvarez et al.,2005; Sapir et al., 2004), and even less is known
about the identity and function of cell types thatarise from other embryonic interneuron classes.
Using functional criteria to classify spinal interneurons may be problematic, given the likelihoodthat several interneuron cell types may together
regulate a particular motor behavior. Consequently, the loss of one cell population may notlead to pronounced behavioral deficits, especiallywhen one is assaying rudimentary motor patterns
such as those produced in the isolated spinal
cord or in hindbrain slices. This is borne out by
experiments that demonstrate the slowing of thelocomotor rhythm caused by V1 depletion is
not reproduced when Renshaw cells alone areAuthor's personal copy
28

silenced (Myers et al., 2005). As such, the complex
organization of spinal motor circuitry in higher
vertebrates may prove problematic for futurefunctional studies due to redundancy and compensation by other spinal interneuron populations
that have similar or overlapping functions.

Comparison with respiratory CPGs
Many of the embryonic interneuron populations
that are present in the spinal cord form longitudinal columns that extend into the hindbrain
where they contribute to other motor networks
including those controlling respiration. Furthermore, many interneuron cell types that are known
to control locomotion appear to have essential
roles in respiration, as inactivating them often
results in perinatal lethality. For example, inhibitory cell types that are derived from the V1
and V2b interneuron populations also have essential roles in establishing a normal respiratory
rhythm (J. M. Zhang, J. Ramirez, M. Goulding,
J. Feldman, unpublished). In Lbx1 mutant mice,the altered organization of some hindbrain excitatory nuclei results in an immature respiratory
rhythm at birth (Pagliardini et al., 2008). Amongthese Lbx1-expressing cells are Tlx3þ glutamatergic neurons that are important for respiration (Cheng et al., 2004). Together, these studies
give heft to the argument that the respiratory
and locomotor CPGs are formed from equivalentneuronal substrates in the hindbrain and spinal
cord, respectively. It also highlights the potential
insights that can be gained by comparative
functional studies between the locomotor andrespiratory CPGs with respect to the cellular organization of motor networks in the CNS. Other
groups have taken the approach of probing thesecircuits using genes that are expressed either inthe hindbrain rhombomeres from which these
CPGs develop or in populations of putative respiratory interneurons (Abbott et al., 2009; Dubreuil
et al., 2009; Gray et al., 2001; McKay andFeldman, 2008; Tan et al., 2008; Thoby-Brisson

et al., 2009). For example, studies analyzing
Phox2b mutant mice have revealed a key rolefor parafacial/retrotrapezoid nucleus in chemosensitivity (Dubreuil et al., 2009). Future experiments to genetically dissect these respiratory
circuits are likely to use combinatorial methodologies to selectively target particular interneuron
populations within a single rhombomere or regionof the medulla, some of which are described in thenext section.

New genetic approaches for studying
motor circuits in the spinal cord
The elaboration of a genetic classification
scheme for the interneuron cell types involved inspinal motor control has opened up new routes
for manipulating the spinal motor system anddetermining how specific motor behaviors are
generated. The approaches used so far involve
deleting or inactivating broad interneuron classes
and assessing how this affects network activity(Table 2). This is usually achieved by generatingtransgenic animals in which a recombination
event such as Cre-mediated excision is used toexpress an effector molecule in a particular groupof neurons (Fig. 3). Class-specific promoters incombination with site-specific recombinases allow
ablating, silencing, or activating a population.
Most of what we know about the function ofgenetically-defined spinal interneuron classes
comes from in vitro analysis using the isolated spinal cord preparation, in which fictive locomotor
activity is determined by extracellular ventral
root recordings (Crone et al., 2008; Gosgnach
et al., 2006; Lanuza et al., 2004; Zhang et al.,2008). While such studies have provided a broadfunctional overview of the cellular makeup of
the spinal locomotor CPG, the finer aspectsof motor control have been largely ignored dueto the limitations of these approaches. Nonetheless an arsenal of second-generation tools is
being assembled that builds on the lessons learnt
from these cruder studies.

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29
Table 2. Genetic tools for studying locomotion and respiration
MethodTarget of action

EffectInducerCaveats

Advantages

Knock out
(a–f)

Gene inactivation

Loss of proteinfunction– Often perinatal lethal

 Many mouse mutants
available Can affect multiple celltypes
 Respecification of cells
 Adaptation possible
DTA (b, g)

Inhibition of RNAtranslation

Cell death– Presynaptic cells may
be affected by loss of
their target
 Adaptation possible
 Not neuron specific Highly efficient method
to remove one defined
population
 Ablation is easy tomonitor

TeNT (h)Cleavage of
VAMP-2 synaptic
proteinSilencing (synaptictransmission)–

 Adaptation possible

 Targeted cells remain in
the circuit Affects only neuronal
cells

AlstR (b, h)Activation of
GIRK channel

Silencing(hyperpolarization)

Alst(peptide) Cannot be injected
systemically (does notcross BBB) Desensitization after
prolonged exposure Inducible/reversible
within minutes
 Local effect byintrathecal application of
Alst Allows analysis of shortterm effects
 Allows analysis in adultmice

ChR2 (i)Expression of lightsensitive cation
channelActivation(depolarization)

Blue
light

 High expression levels
required Few transgenic mice
successfully generated
 Light penetration into
deep tissue

 Inducible/reversible
within milliseconds
 Local excitation possible Allows analysis of shortterm effects

 Silencing difficult toassess

DTA, Diphtheria Toxin A subunit; DTR, Diphtheria Toxin Receptor; TeNT, Tetanus toxin light chain; AlstR, Allatostatin Receptor; Alst, Allatostatin; ChR2, Channelrhodopsin 2; BBB, blood–brain barrier; GIRK, G-protein-coupled inwardly rectifying potassium; VAMP-2, vesicle-associatedmembrane protein 2 (synaptobrevin2). a, Lanuza et al., 2004; b, Gosgnach et al., 2006; c, Pagliardini et al., 2008; d, Rose et al., 2009; e, Dubreuil
et al., 2009; f, Wallen-Mackenzie et al., 2006; g, Crone et al., 2008; h, Zhang et al., 2008; i, Hagglund et al., 2010.

Defective cell generation or cell ablation
One of the routes that has been successfully used
to analyze motor circuits in the hindbrain and spinal cord makes use of mouse mutants in whichtranscription factors or signaling molecules thatspecify neuronal identity and connectivity during
development are inactivated (Fig. 1). Althoughthese factors are required for the generation of
distinct populations of spinal interneurons, they
are often expressed elsewhere in the developing
mouse, which usually leads to perinatal lethality

Author's personal copy
30
On
(a)

Promoter 1

Cre
Off

Promoter 2
Recombination
event
Promoter 2

Stop

Gene of interest

On
Gene of interest(b)Class specific Cre expression
molecular
turns on genemarkerof interest

Cell lineage
permanently
labeledFig. 3. Cre-dependent manipulation of defined neuronal
populations. (a) Two different alleles are used in combination to
genetically manipulate defined populations of cells. First, the sitespecific Cre recombinase is expressed from a promoter(1) specific to one neuronal class. A second allele is used toexpress a gene of interest from a promoter (2) that is, in general,
constitutive and broadly active but can also be neuronal or cell
type specific. Transcriptional STOP sequences flanked by loxPsites (green triangles) are interposed between the promoter andthe coding sequence such that Cre-mediated recombination ofloxP sites and excision of the STOP cassette is required to express
the gene of interest. This strategy can be used to express areporter gene such as GFP for morphological and axonal
characterization, and fate mapping of the genetic lineage or to
express an effector gene like TeNT or DTA, resulting in cellsilencing or ablation for functional studies. (b) Schematic picture
showing genetic lineage labeling in the spinal cord. The promoter(1) of a molecular marker defining a neuronal class (yellow dots)
is used to drive Cre expression in the same cells (green dots).
Recombination induces expression of the gene of interest (bluecircles), which is dependent on promoter (2). At later stages ofdevelopment recombined cells have migrated to settle in their
final location. Although these cells no longer express themolecular marker nor Cre, they are permanently labeled (bluecircles). (For interpretation of the references to color in this figurelegend, the reader is referred to the Web version of this chapter.)when these genes are inactivated. As locomotorlike and respiratory activity can be recorded inneonate mice using the isolated hindbrain–spinal
cord preparation, such mutant animals have

proved useful for studying these behaviors(Gosgnach et al., 2006; Lanuza et al., 2004; Myerset al., 2005). An additional issue is the broad
expression of many factors within the spinal cordor hindbrain, as removing these factors typicallyaffects large neuronal populations or results incomplex cell fate changes that make interpretation of any observed motor deficit difficult. Whileapproaches that selectively ablate a population
may avoid some of the complex cell fate changes
associated with inactivating developmental control genes, in many instances the effects can stillbe quite widespread. Expression of the diphtheria
toxin A protein (DTA) in molecularly definedcells has been used to selectively eliminate entire
classes of interneurons. Examples come from V1
and V2a interneurons, which were successfully
ablated using the Rosa-stop-DTA ablator allele
and a Chx10-stop-DTA allele, respectively (Croneet al., 2008; Gosgnach et al., 2006). However, useof this strategy is limited as molecular markers ofspinal interneurons are often also expressed in
non-neuronal tissues, which causes additional
phenotypes in mice genetically modified to kill
one population. For this reason, cell ablation
approaches are often not possible due to collateral tissue damage that results in embryoniclethality. Examples of this are the V1 and V3
interneuron-specific factors En1 and Sim1 thatare expressed in muscle cells or the V2b marker
Gata3, which is expressed in the placenta andin endothelial cells (Chen and Johnson, 2002;
Coumailleau and Duprez, 2009; Ng et al., 1994).
Using a neuronal driver such as Eno2-StopDTA allele restricts ablation to neurons, thereby
avoiding the effects in nonneural tissue(Kobayakawa et al., 2007).

Manipulating cell activity
A different approach to assessing neuronal function involves silencing or blocking neurotransmission in selected neurons and analyzing the effects
on particular behaviors. Synaptic transmission canAuthor's personal copy
31

be abolished in a particular spinal cell class byexpression of the light chain of the tetanus toxin(TeNT; Yamamoto et al., 2003; Zhang et al.,2008). TeNT prevents neurotransmitter releaseat the synapse, thereby blocking the activation
or inhibition of postsynaptic target neurons. Thisapproach has the advantage that the geneticallytargeted cells are still present. However, expression of TeNT is difficult to assess functionally.
While VAMP2 expression can be used as a readout of TeNT activity, electrophysiological studies
provide the most compelling validation of TeNT
activity. For example, experiments recently
undertaken in the cerebellum revealed that TeNT
expression in cerebellar granule cells disrupts
synaptic transmission to Purkinje cells (Kimet al., 2009). A different strategy for acute silencing involves expressing the insect allatostatin
G-coupled protein receptor, AlstR (Gosgnach
et al., 2006; Zhang et al., 2008). Upon allatostatin
ligand binding, this receptor activates inwardly
rectifying Kþ channels (GIRK channels) leading
to transient hyperpolarization and decreased
excitability of the neurons in which it is expressed(Lechner et al., 2002; Tan et al., 2008). Neuronalsignaling can also be altered by inactivating critical components of the neurotransmission machinery. This approach was successfully used byZagoraiou et al. (2009) since they specificallyknockout choline acetyltransferase (ChAT) in
Pitx2þ V0C cells thus preventing synaptic transmission from this particular neuronal population(Zagoraiou et al., 2009). Finally, genetic systems
that can be used to stimulate or increase the excitability of defined neurons are being developed inmice, and in zebrafish (Armbruster et al., 2007;
Wyart et al., 2009). Neurons can be remotely
activated by the expression of channelrhodopsin
2 protein (ChR2), a microbial light sensitive
monovalent cation channel that allows entranceof Naþ ions in the cell following exposition
to blue light (Luo et al., 2008; Nagel et al.,2003). Thus, illumination of ChR2 expressing
neurons leads to their depolarization and activation (Boyden et al., 2005). ChR2 expression wasrecently targeted to VGlut2 neurons in the hindbrain and the spinal cord and illumination ofeach region was seen to induce and maintainlocomotor-likeactivity in the spinal preparation.
This finding confirms that glutamatergic neurons
have a key role in elaborating the locomotor
rhythm (Hagglund et al., 2010). Approaches using“designer” muscarinic GPCR receptors that canbe exclusively activated by synthetic ligands provide additional avenues for selectively stimulating
particular interneuron cell types (Armbruster
et al., 2007).

Case history: genetic manipulations
involving V1 interneurons
Many of the aforementioned techniques forgenetically manipulating neurons have now beentrialed, each of which has advantages and disadvantages. However, by judiciously tailoring
the approach used, one can undertake elegant
manipulations of these spinal motor networks.
Loss of cells is readily quantifiable, whereassilencing activity requires electrophysiological
techniques to confirm reduced activity in thetargeted neurons. In the Goulding lab, several
genetic approaches have been used to remove
the V1 class from the spinal circuits. These
include developmental deficits in V1 specification(Pax6/), V1-specific cell ablation (En1-DTA),
or blocking transmission/silencing (En1-TeNT
and En1-AlstR, respectively). All four methods
cause a strong slowing of the locomotor-likerhythm while preserving other aspects of the
motor output: rhythmogenesis, left–right andflexor–extensor alternation (Fig. 2c). The
phenotypes of the Pax6 mutant and En1-DTA
mice are very similar, whereas En1-TeNT mice
produce a comparatively slower motor rhythm.
While cell “silencing” with AlstR produces a slow
rhythm when compared to wild-type animals, therhythm is significantly faster than that observedin En1-TeNT cords. This, in all likelihood,
represents partial silencing/inactivation of the V1

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population. The rhythm in the En1-AlstR cords
also returns to its normal frequency approximately 15 min after exposure to allatostatin,
which probably reflects desensitization of the
GIRK channels following prolonged exposure toallatostatin.

Studies in adult behaving mice
Functional analysis in vitro using the isolated spinal cord preparation usually only allows identification of gross defects in motor activity.
Analyzing complex motor behaviors at adultstages in genetically manipulated mice is often
not possible because of neonatal lethality anddeficits in more anterior regions of the CNS orin nonneuronal structures. The transcription factor Lbx1 for instance, which is important for
specifying dorsal neural fate including the dI6
interneurons, is expressed in the diaphragm(Gross et al., 2000; Storm et al., 2009) and hindbrain, where it is involved in respiratory
rhythmogenesis (Pagliardini et al., 2008). In someinstances, genetic approaches have been applied
to adult animals. Silencing the V3 cells in adultmice by using the AlstR system results in defects
in walking (Zhang et al., 2008). Similar to thefindings from in vitro experiments, the conditionalsilencing of these commissural interneurons inadult mice produced less stable and robust locomotor cycles during walking, thus validating thefictive locomotion analysis performed in the neonatal spinal cord.
Studying locomotor behaviors in adult micecan also be used to refine observations that havebeen made using the isolated spinal cord preparation. In vitro studies of V2a interneurons in the
neonatal cord (Crone et al., 2008) were extended
by adult kinematic analyses showing that V2ainterneurons are needed for left–right stepping at faster gait speeds (Crone et al., 2009).
Acute silencing of lumbar level V3 interneurons
was used to confirm the altered rhythm that
occurs in vitro following TeNT blockade of V3

transmission (Zhang et al., 2008). Finer aspectsof motor control have also been tested in behaving mice. For example, analysis of the V0C population in adult mice unraveled the specificfunction of these neurons during particular locomotor behavior. EMG recordings from gastrocnemius and tibialis anterior muscles, respectively
ankle extensor and flexor, during walking andswimming tasks revealed that in the absence of
V0C neuron activity, the task-dependent modulation of the gastrocnemius muscle was impaired(Zagoraiou et al., 2009). In vitro approaches would
not have revealed these kinds of modulatory
functions, particularly with respect to small neuronal populations.

Looking forward
A next and crucial step for understanding thefunctional contribution these spinal interneuron
classes make to motor control involves acutely
eliminating each of them locally from spinal
networks in the adult. It is yet not clear whether
there are developmental compensatory changes
that occur in spinal neuronal networks whena particular neuronal component is removed
from the circuit as it is being wired. Loss of cells,
especially as circuits are forming, may result in
network rearrangements that compensate for theabsence of these cells. In particular, little is
known about the consequences on growing axons
seeking out for synaptic partners when their target cells are absent. Inducing cell ablation after
the circuits have wired and fully matured may
therefore be critical. An elegant approach is torender mouse cells susceptible to the diphtheria
toxin by specific expression of a simian or humandiphtheria toxin receptor (DTR). Selective cell
death can then be achieved by injecting the diphtheria toxin intraperitoneally at desired time
points. Functional compensation has beenreported in the circuits controlling feeding andnociception using this strategy (Cavanaugh et al.,2009; Luquet et al., 2005).

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Most molecular markers of interneurons being
also expressed in supraspinal neurons or other tissue types, new strategies need to be implemented
where genetic targeting is restricted to neuronalclasses only in the spinal cord. This would reveal
the exclusive role these subpopulations have inshaping motor output during walking, without
affecting the functionality of other tissues/organs,or regions of the nervous system. For this purpose, approaches using intersecting combinationsof expressed genes are promising. This has successfully been done using Cre driver lines in combination with Flp driver lines (Awatramani et al.,2003). Several dual recombinase-response alleles
have now been generated that enable cell labelingor silencing (Farago et al., 2006; Jensen et al.,2008; Kim et al., 2009; Yamamoto et al., 2009).
This strategy could allow dissecting the roles ofgenetically defined cells present in multiple
regions in the rostrocaudal axis. For instance,
silencing of all neurons of the Atoh1 lineage,
including granule cells in the cerebellum and the
dI1 class of spinal cells, results in a robust motor
coordination defect in behaving mutant mice
(Kim et al., 2009). Generating Flp lines specificto either the cerebellum or the spinal cord would
allow one to test the contribution each region
makes to motor control. The intersectional strategy also represents an attractive method for subdividing the current broad neuronal classes into
smaller subsets. Within the V1 interneuron classdefined by expression of En1, only the Renshaw
cells coexpress the Ca2þ binding protein calbindin(Carr et al., 1998; Sapir et al., 2004). Using these
two genes in a combinatorial manner to target
this subset of V1 interneurons would thus allow
to functionally testing the role of this subpopulation. Double recombinase approaches could also
be helpful to unravel the functional contributionof specific cell types related to respiration.
Patterning programs in the hindbrain specify
interneurons that integrate different functional
circuitry according to their antero-posterior localization. Manipulating one interneuronal population in specific rhombomeres would allow one tounravel the roles different classes of hindbrain
interneurons play in respiratory rhythmogenesis.
Together with inducible systems for activationand silencing of neurons, the dual recombinase
approach represents an elegant way to study specific neuronal populations in adult mice. Nevertheless, in addition to generating novel genetic
tools for studying locomotion, more effort needs
to be invested in developing robust and precise
behavioral tests that are able to detect the subtle
locomotor phenotypes when smaller groups ofcells are inactivated in adult mice.

Conclusion
The delineation of the transcriptional code forneuronal cell populations in the ventral spinal
cord and the emergence of several genetic
approaches to manipulate cells of interest have
paved the way for genetically and functionally dissecting the neural circuits controlling locomotion.
The embryonic building blocks that make up these
motor circuits are shared between the networks
controlling respiration and locomotion. Thus,
understanding how neuronal cell populations thatcomprise the hindlimb locomotor CPG areorganized should provide important insights into
how other CPGs function. It is also hoped thatefforts to understand motor behaviors such as
locomotion and respiration will provide some of
the keys for unlocking more complex neuronal
networks.
The studies described in this chapter highlightsome of the fundamental findings that have shaped
our understanding of how interneurons in theventral spinal cord form neuronal circuits thatelaborate coordinated rhythmic motor outputs.
However, there is a high degree of complexity inthe composition of these circuits, with recent
studies revealing the existence of specialized
subpopulations within the generic populations ofinterneurons that emerge from the ventral spinal
cord. Although, significant progress has been madein understanding the coarse function of large cell

Author's personal copy
34

groups in controlling particular aspects of locomotor rhythm and pattern generation, many moreinvestigations are waiting to be undertaken which
elucidate the role of smaller subpopulations in
fine-tuning and shaping locomotor output. Suchefforts will depend heavily on the identificationof specialized neuronal cell types and the development of new methodologies to geneticallytarget and manipulate them.

Acknowledgments
K. S. G. is funded by a Feodor Lynen Fellowshipfrom Alexander von Humboldt Foundation. Research in the Goulding lab is supported by grants fromthe National Institutes of Health (NS031249,
NS031978 and NS037075) and the Christopher andDana Reeve Foundation. We would particularly
like to thank Tim Hendricks, Floor Stam, and othermembers of the Goulding Lab for allowing us to cite
their unpublished findings.

Abbreviations
Alst
AlstR
bHLH
BMPs
ChAT
ChR2
CPGs
DTA
DTR
DV
EMG
GIRK
GPCR
HD
RA
Shh
TeNT
TGF

allatostatin
allatostatin receptor
basic helix-loop-helix
bone morphogenetic proteinscholine acetyltransferase
channelrhodopsin 2
central pattern generators
diphtheria toxin A subunit
diphtheria toxin receptor
dorsoventral
electromyogram
G-protein-coupled inwardly
rectifying potassium channel
G-protein-coupled receptor
homeodomain
retinoic acidsonic hedgehog
light chain of the tetanus toxintransforming growth factor

References
Abbott, S. B., Stornetta, R. L., Fortuna, M. G., Depuy, S. D.,
West, G. H., Harris, T. E., et al. (2009). Photostimulationof retrotrapezoid nucleus phox2b-expressing neurons
in vivo produces long-lasting activation of breathing in rats.
The Journal of Neuroscience, 29(18), 5806–5819.
Al-Mosawie, A., Wilson, J. M., & Brownstone, R. M. (2007).
Heterogeneity of V2-derived interneurons in the adultmouse spinal cord. The European Journal of Neuroscience,
26(11), 3003–3015.
Alvarez, F. J., Jonas, P. C., Sapir, T., Hartley, R.,
Berrocal, M. C., Geiman, E. J., et al. (2005). Postnatal
phenotype and localization of spinal cord V1 derived
interneurons. The Journal of Comparative Neurology, 493(2), 177–192.
Armbruster, B. N., Li, X., Pausch, M. H., Herlitze, S., &Roth, B. L. (2007). Evolving the lock to fit the key to createa family of G protein-coupled receptors potently activatedby an inert ligand. Proceedings of the National Academy of
Sciences of the United States of America, 104(12), 5163–5168.
Awatramani, R., Soriano, P., Rodriguez, C., Mai, J. J., &Dymecki, S. M. (2003). Cryptic boundaries in roof plateand choroid plexus identified by intersectional gene activation. Nature Genetics, 35(1), 70–75.
Ballion, B., Morin, D., & Viala, D. (2001). Forelimb locomotor
generators and quadrupedal locomotion in the neonatal rat.
The European Journal of Neuroscience, 14(10), 1727–1738.
Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G., &Deisseroth, K. (2005). Millisecond-timescale, geneticallytargeted optical control of neural activity. Nature Neuroscience, 8(9), 1263–1268.
Bracci, E., Ballerini, L., & Nistri, A. (1996). Localization ofrhythmogenic networks responsible for spontaneous bursts
induced by strychnine and bicuculline in the rat isolated spinal cord. The Journal of Neuroscience, 16(21), 7063–7076.
Briscoe, J., Sussel, L., Serup, P., Hartigan-O'Connor, D.,
Jessell, T. M., Rubenstein, J. L., et al. (1999). Homeobox geneNkx2.2 and specification of neuronal identity by graded Sonic
hedgehog signalling. Nature, 398(6728), 622–627.
Burrill, J. D., Moran, L., Goulding, M. D., & Saueressig, H.
(1997). PAX2 is expressed in multiple spinal cord interneurons, including a population of EN1þ interneurons that
require PAX6 for their development. Development, 124(22), 4493–4503.
Carr, P. A., Alvarez, F. J., Leman, E. A., & Fyffe, R. E. (1998).
Calbindin D28k expression in immunohistochemically
identified Renshaw cells. NeuroReport, 9(11), 2657–2661.
Cavanaugh, D. J., Lee, H., Lo, L., Shields, S. D., Zylka, M. J.,
Basbaum, A. I., et al. (2009). Distinct subsets of unmyelinated primary sensory fibers mediate behavioral responses
to noxious thermal and mechanical stimuli. Proceedings of
the National Academy of Sciences of the United States ofAmerica, 106(22), 9075–9080.

Author's personal copy
35
Cazalets, J. R., Borde, M., & Clarac, F. (1995). Localizationand organization of the central pattern generator for
hindlimb locomotion in newborn rat. The Journal of Neuroscience, 15(7 Pt 1), 4943–4951.
Chen, H., & Johnson, R. L. (2002). Interactions between
dorsal-ventral patterning genes lmx1b, engrailed-1 andwnt-7a in the vertebrate limb. The International Journal of
Developmental Biology, 46(7), 937–941.
Cheng, L., Arata, A., Mizuguchi, R., Qian, Y., Karunaratne, A.,
Gray, P. A., et al. (2004). Tlx3 and Tlx1 are post-mitotic
selector genes determining glutamatergic over GABAergic
cell fates. Nature Neuroscience, 7(5), 510–517.
Chiang, C., Litingtung, Y., Lee, E., Young, K. E.,
Corden, J. L., Westphal, H., et al. (1996). Cyclopia anddefective axial patterning in mice lacking Sonic hedgehog
gene function. Nature, 383(6599), 407–413.
Coumailleau, P., & Duprez, D. (2009). Sim1 and Sim2 expression during chick and mouse limb development. The
International Journal of Developmental Biology, 53(1), 149–157.
Crone, S. A., Quinlan, K. A., Zagoraiou, L., Droho, S.,
Restrepo, C. E., Lundfald, L., et al. (2008). Genetic ablationof V2a ipsilateral interneurons disrupts left–right locomotor
coordination in mammalian spinal cord. Neuron, 60(1), 70–83.
Crone, S. A., Zhong, G., Harris-Warrick, R., & Sharma, K.
(2009). In mice lacking V2a interneurons, gait depends on
speed of locomotion. The Journal of Neuroscience, 29(21),7098–7109.
Del Barrio, M. G., Taveira-Marques, R., Muroyama, Y.,
Yuk, D. I., Li, S., Wines-Samuelson, M., et al. (2007).
A regulatory network involving Foxn4, Mash1 and delta-like
4/Notch1 generates V2a and V2b spinal interneurons from acommon progenitor pool. Development, 134(19), 3427–3436.
Dubreuil, V., Thoby-Brisson, M., Rallu, M., Persson, K.,
Pattyn, A., Birchmeier, C., et al. (2009). Defective respiratory rhythmogenesis and loss of central chemosensitivity in
Phox2b mutants targeting retrotrapezoid nucleus neurons.
The Journal of Neuroscience, 29(47), 14836–14846.
Ericson, J., Rashbass, P., Schedl, A., Brenner-Morton, S.,
Kawakami, A., van Heyningen, V., et al. (1997). Pax6 controls progenitor cell identity and neuronal fate in response
to graded Shh signaling. Cell, 90(1), 169–180.
Farago, A. F., Awatramani, R. B., & Dymecki, S. M. (2006).
Assembly of the brainstem cochlear nuclear complex is revealed by intersectional and subtractive genetic fate maps.
Neuron, 50(2), 205–218.
Fujiyama, T., Yamada, M., Terao, M., Terashima, T.,
Hioki, H., Inoue, Y. U., et al. (2009). Inhibitory and excitatory subtypes of cochlear nucleus neurons are defined by
distinct bHLH transcription factors, Ptf1a and Atoh1. Development, 136(12), 2049–2058.
Gosgnach, S., Lanuza, G. M., Butt, S. J., Saueressig, H.,
Zhang, Y., Velasquez, T., et al. (2006). V1 spinal neurons
regulate the speed of vertebrate locomotor outputs. Nature,440(7081), 215–219.

Goulding, M. (2009). Circuits controlling vertebrate locomotion: Moving in a new direction. Nature Reviews Neuroscience, 10(7), 507–518.
Gray, P. A. (2008). Transcription factors and the genetic organization of brain stem respiratory neurons. Journal ofApplied Physiology, 104(5), 1513–1521.
Gray, P. A., Janczewski, W. A., Mellen, N., McCrimmon, D. R.,& Feldman, J. L. (2001). Normal breathing requires
preBotzinger complex neurokinin-1 receptor-expressing
neurons. Nature Neuroscience, 4(9), 927–930.
Grillner, S., & Jessell, T. M. (2009). Measured motion:
Searching for simplicity in spinal locomotor networks. Current Opinion in Neurobiology, 19(6), 572–586.
Gross, M. K., Dottori, M., & Goulding, M. (2002). Lbx1
specifies somatosensory association interneurons in thedorsal spinal cord. Neuron, 34(4), 535–549.
Gross, M. K., Moran-Rivard, L., Velasquez, T., Nakatsu, M. N.,
Jagla, K., & Goulding, M. (2000). Lbx1 is required for muscle
precursor migration along a lateral pathway into the limb.
Development, 127(2), 413–424.
Hagglund, M., Borgius, L., Dougherty, K. J., & Kiehn, O.
(2010). Activation of groups of excitatory neurons in the
mammalian spinal cord or hindbrain evokes locomotion.
Nature Neuroscience, 13(2), 246–252.
Jensen, P., Farago, A. F., Awatramani, R. B., Scott, M. M.,
Deneris, E. S., & Dymecki, S. M. (2008). Redefining theserotonergic system by genetic lineage. Nature Neuroscience,
11(4), 417–419.
Jessell, T. M. (2000). Neuronal specification in the spinal cord:
Inductive signals and transcriptional codes. Nature Reviews
Genetics, 1(1), 20–29.
Karunaratne, A., Hargrave, M., Poh, A., & Yamada, T.
(2002). GATA proteins identify a novel ventral interneuron
subclass in the developing chick spinal cord. Developmental
Biology, 249(1), 30–43.
Kiehn, O. (2006). Locomotor circuits in the mammalian spinal
cord. Annual Review of Neuroscience, 29, 279–306.
Kim, J. C., Cook, M. N., Carey, M. R., Shen, C.,
Regehr, W. G., & Dymecki, S. M. (2009). Linking genetically defined neurons to behavior through a broadly applicable silencing allele. Neuron, 63(3), 305–315.
Kimura, Y., Okamura, Y., & Higashijima, S. (2006). alx, azebrafish homolog of Chx10, marks ipsilateral descending
excitatory interneurons that participate in the regulation of
spinal locomotor circuits. The Journal of Neuroscience, 26(21), 5684–5697.
Kimura, Y., Satou, C., & Higashijima, S. (2008). V2a andV2b neurons are generated by the final divisions ofpair-producing progenitors in the zebrafish spinal cord.
Development, 135(18), 3001–3005.
Kjaerulff, O., & Kiehn, O. (1996). Distribution of networks
generating and coordinating locomotor activity in the neonatal rat spinal cord in vitro: A lesion study. The Journalof Neuroscience, 16(18), 5777–5794.

Author's personal copy
36
Kjaerulff, O., & Kiehn, O. (1997). Crossed rhythmic synaptic
input to motoneurons during selective activation of thecontralateral spinal locomotor network. The Journal of
Neuroscience, 17(24), 9433–9447.
Kobayakawa, K., Kobayakawa, R., Matsumoto, H., Oka, Y.,
Imai, T., Ikawa, M., et al. (2007). Innate versus learned
odour processing in the mouse olfactory bulb. Nature, 450(7169), 503–508.
Kudo, N., & Yamada, T. (1987). Morphological and physiological studies of development of the monosynaptic reflex pathway in the rat lumbar spinal cord. Journal de Physiologie,
389, 441–459.
Lanuza, G. M., Gosgnach, S., Pierani, A., Jessell, T. M., &Goulding, M. (2004). Genetic identification of spinal interneurons that coordinate left–right locomotor activity necessary for walking movements. Neuron, 42(3), 375–386.
Lechner, H. A., Lein, E. S., & Callaway, E. M. (2002).
A genetic method for selective and quickly reversible silencing of Mammalian neurons. The Journal of Neuroscience, 22(13), 5287–5290.
Li, S., Misra, K., Matise, M. P., & Xiang, M. (2005). Foxn4 acts
synergistically with Mash1 to specify subtype identity of V2
interneurons in the spinal cord. Proceedings of the National
Academy of Sciences of the United States of America, 120(30), 10688–10693.
Liem, K. F., Jr., Tremml, G., & Jessell, T. M. (1997). A role for theroof plate and its resident TGFbeta-related proteins in neuronal patterning in the dorsal spinal cord. Cell, 91(1), 127–138.
Lundfald, L., Restrepo, C. E., Butt, S. J., Peng, C. Y.,
Droho, S., Endo, T., et al. (2007). Phenotype of V2-derived
interneurons and their relationship to the axon guidance
molecule EphA4 in the developing mouse spinal cord. The
European Journal of Neuroscience, 26(11), 2989–3002.
Luo, L., Callaway, E. M., & Svoboda, K. (2008). Genetic dissection of neural circuits. Neuron, 57(5), 634–660.
Luquet, S., Perez, F. A., Hnasko, T. S., & Palmiter, R. D.
(2005). NPY/AgRP neurons are essential for feeding in
adult mice but can be ablated in neonates. Science, 310(5748), 683–685.
McKay, L. C., & Feldman, J. L. (2008). Unilateral ablation of
pre-Botzinger complex disrupts breathing during sleep but
not wakefulness. American Journal of Respiratory and Critical Care Medicine, 178(1), 89–95.
Mizuguchi, R., Kriks, S., Cordes, R., Gossler, A., Ma, Q., &Goulding, M. (2006). Ascl1 and Gsh1/2 control inhibitoryand excitatory cell fate in spinal sensory interneurons.
Nature Neuroscience, 9(6), 770–778.
Moran-Rivard, L., Kagawa, T., Saueressig, H., Gross, M. K.,
Burrill, J., & Goulding, M. (2001). Evx1 is a postmitotic
determinant of v0 interneuron identity in the spinal cord.
Neuron, 29(2), 385–399.
Muller, T., Brohmann, H., Pierani, A., Heppenstall, P. A.,
Lewin, G. R., Jessell, T. M., et al. (2002). The homeodomain

factor lbx1 distinguishes two major programs of neuronaldifferentiation in the dorsal spinal cord. Neuron, 34(4),551–562.
Myers, C. P., Lewcock, J. W., Hanson, M. G., Gosgnach, S.,
Aimone, J. B., Gage, F. H., et al. (2005). Cholinergic input isrequired during embryonic development to mediate proper
assembly of spinal locomotor circuits. Neuron, 46(1), 37–49.
Nagel, G., Szellas, T., Huhn, W., Kateriya, S., Adeishvili, N.,
Berthold, P., et al. (2003). Channelrhodopsin-2, a directly
light-gated cation-selective membrane channel. Proceedingsof the National Academy of Sciences of the United States ofAmerica, 100(24), 13940–13945.
Ng, Y. K., George, K. M., Engel, J. D., & Linzer, D. I. (1994).
GATA factor activity is required for the trophoblast-specifictranscriptional regulation of the mouse placental lactogen I
gene. Development, 120(11), 3257–3266.
Onimaru, H., & Homma, I. (1987). Respiratory rhythm generator neurons in medulla of brainstem–spinal cord preparation from newborn rat. Brain Research, 403(2), 380–384.
Pagliardini, S., Ren, J., Gray, P. A., Vandunk, C., Gross, M.,
Goulding, M., et al. (2008). Central respiratory
rhythmogenesis is abnormal in lbx1- deficient mice. The
Journal of Neuroscience, 28(43), 11030–11041.
Peng, C. Y., Yajima, H., Burns, C. E., Zon, L. I., Sisodia, S. S.,
Pfaff, S. L., et al. (2007). Notch and MAML signaling drives
Scl-dependent interneuron diversity in the spinal cord. Neuron, 53(6), 813–827.
Pierani, A., Moran-Rivard, L., Sunshine, M. J., Littman, D. R.,
Goulding, M., & Jessell, T. M. (2001). Control of interneuron fate in the developing spinal cord by the progenitor
homeodomain protein Dbx1. Neuron, 29(2), 367–384.
Rabe, N., Gezelius, H., Vallstedt, A., Memic, F., &Kullander, K. (2009). Netrin-1-dependent spinal interneuron subtypes are required for the formation of left–rightalternating locomotor circuitry. The Journal of Neuroscience, 29(50), 15642–15649.
Rose, M. F., Ren, J., Ahmad, K. A., Chao, H. T., Klisch, T. J.,
Flora, A., et al. (2009). Math1 is essential for the development of hindbrain neurons critical for perinatal breathing.
Neuron, 64(3), 341–354.
Sander, M., Paydar, S., Ericson, J., Briscoe, J., Berber, E.,
German, M., et al. (2000). Ventral neural patterning by
Nkx homeobox genes: Nkx6.1 controls somatic motor neuron and ventral interneuron fates. Genes & Development,
14(17), 2134–2139.
Sapir, T., Geiman, E. J., Wang, Z., Velasquez, T., Mitsui, S.,
Yoshihara, Y., et al. (2004). Pax6 and engrailed 1 regulate
two distinct aspects of Renshaw cell development. The Journal of Neuroscience, 24(5), 1255–1264.
Saueressig, H., Burrill, J., & Goulding, M. (1999). Engrailed-1
and netrin-1 regulate axon pathfinding by associationinterneurons that project to motor neurons. Development,
126(19), 4201–4212.

Author's personal copy
37
Smith, J. C., Ellenberger, H. H., Ballanyi, K., Richter, D. W.,& Feldman, J. L. (1991). Pre-Botzinger complex: A
brainstem region that may generate respiratory rhythm inmammals. Science, 254(5032), 726–729.
Smith, J. C., & Feldman, J. L. (1987). In vitro brainstem–spinal
cord preparations for study of motor systems for mammalian respiration and locomotion. Journal of Neuroscience
Methods, 21(2–4), 321–333.
Stepien, A. E., & Arber, S. (2008). Probing the locomotor
conundrum: Descending the ‘V’ interneuron ladder. Neuron,60(1), 1–4.
Storm, R., Cholewa-Waclaw, J., Reuter, K., Brohl, D.,
Sieber, M., Treier, M., et al. (2009). The bHLH transcription
factor Olig3 marks the dorsal neuroepithelium of the hindbrain and is essential for the development of brainstem
nuclei. Development, 136(2), 295–305.
Tan, W., Janczewski, W. A., Yang, P., Shao, X. M.,
Callaway, E. M., & Feldman, J. L. (2008). Silencing preBotzinger complex somatostatin-expressing neurons induces
persistent apnea in awake rat. Nature Neuroscience, 11(5),
538–540.
Thoby-Brisson, M., Karlen, M., Wu, N., Charnay, P.,
Champagnat, J., & Fortin, G. (2009). Genetic identificationof an embryonic parafacial oscillator coupling to the
preBotzinger complex. Nature Neuroscience, 12(8), 1028–1035.
Timmer, J. R., Wang, C., & Niswander, L. (2002). BMP signaling patterns the dorsal and intermediate neural tube via regulation of homeobox and helix-loop-helix transcription
factors. Development, 129(10), 2459–2472.
Vallstedt, A., Muhr, J., Pattyn, A., Pierani, A.,
Mendelsohn, M., Sander, M., et al. (2001). Different levelsof repressor activity assign redundant and specific roles toNkx6 genes in motor neuron and interneuron specification.
Neuron, 31(5), 743–755.
Wallen-Mackenzie, A., Gezelius, H., Thoby-Brisson, M.,
Nygard, A., Enjin, A., Fujiyama, F., et al. (2006). Vesicularglutamate transporter 2 is required for central respiratory
rhythm generation but not for locomotor central pattern generation. The Journal of Neuroscience, 26(47), 12294–12307.
Wang, Z., Li, L., Goulding, M., & Frank, E. (2008). Early postnatal development of reciprocal Ia inhibition in the murine
spinal cord. Journal of Neurophysiology, 100(1), 185–196.
Wildner, H., Muller, T., Cho, S. H., Brohl, D., Cepko, C. L.,
Guillemot, F., et al. (2006). dILA neurons in the dorsal spinal cord are the product of terminal and non-terminal asymmetric progenitor cell divisions, and require Mash1 for theirdevelopment. Development, 133(11), 2105–2113.
Wyart, C., Del Bene, F., Warp, E., Scott, E. K., Trauner, D.,
Baier, H., et al. (2009). Optogenetic dissection of a behavioural module in the vertebrate spinal cord. Nature, 461(7262), 407–410.
Yamamoto, M., Shook, N. A., Kanisicak, O., Yamamoto, S.,
Wosczyna, M. N., Camp, J. R., et al. (2009). A multifunctional reporter mouse line for Cre- and FLP-dependent lineage analysis. Genesis, 47(2), 107–114.
Yamamoto, M., Wada, N., Kitabatake, Y., Watanabe, D.,
Anzai, M., Yokoyama, M., et al. (2003). Reversible suppression of glutamatergic neurotransmission of cerebellar granule cells in vivo by genetically manipulated expression oftetanus neurotoxin light chain. The Journal of Neuroscience,
23(17), 6759–6767.
Zagoraiou, L., Akay, T., Martin, J. F., Brownstone, R. M.,
Jessell, T. M., & Miles, G. B. (2009). A cluster of cholinergic
premotor interneurons modulates mouse locomotor activity.
Neuron, 64(5), 645–662.
Zhang, Y., Narayan, S., Geiman, E., Lanuza, G. M.,
Velasquez, T., Shanks, B., et al. (2008). V3 spinal neurons
establish a robust and balanced locomotor rhythm during
walking. Neuron, 60(1), 84–96.
Zhou, Y., Yamamoto, M., & Engel, J. D. (2000). GATA2 isrequired for the generation of V2 interneurons. Development, 127(17), 3829–3838.