Neurite outgrowth traced by means of horseradish peroxidase inherited from neuronal ancestral cells in frog embryos

DEVELOPMENTAL

BIOLOGY

110,

102-113

(1985)

Neurite Outgrowth Traced by Means of Horseradish Peroxidase Inherited from Neuronal Ancestral Cells in Frog Embryos’ MARCUS JACOBSONAND Department

of Anatmy,

Received

University

September

of Utah

School

21, 1984;

SEN HUANG~ of Medicine,

accepted

January

Salt Lake

City,

Utah

841.32

25, 1985

Outgrowing neurites in Xenopus embryos were labeled with horseradish peroxidase which had been injected into a single blastomere at the 32-cell stage and had been inherited by all the descendants, including neurons. Neurite outgrowth was traced from labeled trigeminal ganglion cells and most or all types of neurons present in the spinal cord at embryonic stages 20-30: primary motoneurons, commissural, dorsal longitudinal, ventral longitudinal, and Rohon-Beard neurons. All types of nerve fibers grew by the most direct pathway, apparently without errors of initial outgrowth, pathway selection, or target selection. An initial transient phase of outgrowth of filopodial processes from neuronal cell bodies and shafts of short neurites was observed which disappeared after further elongation of the neurites. The first pioneer fibers grew out from all types in a 2-hr period, from stage 20 to 22, and these fibers arrived at the targets within 3.5 hr after initial outgrowth. Additional fibers grew later in contact with the pioneers to form fascicles. Nerve fibers elongated without branching until they neared or contacted their targets. The rate of elongation at 20°C was 30-75 pm/hr. The rapid, unbranched, error-free initial outgrowth and elongation of neurites to their targets is discussed in relation to theories of development of nerve pathways. o 1985 Academic press, IK

INTRODUCTION

studying axonal regeneration in older embryos or adults or after experimental surgery on embryos at stages of development after the initial outgrowth of nerve fibers. We have tried to avoid many of these limitations by studying neurite outgrowth in the central and peripheral nervous system of normal embryos without any experimental interference with the outgrowing nerve fibers, their pathways, or their targets. We have taken advantage of the observation that horseradish peroxidase (HRP) is a heritable cell lineage tracer in Xenopus embryos (Hirose and Jacobson, 1979; Jacobson and Hirose, 1981; Jacobson, 1981, 1983). After injection of HRP into a single blastomere, the tracer is transmitted during mitosis to all the descendants and can be seen up to a week later in well-differentiated cells, including neurons and their peripheral targets. The tracer enters the outgrowing neurites and shows the position and direction of initial outgrowth of axons as well as dendrites, the pattern of neurite branching, and the relation between axon terminals and specific targets (Jacobson, 1983; Moody and Jacobson, 1983). The inherited HRP is confined intracellularly so that the problem of selective and prolonged uptake from an extracellular pool of tracer is completely avoided. The foundations for the present investigation have been laid by studies of the structure and function of the developing spinal cord in Xenms embryos (Hughes, 1957, 1959; Macklin and Wojtkowski, 1973; Hayes and Roberts, 1974; Roberts and Smyth, 1974; Muntz, 1975; Kevetter and Lasek, 1980; Lamborghini, 1980; Forehand

Harrison (1910) clearly defined the problem of development of nerve circuit pathways and thought that formation of the initial connections could be explained in terms of predetermination of the initial direction of axon outgrowth, the motive force of axon elongation, guidance along the pathway, and specific attraction between outgrowing axons and peripheral targets. But he conceded that “the specific arrangement of fibers within the central nervous system affords a morphogenetic problem of much greater difficulty” than the problem of development of peripheral nerve pathways. In addition to the conceptual difficulties there are technical problems with tracing axonal outgrowth in small embryos. In small and rapidly developing embryos it is not possible to obtain the necessary spatial and temporal resolution by extracellular injection of HRP or other tracers: spatial resolution is limited by diffusion from the injection site and temporal resolution is limited by relatively prolonged uptake of the tracer from an extracellular pool (Mesulam, 1982). Because of the difficulties of studying the initial development of nerve circuit pathways in early embryos of vertebrates, the problem has been approached indirectly by i This work was supported by Grant BNS 8116768 from the National Science Foundation. We thank Hannelore Mueller and Karen Evans for their expert histological and secretarial sssistance. ’ Present address: Shanghai Brain Research Institute, Academia Sinica, 319 Yo-Yang Road, Shanghai, Peoples Republic of China.

0012-1606/85 Copyright All rights

$3.00

0 1985 by Academic Press. Inc. of reproduction in any form reserved.

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and Farel, 1982a,b; Spitzer, 1982; Roberts and Clarke, 1982; Nordlander, 1984). The rostra1 part of the spinal cord develops earliest and spinal reflex circuits apparently start functioning at about stage 24 of Nieuwkoop and Faber (1967). Neuromuscular junctions start becoming functional at stage 24 (Blackshaw and Warner, 1976; Kullberg et al., 1977). Innervation of the trunk skin by peripheral neurites of Rohon-Beard neurons starts at stage 24 (Roberts and Taylor, 1982) and peripheral trigeminal axons reach the ventral head skin and cement gland at stage 26 (Roberts and Blight, 1975; Davies et al., 1982). By stage 37-38 there are several types of spinal interneurons with local connections, neurons with long intraspinal axons, and primary sensory and motor neurons (Roberts and Clarke, 1982; Nordlander, 1984). However, little information is available about the times of initial axon outgrowth from each type of neuron, rates of axonal elongation, and the paths of pioneer axons. The main aim of this investigation is to trace the spatial and temporal pattern of outgrowth of neurites from identified types of neurons in Xenopus embryos from the earliest stages at which neurites start growing out of the young neurons. METHODS

Embryo product&m and treatment were the same as published previously (Jacobson, 1983). Xenopus laevis embryos were obtained by matings induced by injection of human chorionic gonadotrophin. At the 2- to 4-cell stage the jelly coat was removed by washing the eggs for 4 min in 9 mM dithiothreitol buffered with Hepes to pH 8.9, followed by three washes in 100% Steinberg solution. Eggs were selected with well-marked pigment gradients and regular cleavage patterns at the 32-cell stage for intracellular injection (Jacobson, 1981; Jacobson and Hirose, 1981). Injection of tracers. An injection of horseradish peroxidase (HRP, Type IX, Sigma, l-2 nl, about 10% in distilled water) or biotinylated horseradish peroxidase (Vector laboratories, l-2 nl, about 10% in distilled water) was given by means of a micropipet (tip diameter l-5 pm) into a single blastomere. Blastomeres D1.l.l, D1.1.2, D1.2.1, D1.2.2, D2.1.2, D2.2.2, V1.2.1, V1.2.2 were injected at the 32-cell stage as previously described (Jacobson, 1981; Jacobson and Hirose, 1981). These blastomeres were selected because their descendants occupied overlapping clonal domains in the central nervous system and they were known to be ancestral to a number of types of neurons, especially in the rhombencephalon and spinal cord (Jacobson, 1981; Jacobson and Hirose, 1981). The 336 embryos injected with HRP and 37 embryos injected with biotinylated HRP developed normally through gastrulation and

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were fixed at embryonic stages 18 to 30. The small percentage of cases that failed to develop normally were due to arrest of mitosis in the injected cell and those cases were rapidly detected and excluded. Embryos were raised in 20% Steinberg solution until they reached embryonic stages 18 to 30 and they were then fixed for 6 hr at 4°C in 0.5% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4. Embryos were processed histologically as described below. Histochemistry. After fixation, embryos were washed twice in sucrose solution (5% for 6 to 24 hr and 15% for 1 to 2 hr, both in 0.1 M phosphate buffer, pH 7.4, at 4°C). Frozen sections were cut at 28 pm and mounted on subbed slides. Specimens injected with HRP were soaked for 10 min in 0.1 M phosphate buffer, pH 7.4, containing 12.5 mg diaminobenzidine tetrahydrochloride/100 ml solution, after which hydrogen peroxide (0.3%, 0.5 ml/100 ml soaking solution) was added. The reaction was monitored at intervals until a brown reaction product, which will be referred to as label, became visible under the microscope (5 to 10 min). The slides were washed, dehydrated, and coverslipped for further study. Specimens injected with biotinylated HRP were sectioned as stated above. Sections were reacted with avidin-biotynilated HRP (Vectastain, standard ABC kit) for 60 min, washed three times (15 min) in phosphate buffer, pH 7.4, and reacted with diaminobenzidine tetrahydrochloride and hydrogen peroxide as described above until a brown reaction product became visible under the microscope (10 to 20 min). Whole mount preparaticms. After fixation the central nervous system was dissected out. In some specimens the skin and somites on the labeled side were left attached to the CNS. The dissected specimens were reacted by soaking for 20 min in 0.1 M phosphate buffer containing 25 mg diaminobenzidine hydrochloride/100 ml solution, after which hydrogen peroxide was added (0.3%, 0.5 ml/100 ml soaking solution) and the reaction was terminated after 20 min. The specimens were washed, dehydrated, cleared in xylene and Canada balsam, and mounted between two coverslips. RESULTS

In agreement with earlier reports, labeling a single blastomere at the 32-cell stage resulted in labeling of a large fraction of the total population of neurons which were mingled with unlabeled cells (Jacobson and Hirose, 1981; Jacobson and Moody, 1984). All blastomeres of the 32-cell stage that gave rise to CNS were labeled in this series of embryos. Therefore, it is highly probable that spinal cord neurons of all types present at embryonic stages 20 to 30 were represented in the

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sample of labeled cells (Jacobson and Hirose, 1981). We have observed neurites growing out of trigeminal ganglion cells and all types of spinal cord neurons that have previously been described (Table 1): Rohon-Beard neurons (Hughes, 1957; Lamborghini, 1980; Roberts and Clarke 1982); primary motoneurons (Forehand and Farel, 1982; Moody and Jacobson, 1983); commissural neurons (Roberts and Clarke 1982); dorsal and ventral longitudinal neurons which appear to be early stages of the dorsolateral and ventrolateral ascending and descending interneurons of Roberts and Clarke (1982). For each type of neuron, a complete series of stages of initial outgrowth of the nerve fiber could be seen, starting with emergence of the growth cone from the cell body (Fig. 1) followed by progressive elongation of the unbranched neurite which ended in a typical growth cone (Fig. 1). At the earliest stage of outgrowth, several fine processes resembling filopodia could be seen emerging directly out of the nerve cell body of the pioneer neurons and the neurons whose initial neurite outgrowth occurred later (Fig. 1). Many fine processes extended from the shafts of neurites during the initial period of outgrowth when the neurite was less than 100~pm long (Figs. lB, 2). Filopodial processes were not present on the shafts of longer neurites but were confined to the growth cones (Figs. l-3). No outgrowth of neurites was seen at stages 18 and 19. The first neurites emerged at stages 20 and 21. The neurites grew out about 1 hr earlier from commissural neurons, Rohon-Beard neurons, and dorsal longitudinal neurons in the dorsal part of the rhombencephalon and rostra1 spinal cord than from neurons in the

DEVELOPMENT Initial stage

5w Rohon-Beard

110, 1985

ventral spinal cord (Table 1). By stage 22, five types of labeled neurons with labeled neurites could be identified in the CNS. In addition to the three mentioned above, located dorsally, primary motoneurons and ventral longitudinal neurons were located in the ventral half of the rhombencephalon and spinal cord. The central processes of trigeminal ganglion cells started growing out at stage 21 and their peripheral processes were first seen at stage 22. The earliest fibers to grow out of each type of neuron in virgin pathways are called pioneer fibers (Harrison, 1910), and those which followed later are called secondary fibers. The pioneer fibers were very sparse at stages 20 to 23: only 11 labeled neurites were seen in 8 specimens at stage 20; 52 in 15 specimens at stage 21; 99 in 22 specimens at stage 22; and 157 neurites in 25 specimens at stage 23. It became difficult to count fibers at later stages because the numbers increased very rapidly and fibers formed fascicles. The secondary fibers grew in proximity to the pioneer fibers and to one another, often with their growth cones and filopodia in contact with other axons or growth cones (Fig. 3). By stage 24 the pioneer fibers of all types of neurons had contacted their targets or putative targets or were in the target zone. The peripheral targets of trigeminal ganglion cells and Rohon-Beard neurons in the epidermis have been well defined (Davies et al., 1982; Roberts and Taylor, 1982) and the peripheral targets of primary motoneurons in somitic muscle are known (Kullberg et al., 1977; Moody and Jacobson, 1983) but the central targets of these neurons and the targets of spinal interneurons are not known with certainty (Roberts

TABLE 1 OF PIONEER NEURITES

IN Xenqnq

Contact target stage (time, hr) ’

EMBRYOS Neurite

length

Elongation (pm/hr)”

(rmP

St 20 (0)

St 23 (3)

122 at St 23

41

Commissural

St 20 (0)

St 22 (2)

71 at St 21 145 at St 22

71 73

Dorsal

St 20 (0)

St 22 (2)

150 at St 22 268 at St 24

75 60

motoneuron

St 21 (1)

St 24 (3.5)

110 at St 24

31

V sensory

central

St 21 (1)

St 22-24

900 at St 30

64

V sensory

peripheral

St 22 (2)

St 24 (2.5)

76 at St 24 300 at St 26

30 50

St 22 (2)

St 24 (2.5)

323 at St 26 350 at St 28

54 35

longitudinal

Primary

Ventral

(peripheral)

outgrowth (time, hr)”

VOLUME

longitudinal

(l-3.5)

a Time from stage 20. See text for identity of targets. *Length of longest neurite at given stage (gives a lower limit of elongation rate). “Time from stage 20 to 21, 22, 23, 24, 26, 28, 30, was 1, 2, 3, 4.5, 8, 12, 15 hr at 20°C.

rate

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Neurite

Outgrowth

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FIG. 1. Whole mount of trunk spinal cord of Xenopus embryo at stage 24 after injection of HRP into blastomere V1.2.1 at the 32-cell stage. Neuron (N) at stage of initiation of neurite outgrowth, with filopodial processes on the cell body. Dorsal longitudinal neuron (D) with neurites extending rostrally and caudally terminating in growth cones with filopodia (arrowheads). Double arrowheads point at filopodia on the neurite shaft. Commissural neurons (C) with short, broad neurite extending ventrally, terminating in a growth cone with filopodia. Rostra1 to the left, dorsal to the top of photographs. Magnification bar equals 100 pm (A); 20 pm (B). FIG. 2. Rohon-Beard neuron at stage 24, labeled with HRP inherited from its ancestral cell. The long central neurite (L) with smooth shaft, terminates in a growth cone with filopodia (arrowhead) extending rostrally. The short neurite (S) has many filopodia on the shaft (double arrowheads) as well as on the terminal growth cone (arrowhead). Arrow points at peripheral neurite. Whole mount. Magnification bar, 50 pm.

and Clarke, 1982). Neurites with defined targets grew directly toward them, without branching and without deviating from the most direct pathway. This was

clearly evident for the peripheral processes of RohonBeard neurons, primary motoneurons, and trigeminal ganglion cells whose normal pathways and targets are

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FIG. 3. Dorsal longitudinal neurons at stage 24 labeled with HRP inherited from a single ancestral cell (D1.2.1) Several growth cones (arrowheads) are in direct contact or within filopodial reach of one another or of other neurites or opposite direction. Rostra1 is to the left. Whole mount. Magnification bar: (B) 100 pm; (A and C) 20 pm.

well known. The peripheral neurite of the Rohon-Beard neuron started outgrowth at stage 20 and reached the epidermis, the definitive target, at stage 23, approximately 3 hr later. The fiber always grew out in the correct direction, and by the shortest route, without branching (Fig. 4). At stages 24-30, short branches to the epidermis sprouted from the peripheral end of the Rohon-Beard neurite. The central neurites of RohonBeard neurons extended, without branching, longitudinally in both directions in the dorsolateral spinal cord (Fig. 2). Similarly, the neurites of all other types grew directly toward their targets or putative targets without branching until close to or in contact with the targets (Figs. l-8). The peripheral neurites of trigeminal ganglion cells grew straight, without branching for lengths of more than 400 pm (Fig. 8A) before reaching the epidermis where branching occurred (Fig. 8B).

at the growing

32-cell stage. in the same

All neurites grew in a stereotyped pathway that was characteristic for each type of neuron. It is remarkable that no aberrant neurites were seen at any of the stages examined. Outgrowth of secondary neurites continued from neurons of all types so that increasing numbers of neurites in all stages of outgrowth were seen at stages 24 to 30 in the rhombencephalon and rostra1 spinal cord, which were the regions of most advanced development. The events described above were delayed by a few hours more rostrally and in the caudal trunk spinal cord (Fig. 5). We made no observations on the tail spinal cord which originates considerably later (Nordlander, 1984) by different developmental processes of secondary neural induction and secondary neurulation (Vogt, 1926; Kingsbury, 1932; Holmdahl, 1933; Bij tel, 1958). Rate of pioneer neurite elongation could be calculated from measurements of the longest fibers of each type

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FIG. 4. Rohon-Beard neuron showing single, unbranched, peripheral neurite (arrowhead) extending directly to the epidermis (E). Coronal section through trunk spinal cord at stage 24. The label was injected into blastomere V1.2.1 at the 32-cell stage. M, myotome; S, spinal cord. Magnification bar 50 pm.

at progressively later stages of outgrowth. The elongation rate was calculated on the assumption that initial outgrowth of all the longest fibers started at stage 20, but some may have started elongating after stage 20 and there may have been longer fibers than those observed, so that these calculations gave lower limits. Elongation rate for different types of neurites ranged from 30 to 75 pm/hr (Table 1). No significant differences between preparations injected with HRP or biotinylated HRP were seen at stages 18 to 30. Biotinylated HRP yielded a more uniformly dispersed and very fine reaction product which labeled filopodia very effectively. The biotinylated HRP could be detected in neurites of many types of neurons by means of the biotin-avidin technique at later stages (up to stage 45), whereas few neurites were labeled at stage 45 after HRP injection. In spite of the greater detectability of the biotinylated HRP at later stages, the appearance and number of outgrowing neurites and the times of their initial outgrowth at stages 20 to 30 were the same when either HRP or biotinylated HRP was used as a tracer. DISCUSSION

The developmental timetable of initial outgrowth of pioneer neurites from several types of neurons has been elucidated during early development of the central and peripheral nervous system in a vertebrate embryo. Young neurons were labeled with HRP inherited from their ancestral cells so that the outgrowth of neurites could be followed from the earliest stages. One advan-

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tage of this method is that the embryos were not injured or manipulated in any way during the period of neuron origin, differentiation, and neurite outgrowth. Stages of neurite outgrowth could be traced from before outgrowth, through the stages of initial outgrowth, elongation and pathway selection, approach to and contact with the targets, and branching. A single neuron or a few neurons of each type were the first to initiate neurite outgrowth. These pioneer neurites grew out into virgin pathways, always correctly oriented, straight toward their targets without branching until the target was contacted. No phase of trial and error or excessive sprouting was ever seen and no neurites were seen growing in aberrant directions. During initial outgrowth, until the neurite was about 100 pm in length its surface was covered with many filopodial outgrowths of different lengths up to about 20 pm (Fig. 2). A few filopodial outgrowths were seen on some nerve cell bodies during the phase of initial neurite outgrowth (Fig. 1). During this phase, the entire neurite and parts of the cell body appeared to be sensing the local environment by means of filopodia, or these filopodia may represent a response of the neuron to growth-promoting conditions in their local environment. The initial stage of generalized filopodial outgrowth was seen in pioneer neurons and secondary neurons that initiated neurite outgrowth later. These appearances were remarkably like those seen during initiation of neurite outgrowth in chick ciliary ganglion cells in culture (Collins, 1978a). The filopodial processes were not seen on the cell body or neurite shaft after the neurite had reached about 100 grn in length, about 2 hr after initial outgrowth of the neurite. At that stage the filopodia were confined to the expanded terminal portion of the neurite (Figs. 1, 3), which had the typical appearance of a growth cone (Bray and Bunge, 1973; Wessels et al., 1980; Letourneau, 1982). The sudden onset and short duration of the phase of filopodial outgrowth from the cell body and neurite shafts raises questions about the functions and developmental control of the outgrowths. Are they the result of a sudden transient change in extracellular conditions or an abrupt change in the neuron’s ability to respond to constant external conditions or a combination of intrinsic and extrinsic changes? The observation that both types of neurites, smooth and filopodial, were often seen growing simultaneously from the same neuron (Figs. 1,2) shows that they were probably not responses to changes in external conditions but represented differences in the neuron itself. The problem may be studied by heterochronic transplantation of neurons from the stage prior to pioneer neurite initiation to older embryos at stages after completion of pioneer neurite outgrowth.

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All these events occurred rapidly: no neurites were seen at stage 19 but within a 2-hr period, from stage 19 to 21, initiation of neurite outgrowth had occurred in five types of CNS neurons as well as trigeminal ganglion cells. Three to four hours later all these pioneer axons had arrived at their targets (Table 1). The rate of elongation was 30-75 ym/hr, which was close to the maximum rate expected in amphibians (Jacobson, 1978, p. 131). These findings have paved the way for studies of the fine structures of the initial neurite outgrowths and the relations between neurites and their environment. One of the questions which may be answered definitively is whether preformed extracellular tunnels between glial cells or neuroepithelial cells act as guides for pioneer fibers at the time of initial outgrowth in Xenopus embryos. Such tunnels have been reported in Xenopus spinal cord at stages considerably after initial outgrowth of pioneer axons (Singer et al., 1979; Nordlander and Singer, 1982) and after injury and regeneration (Egar and Singer, 1972; Nordlander and Singer, 1978). No extracellular tunnels were seen during outgrowth of optic axons in fish (Bodick and Levinthal, 1980; Easter et at, 1984). By contrast with pioneer neurites which grew into virgin pathways, the secondary neurites were frequently seen with growth cones and filopodia in close contact with the pioneers and with one another (Figs. 1, 3). This suggests that secondary fibers are guided by conditions along the pathway such as mechanical support by pioneers (Harrison, 1914; Weiss, 1941) and adhesion between neurites mediated by specific nerve cell adhesion molecules (Rutishauser, 1983; Edelman, 1984). Pioneer fibers of dorsal longitudinal neurons, ventral longitudinal neurons, and commissural neurons grew at the periphery of the neural tube, apparently in association with the basal lamina or with end feet of neuroepithelial cells (Fig. 5). This suggests that their outgrowth may be guided by components of the basal lamina or factors bound to it (Collins and Garrett, 1980; Lander et al., 1982; Rogers et al, 1983) or by the neuroepithelial end feet (Silver and Rutishauser, 1984). However, these relationships cannot be resolved without electron microscopy. By contrast, neurites of trigeminal ganglion cells, Rohon-Beard neurons, and primary mo-

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toneurons penetrated through the basal lamina of the neural tube to reach their targets. Pioneer neurites of primary motoneurons grew out of the spinal cord at the positions of intermyotomal septa, and motor axons that grew later fasciculated to form the ventral spinal roots (Figs. 6, 7). Apparently, there are special conditions at the septa between the myotomes that favor outgrowth of pioneer motor axons. Neurite outgrowth in Xenopus embryos is not a process of trial and error but appears to be under remarkably efficient temporal and spatial constraints on initiation, elongation, and targeting. These findings raise the possibility that when errors of axon growth and excess collateral sprouting have been seen, especially during axon regeneration, they may be pathological responses to injury. The observations on Xenopus embryos are in agreement with those of Lance-Jones and Landmesser (1981) who have shown that axons of spinal motoneurons in the chick embryo grew along specific paths to the limb muscles without any phase of diffuse projections. Such observations are not consistent with models in which axons branch profusely to reach their targets or in which errors of axonal pathway selection occur and are later corrected by elimination of axon branches or of the entire axon or neuron. Another finding which bears on the possible mechanisms required to guide neurites to their targets without errors and without redundant branching, is that the distances over which the pioneer neurites have to navigate are very short, 50-150 pm, at the time of initial outgrowth. This is well within the distance over which a concentration gradient of a diffusible factor can be set up within an hour in the embryo (Crick, 1970; Munro and Crick, 1971). It is conceivable that the initiation, elongation, and target selection of pioneer fibers could be controlled by diffusible factors released from the targets (Rambn y Cajal, 1910, 1928, pp. 278, 378), and growing evidence implicates factors produced by the target (Charnley et al., 1973; Coughlin, 1975; Ebendal and Jacobson, 1977; Letourneau, 1978; Gundersen and Barrett, 1979, 1980; Lumsden and Davies, 1983; Berg, 1984). The possibility of such diffusible factors released from the targets is being tested in Xenopus embryos by removing the targets (Huang and

FIG. 5. Coronal section through rostra1 spinal cord (A) and caudal spinal cord (B) of Xenops embryo at stage 24 showing pioneer commissural neurons (arrowheads) labeled with HRP inherited from an ancestral cell. Dark granular material is melanin. Magnification bar 50 pm. FIG. 6. Ventral longitudinal neuron (V) and primary motoneurons (M) labeled in the trunk spinal at stage 28 after injection of HRP into blastomere D1.1.2 at the 32-cell stage. Whole mount, rostra1 to left. A single, unbranched neurite (double arrowheads) extends rostrally from the VL neuron. Neurites of primary motoneurons extend rostrally and caudally, and some axons have formed ventral spinal roots (arrowheads). Magnification bar 50 pm. FIG. 7. Whole mount of spinal cord (S) and notochord (N) at embryonic stage 28, showing primary motoneurons (M) and motoneuron axons fasciculating to form ventral spinal roots (arrowheads). Magnification bar, 50 Frn.

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Jacobson, in preparation). Lack of branching until the axon reaches the target also implicates the target in some way in the outgrowth of branches. The target may stimulate axonal branching by direct contact or by releasing diffusible factors which have a limited range because they are bound to the substrate, used up, or destroyed locally (Brown et ok, 1981; Nurcombe and Bennett, 1983). It is conceivable that branching occurs only after the axon has attained a certain length but this is contradicted by our observations that surgical removal of targets from Xenopus embryos before the time of initial axon outgrowth resulted in growth of pioneer axons to several times their normal length without branching (Huang and Jacobson, in preparation). Outgrowth of pioneer fibers is comparable in Xenow and insects. In both, pioneer neurites grow out along stereotyped pathways, and the pioneers are later joined by secondary neurites to form fascicles in the central as well as peripheral nervous system (Bate, 1978; Bentley and Kishishian, 1982; Edwards, 1982; Goodman et al., 1982). Although the filopodia on growth cones of vertebrates are much shorter than those of insects, they may both function in a similar manner, reaching out to seek and adhere to a suitable substrate or target. The pioneer fibers are the substrate on which the secondary fibers normally grow but we do not know whether the secondary fibers in Xenopus could successfully navigate a virgin pathway as they can do in the grasshopper after ablation of the pioneer neurons (Bentley and Kishishian, 1982). Until similar ablation experiments are done in Xenopus embryos it remains an open question whether the pioneers are qualitatively different from the secondary fibers in their ability to navigate pathways or whether they merely differ in the time of initial outgrowth. Much evidence shows that the specificity of the cellular interactions involved in neurite guidance is relative and graded rather than absolute and all-ornone (summarized in Jacobson, 1978, p. 348). For example, optic axons growing from an eye grafted close to the spinal cord in Xenopus tadpoles, preferred to associate with axons of the longitudinal spinal tracts but showed no preference for growing rostrally or caudally (Constantine-Paton and Capranica, 1976; Katz and Lasek, 1978, 1979). However, some of those optic axons ultimately reached the tectum (Giorgi and van der Loos, 1978). All those observations were made

FIG. 8. Peripheral trigeminal neurites labeled at stage 26 after injection of HRP into blastomere D1.2.1 at the 32-cell stage. Coronal sections. (A) shows fibers extending from ganglion cells (arrowheads), without branching, in the direction of the cement gland. (B) shows fibers branching in the cement gland. Magnification bar 50 pm.

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after stage 45, long after pioneer fibers had traversed the pathways. Therefore, they do not address the question directly of whether there are cues in the virgin pathways which guide pioneer axons to their targets. Such preformed pathways have been called “substrate pathways” (Katz and Lasek, 1978, 1979; Katz et al., 1980). Direct evidence that optic nerve fibers prefer to follow a preformed pathway of N-CAM during their initial outgrowth in normal chick embryos has been reported by Silver and Rutishauser (1984). Neurite guidance along a pathway of material bound to the substrate has been demonstrated definitively in tissue culture (Collins and Lee, 1984). Now that the events of neurite outgrowth have been accurately timed in the rhombencephalon and trunk spinal cord of Xenopus embryos it is possible to investigate development of conditions which promote initial outgrowth and elongation, guide the neurite along the pathway, and promote branching. It is evident that neurites develop under control of many different factors (Letourneau, 1982; Collins 1984; Rutishauser, 1984). Immunocytochemical studies, using antibodies to wellcharacterized molecules that promote initial outgrowth, elongation, and guide neurites, may now be done in relation to the precise timetable of neurite outgrowth from identified neurons in Xenopus. These morphological changes may be related to the expression of special nerve cell adhesion molecules that promote specific adhesion between neurons or between neurons and glial cells (Rutishauser, 1983, 1984; Edelman, 1984), and development of materials bound to the substrate which have been shown to initiate, guide and stimulate neurite elongation (Collins, 1978a,b; Collins and Garrett, 1980; Collins and Dawson, 1982; Collins and Lee, 1984). Antibodies to such materials and factors may be tested for their ability to perturb neurite outgrowth, elongation, orientation, and branching (Fraser et al., 1984; Silver and Rutishauser, 1984). Xenopus embryos are especially favorable subjects for such studies because of the knowledge already available about earlier stages of development of the nervous system and because the pathways of pioneer fibers and the timetable of their development are now precisely known in Xenopus embryos. CONCLUSIONS

Five types of spinal cord neurons and trigeminal ganglion cells sprouted pioneer fibers during the same 2-hr period from embryonic stage 20 to 22. Additional neurites followed the pioneers at later stages. Therefore, a distinction could be made between pioneer neurites that grow out initially into virgin pathways, within a tightly controlled period of early development and a more extended second period of outgrowth of neurites

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that follow the pioneers. Whether these two periods of neurite outgrowth have different mechanisms and developmental controls remains to be investigated. The earliest stage of outgrowth of pioneer as well as secondary neurons was typified by outgrowth of numerous filopodial processes from the cell body and neurite shaft. We interpret these as tentative sensing of the local environment prior to selection of the direction of initial outgrowth of the neurite and of the definitive pathway. It remains to be determined whether the factors controlling initiation of neurite outgrowth are the intrinsic state of differentiation of the neuron, the state of the local environment, or both. From the stereotyped routes taken by the pioneer neurites, straight to their targets, rapidly (30-75 pm/ hr), without errors or branching, we conclude that elongation and guidance of pioneer neurites is tightly controlled by factors along the route. Mechanisms of trial and error or elimination of excessive sprouts can be excluded from consideration in Xenopus during normal development. The short distances between neurons and targets (50-150 Km) at the time of initial outgrowth is consistent with the possibility of release of diffusible neurite elongation factors from the target, but these remain to be investigated in Xenolpus embryos. Absence of branching of neurites until they are close to or in contact with the target implicates the target in the mechanism of branching. These results indicate that conditions along the virgin pathway into which pioneer neurites penetrate are responsible for guiding and stimulating neurite elongation, but branching is under different controls that are related to proximity to the target.

REFERENCES BATE, C. M. (1978). Development of sensory systems in arthropods. In “Development of Sensory Systems” (M. Jacobson, ed.), pp. l53. Springer-Verlag, New York. BENTLEY, D., and KESHISHIAN, H. (1982). Pioneer neurons and pathways in insect appendages. Trends Neurosci. 5, 364-367. BERG, D. K. (1984). New neuronal growth factors. Annu. Rev. Neurosci. 7, 149-170. BIJTEL, J. H. (1958). The mode of growth of the tail in urodele larvae. J. EmlnyoL Exp. MorphoL 6, 466-478. BLACKSHAW, S., and WARNER, A. (1976). Onset of acetylcholine sensitivity and end-plate activity in developing myotome muscles of Xenopus. Nature (London) 262, 217-218. BODICK, N., and LEVINTHAL, C. (1980). Growing optic nerve fibers follow neighbors during embryogenesis. Proc. Natl. Acad. Sci. USA 77, 4374-4378. BRAY, D., and BUNGE, M. B. (1973). The growth cone in neurite extension. C&r Found. Symp. 14, 195-209. BROWN, M. C., HOLLAND, R. L., and HOPKINS, W. G. (1981). Motor nerve sprouting. Annu. Rev. Neurosci. 4, 17-42. c HAMLEY, J. H., GOLLER, I., and BURNSTOCK, G. (1973). Selective growth of sympathetic nerve fibers to explants of normally densely

112

DEVELOPMENTAL

BIOLOGY

innervated autonomic effector organs in tissue culture. Dm. Biol. 31, 362-379. CONSTANTINE-PATON, M., and CAPRANICA, R. P. (1976). Axonal guidance of developing optic nerves in the frog. I. Anatomy of the projection from transplanted eye primordia. J. Camp. Neural. 170, 17-32. COLLINS, F. (1978a). Axon initiation by ciliary neurons in culture. Dev. BioL 65,50-57. COLLINS, F. (1978b). The induction of neurite outgrowth by a conditioned medium factor bound to the culture substratum. Proc. Notl. Acad. Sci. USA 75, 5210-5213. COLLINS, F. (1984). Multiple sites for the regulation of neurite outgrowth. In “Cellular and Molecular Biology of Neuronal Development” (Ira Black, ed.), pp. 217-230. Plenum, New York. COLLINS, F., and DAWSON, A. (1982). Conditioned medium increases the rate of neurite elongation: Separation of this activity from the substratum-bound inducer of neurite outgrowth. J. Neurosci. 2, 1005-1010. COLLINS, F., and GARRETT, J. E. (1980). Elongating nerve fibers are guided by a conditioned medium factor bound to the culture substratum. Proa NatL Acad. ScZ: USA 77, 6226-6228. COLLINS, F., and LEE, M. R. (1984). The spatial control of ganglionic neurite growth by the substrate associated material from conditioned medium: An experimental model of haptotaxis. J. Neurosci. 4,2823-2829. COUGHLIN, M. D. (1975). Target organ stimulation of parasympathetic nerve growth in the developing mouse submaxillary gland. Dev. Biol. 43, 140-158. CRICK, F. (1970). Diffusion in embryogenesis. Nature (London) 225, 420-422. DAVIES, S. N., KITSON, D. L., and ROBERTS, A. (1982). The development of the peripheral trigeminal innervation in Xenopus embryos. J. Embrl/ol. Exp. Morphol. 70,215-224. EASTER, S. S., BRATTON, B., and SCHERER, S. S. (1984). Growthrelated order of the retinal fiber layer in goldfish. J. Neurosci. 4, 2173-2190. EBENDAL, T., and JACOBSON, C. 0. (1977). Tissue explants affecting extension and orientation of axons in cultured chick embryo ganglia. Exp. Cell Res. 105, 379-387. EDELMAN, G. M. (1984). Modulation of cell adhesion during induction, histogenesis, and perinatal development of the nervous system. Annu. Rev. Neurosci 7,319-377. EDWARDS, J. S. (1982). Pioneer fibers. The case for guidance in the embryonic nervous system of the cricket. In “Neuronal Development” (N. C. Spitzer, ed.), pp. 255-266. Plenum, New York. EGAR, M., and SINGER, M. (1972). The role of ependyma in spinal cord regeneration in the urodele, Triturus. Ezp. Neural 37, 422430. FOREHAND, C. J., and FAREL, P. B. (1982a). Spinal cord development in anuran larvae. I. Primary and secondary neurons. J. Cmnp. Neural 209,386-394. FOREHAND, C. J., and FAREL, P. B. (1982b). Spinal cord development in anuran larvae. II. Ascending and descending pathways. J. Cinnp. NeuroL 209, 395-408. FRASER, S. E., MURRAY, B. A., CHUONG, C.-M., and EDELMAN, G. M. (1984). Alteration of the retinotectal map in Xenopus by antibodies to cell adhesion molecules. Proc Natl. Acod. Sci USA 81, 42224226. GIORGI, P. P., and VAN DER Loos, H. (1978). Axons from eyes grafted in Xenopus can grow into the spinal cord and reach the optic tectum. Nature (London) 275, 746-748. GOODMAN, C. S., RAPER, J. A., Ho, R. K., and CHANG, S. (1982). Pathfinding by neuronal growth cones in grasshopper embryos. In “Developmental Order: Its Origin and Regulation” (S. Subtelny and P. G. Green, eds.), pp. 275-316. Liss, New York. GUNDERSEN, R. W., and BARRETT, J. N. (1979). Neuronal chemotaxis:

VOLUME

110, 1985

Chick dorsal root axons turn toward high concentrations of nerve growth factor. Science (Washingtm, D.C.) 206, 1079-1080. GUNDERSEN, R. W., and BARRETT, J. N. (1980). Characterization of the turning response of dorsal root neurites towards nerve growth factor. J. Cell Biol 87, 546-554. HARRISON, R. G. (1910). The outgrowth of nerve fiber as a mode of protoplasmic movement. J. Exp. Zool. 9, 787-848. HARRISON, R. G. (1914). The reaction of embryonic cells to solid structures. J. Exp. Zool. 17, 521-544. HAYES, B. P., and ROBERTS, A. (1974). The distribution of synapses along the spinal cord of an amphibian embryo: An electron microscopic study of junction development. Cell Tissue Res. 153, 227-244. HIROSE, G., and JACOBSON, M. (1979). Clonal organization of the central nervous system of the frog. I. Clones stemming from individual blastomeres of the 16-cell and earlier stages. Den Biol. 71.191-202. HOLMDAHL, D. E. (1933). Die zweifache Bildungsweise des zentralen Nervensystems bei den Wirbeltieren. Eine formgeschichtliche und materialgeschichtliche Analyse. Wilhelm Roux’ Arch. Entwicklungsmech. Ch-g. 139, 191-226. HUGHES, A. F. W. (1957). The development of the primary sensory system in Xenqpus laevis. J. Anat. 91, 323-338. HUGHES, A. (1959). Studies in embryonic and larval development in amphibia. II. The spinal motor root. J. Emb?yol. Ezp. Morphol. 7, 128-145. JACOBSON, M. (1978). “Developmental Neurobiology” ed. 2. Plenum, New York. JACOBSON, M. (1981). Rohon-Beard neuron origin from blastomeres of the 16-cell frog embryo. J. Neurosci. 1, 918-922. JACOBSON, M. (1983). Clonal organization of the central nervous system of the frog. III. Clones stemming from individual blastomeres of the 128-, 256-, and 512-cell stages. J. Neurosci. 3, 10191038. JACOBSON, M., and HIROSE, G. (1981). Clonal organization of the central nervous system of the frog. II. Clones stemming from individual blastomeres of the 32- and 64-cell stages. J. Neurosci. 1, 271-285. KATZ, M. J., and LASEK, R. J. (1978). Eyes transplanted to tadpole tails send axons rostrally in two spinal cord tracts. Science (Washington, D.C.) 199, 202-203. KATZ, M. J., and LASEK, R. J. (19’79). Substrate pathways which guide growing axons in Xenopus embryos. J. Camp. NeuroL 183, 817-832. KATZ, M. J., LASEK, R. J., and NAUTA, H. J. W. (1980). Ontogeny of substrate pathways and the origin of the neural circuit pattern. Neuroscience 5, 821-833. KEVETTER, G. A., and LASEK, R. J. (1980). Segmental pattern of the events associated with early development of marginal zones in Xenopus spinal cord. Neuroaci. Abstr. 6, 846. KINGSBURY, B. F. (1932). The “law” of cephalocaudal differential growth in its application to the nervous system. J. Comp. Neural. 56, 431-463. KULLBERG, R. W., LENTZ, T. L., and COHEN, M. W. (1977). Development of the myotomal neuromuscular junction in Xenopus laevis: An electrophysiological and fine structural study. Dev. BioL 60, 101129. LAMBORGHINI, J. L. (1980). Rohon-Beard cells and other large neurons in Xenqpus embryos originate during gastrulation. J. Camp. Neural 189, 323-333. LANCE-JONES, C., and LANDMESSER, L. (1981). Pathway selection by chick lumbosacral motoneurones during normal development. Proc R. Sot. London Ser. B 214, 1-18. LANDER, A. J., DFUJII, D. K., GOSPODAROWICZ, D., and REICHARDT, L. F. (1982). Characterization of a factor that promotes neurite

JACOBSON

AND

HUANC

outgrowth-evidence linking activity to a heparan sulfate proteoglycan. J. Cell Biol. 94, 5’74-585. LETOURNEAU, P. C. (1978). Chemotactic response of nerve fiber elongation to nerve growth factor. Dev. Biol. 66, 183-196. LETOURNEAU, P. C. (1982). Nerve fiber growth and its regulation by extrinsic factors. In “Neuronal Development” (N. C. Spitzer, ed.), pp. 213-254. Plenum, New York. LUMSDEN, A. G. S., and DAVIES, A. M. (1983). Earliest sensory nerve fibers are guided to peripheral targets by attractants other than nerve growth factor. Nature (London) 306, 786-788. MACKLIN, M., and WOJTKOWSKI, W. (1973). Correlates of electrical activity in Xenopus laevis embryos. J. Comp. Physiol. 84, 41-58. MESULAM, M.-M. (1982). “Tracing Neural Connections with Horseradish Peroxidase.” Wiley-Interscience, New York. MOODY, S. A., and JACOBSON, M. (1983). Compartmental relationships between anuran primary spinal motoneurons and somitie muscle fibers that they first innervate. J. Neurosci. 3, 1670-1682. MUNRO, M., and CRICK, F. H. C. (1971). The time needed to set up a gradient: Detailed calculations. Symp. Sot. Exp. Biol. 25, 439-453. MUNTZ, L. (1975). Myogenesis in the trunk and leg during development of the tadpole of Xenopus la&s (Daudin 1802). J. Embryo1 Exp. Mwphol. 33, 757-774. NIEUWKOOP, P. D., and FABER, J. (1967). “Norma1 table of Xenopus laevis (Daudin),” 2nd ed., North-Holland, Amsterdam. NORDLANDER, R. H. (1984). Developing descending neurons of the early Xenopus tail spinal cord in the caudal spinal cord of early Xenopua J. Comp. Neural. 228, 117-128. NORDLANDER, R., and SINGER, M. (1978). The role of ependyma in regeneration of the spinal cord in the urodele amphibian tail. J. Camp. Neural. 180, 349-374. NORDLANDER, R., and SINGER, M. (1982). Morphology and position of growth cones in the developing Xenopus spinal cord. Dew. Brain Res. 4, 181-193. NURCOMBE, V., and BENNETT, M. R. (1983). The growth of neurites from explants of brachial spinal cord exposed to different components of wing bud mesenchyme. J. Camp. NeuroL 219, 133-142. RAMON Y CAJAL, S. (1910). Algunas observaciones favorables a la hipotesis neurotropica. Trab. Lab. Invest. Biol. Univ. Madrid 8, 63134. RAMON Y CAJAL, S. (1928). “Degeneration and Regeneration of the Nervous System” (R. M. May, trans.), Hafner, New York. ROBERTS, A., and BLIGHT, A. R. (1975). Anatomy, physiology and

Pioneer

Neurite

113

Outgrowth

behavioural role of sensory nerve endings in the cement gland of embryonic Xenopua Proc. R. Sot. Land Ser. B 192, 111-127. ROBERTS, A., and CLARKE, J. D. W. (1982). The neuroanatomy of an amphibian embryo spinal cord. Phil. Trans. R. Sot. Land. Ser. B 296, 195-212. ROBERTS, A., and HAYES, B. P. (1977). The anatomy and function of “free” nerve endings in an amphibian skin sensory system. Proc. R. Sot. Lond Ser. B 196, 415-429. ROBERTS, A., and SMYTH, D. (1974). The development of a dual touch sensory system in embryos of the amphibian Xenopus laevti. J. Comp. Physiol 88, 31-42. ROBERTS, A., and TAYLOR, J. S. H. (1982). A scanning electron microscope study of the development of a peripheral sensory neurite network. J. Embryol. Exp. Morphol. 69,237-250. ROGERS, S. L., LETOURNEAU, P. C., PALM, S. L., MCCARTHY, J., and FURCHT, L. (1983). Neurite extension by peripheral and central nervous system neurons in response to substratum-bound fibronectin and laminin. Dew. Biol 98, 212-220. RUTISHAUSER, U. (1983). Molecular and biological properties of a neural cell adhesion molecule. Cold Spring Harbor Symp. Quant. Biol. 48, 501-514. RUTISHAUSER, U. (1984). Developmental biology of a neural cell adhesion molecule. Nature (London) 310, 549-554. SILVER, J., and RUTISHAUSER, U. (1984). Guidance of optic axons by a preformed adhesive pathway on neuroepithelial end feet. Dev. Biol. 106, 485-499. SINGER, M., NORDLANDER, R. H., and EGAR, M. (1979). Axonal guidance during embryogenesis and regeneration in the spinal cord of the newt; the blueprint hypothesis of neuronal pathway patterning. J. Camp. NeuroL 185, l-22. SPITZER, N. C. (1982). Voltage Rohon-Beard neurones during tadpoles. J. Physiol. (London)

and stage dependent uncoupling of embryonic development of Xenopus 330, 513-536.

VOGT, W. (1926). Uber Wachstum und Gestaltungsbewegungen hintern Korperende der Amphibien. Anat. Anz. 61, 62-75. WEISS, P. (1941). Nerve patterns: The mechanics of nerve Third Growth Symp. 5, 163-203.

am growth.

WESSELS, N. K., LETOURNEAU, P. C., NUTTALL, R. P., LUDUENAANDERSON, M., and GEIDUSCHEK, J. M. (1980). Responses to cell contacts between growth cones, neurites and ganglionic nonneuronal cells. J. Neurocytot. 9, 647-664.