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. Author manuscript; available in PMC: 2012 Nov 14.
Published in final edited form as: Dev Biol. 2009 Dec 28;339(1):114–125. doi: 10.1016/j.ydbio.2009.12.022

Vascular Endothelial Growth Factor Regulates Cranial Neural Crest Migration In Vivo

Rebecca McLennan 1, Jessica M Teddy 1, Jennifer C Kasemeier-Kulesa 1, Morgan H Romine 1, Paul M Kulesa 1,2
PMCID: PMC3498053  NIHMSID: NIHMS231099  PMID: 20036652

Abstract

The neural crest is an excellent model to study embryonic cell migration, since cell behaviors can be studied in vivo with advanced optical imaging and molecular intervention. What is unclear is how molecular signals direct neural crest cell (NCC) migration through multiple microenvironments and into specific targets. Here, we tested the hypothesis that the invasion of cranial NCCs, specifically the rhombomere 4 (r4) migratory stream into branchial arch 2 (ba2), is due to chemoattraction through neuropilin-1-vascular endothelial growth factor (VEGF) interactions. We found that the spatio-temporal expression pattern of VEGF in the ectoderm correlated with the NCC migratory front. RT-PCR analysis of the r4 migratory stream showed that ba2 tissue expressed VEGF and r4 NCCs expressed VEGF receptor 2. When soluble VEGF receptor 1 (sVEGFR1) was injected distal to the r4 migratory front, to bind up endogenous VEGF, NCCs failed to completely invade ba2. Time-lapse imaging revealed that cranial NCCs were attracted to ba2 tissue or VEGF sources in vitro. VEGF-soaked beads or VEGF-expressing cells placed adjacent to the r4 migratory stream caused NCCs to divert from stereotypical pathways and move towards an ectopic VEGF source. Our results suggest a model in which NCC entry and invasion of ba2 is dependent on chemoattractive signaling through neuropilin-1-VEGF interactions.

Keywords: VEGF, neural crest, cell migration, cranial, chick, confocal, time-lapse imaging, chemoattraction

Introduction

NCCs exit all along the dorsal neural tube and in the head are directed towards specific peripheral targets that include the branchial arches (Schilling and Kimmel, 1994; Kulesa and Fraser, 1998; Farlie et al, 1999; Kulesa and Fraser, 2000; Golding et al, 2002; Trainor et al, 2002). Prevailing models suggest that loosely connected, discrete cranial NCC migratory streams are directed from the neural tube to their specific destinations by a combination of intrinsic and extrinsic cues (Lumsden et al, 1991; Graham et al, 1993; Kulesa and Fraser, 1998; Le Douarin and Kalcheim, 1999; Kulesa and Fraser, 2000; Golding et al, 2002; Trainor et al, 2002; Teddy and Kulesa, 2004). In cell contact-based models, mechanisms such as contact inhibition of movement (Carmona-Fontaine et al, 2008) and population pressure (Newgreen et al, 1996) are thought to stimulate cell movements. When combined with instructions from the neural tube, NCC streams emerge from discrete locations of the neural tube and travel to specific branchial arches.

In contrast, some models suggest that external cues within the multiple microenvironments through which the neural crest travel, permit or inhibit cell movements to dynamically sculpt the cranial NCC migratory pattern. The explosion of molecular data on genes that appear to guide NCCs, primarily by restricting their movement to a particular migratory pathway has revealed the importance of cell-microenvironment signaling (Smith et al, 1997; Eickholt et al, 1999; Santiago and Erickson, 2002; De Bellard et al, 2003; Golding et al, 2004; Harris et al, 2008; Toyofuku et al, 2008). There is now a critical need for information about whether microenvironmental signals attract cranial NCCs towards the branchial arches and regulate entry to colonize the target microenvironment.

Prior studies have implicated neuropilins in the proper migration of NCCs throughout the head and trunk (Eickholt et al, 1999; Chilton and Guthrie, 2003; Osborne et al, 2005; Yu and Moens, 2005; Gammill et al, 2006; Gammill et al, 2007; McLennan and Kulesa, 2007; Schwarz et al, 2008; Roffers-Agarwal and Gammill, 2009; Schwarz et al, 2009a; Schwarz et al, 2009b). Both neuropilin-1 and neuropilin-2 are expressed by cranial NCCs, and have been shown to be involved in sculpting the early migratory stream of mid-rhombomere 3 (r3) to mid-rhombomere 5 (r5) NCCs, referred to as the rhombomere 4 (r4) migratory stream (Eickholt et al, 1999; Chilton and Guthrie, 2003; Osborne et al, 2005; Yu and Moens, 2005; Gammill et al, 2007; McLennan and Kulesa, 2007; Schwarz et al, 2008). Neuropilins act as co-receptors with plexins and vascular endothelial growth factor (VEGF) receptors to interact with class 3 semaphorins and isoforms of VEGF-A, respectively (Tamagnone and Comoglio, 2000; He and Tessier-Lavigne, 1997; Kolodkin et al, 1997; Soker et al, 1998; Neufeld et al, 2002). Although several different isoforms of VEGF-A exist, neuropilin-1 is a functional receptor for only the VEGF165 isoform, commonly referred to as VEGF. Neuropilin-1 interactions with Semaphorin-3A (Sema3A) or VEGF can result in opposite cellular reactions (Bagnard et al, 2001).

We have shown that neuropilin-1 signaling is critical for the invasion of the 2nd avian r4 NCC migratory stream into the branchial arch (ba2) microenvironment (McLennan and Kulesa, 2007); neuropilin-1 siRNA-EGFP (Np-1 siRNA) (Bron et al, 2004) transfected cranial NCCs failed to enter the ba2 microenvironment. Together, these recent studies demonstrate the functional diversity and importance of this family of guidance molecules and need to better understand their role in neural crest migration.

In this paper, we studied the invasion of the r4 NCC migratory stream into ba2 using the chick as a model system. First, we determined whether the spatio-temporal expression of neuropilin-1 co-receptors and ligands correlated with r4 NCC migratory events through RT-PCR, in situ hybridizations and immunohistochemistry. To address how r4 NCCs were attracted into their target microenvironment, we analyzed cell migration in response to co-cultures with ba2 tissue using a novel in vitro culture system and confocal time-lapse imaging. We used the same culture technique to determine whether VEGF influenced migration and proliferation of r4 NCCs in vitro. Using a combination of molecular perturbations to block neuropilin-1-VEGF signaling and transplantation of ectopic VEGF signaling sources, we analyzed the in vivo dynamic response of NCCs to accurately invade ba2. Our results implicate VEGF signaling through neuropilin-1 to mediate cranial NCC entry into and invasion of ba2 in a chemotactic manner.

Materials and Methods

Embryos and in ovo cell labeling

Fertilized White Leghorn chicken eggs (supplied by Placid Acre Poultry, Jasper, MO) were incubated at 38°C in a humidified incubator until the desired Hamburger and Hamilton (HH) (Hamburger and Hamilton, 1951) stages of development, and prepared for experiments as previously described (McLennan and Kulesa, 2007). NCCs were labeled in vivo as previously described (McLennan and Kulesa, 2007). The dyes used to label the NCCs were DiI and DiO (V22889, Invitrogen, Carlsbad, CA). The plasmid constructs electroporated into NCCs were pMES, which is a control EGFP vector (a kind gift from Cathy Krull, University of Michigan), and siRNA against neuropilin-1 (a kind gift from Frances Lefcort, Montana State University, originally made in the J. Cohen laboratory, MRC Centre for Developmental Neurobiology, London, UK) (Bron et al., 2004).

RT-PCR analysis

RT-PCR was used to examine the expression of neuropilin-1 co-receptors and ligands in the r4 NCCs. RNA was isolated from r4 NCCs isolated via FACS. The r4 NCCs were electroporated with an EGFP control construct, pMES (a kind gift from Cathy Krull, University of Michigan) 18 hours prior to sorting and were isolated based on GFP signal. RT-PCR was performed using primers for plexinA1, plexinA2, plexinB1, plexinB2, plexinC1, plexinD1, Semaphorin3A, Semaphorin3C, Semaphorin3D, neuropilin-1, neuropilin-2, L1-CAM, VEGF, VEGFR1, VEGFR2, VEGFR3 and beta-actin (control). Primers are listed in Table S1. Primers were purchased from Invitrogen. All resulting DNA products were approximately 500 base pairs in length. Touchdown PCR was performed as previously described (McLennan and Kulesa, 2007). The resulting PCR products were analyzed by gel electrophoresis.

In vitro culture experiments

For ba2 in vitro cultures, glass bottom dishes (P35G-1.5-20-C, MatTek Corporation, Ashland, MA) were coated with 1 mg/ml of poly-l-lysine (P6282, Sigma-Aldrich, St Louis, MO) and 1 mg/ml of fibronectin (F1141, Sigma-Aldrich). Cranial neural tubes from HH Stage 9–10 embryos were excised from the axial segment of interest (r3–r5) using a sharpened tungsten needle and isolated from surrounding tissue by 1 mg/ml dispase for 15 minutes on ice. To help with cell tracking, cranial NCCs were often pre-labeled with DiI, or electroporated with EGFP control or H2B-mRFP as previously described (McLennan and Kulesa, 2007). During this time, the ba2 tissue was excised from HH Stage 10–12 donor embryos. The neural tubes were washed in Neurobasal media (21103-049, Invitrogen) supplemented with B27 (17504044, Invitrogen) before being placed onto the coated glass bottom dishes. The ba2 tissue segments were also plated at this time and the tissue was allowed to adhere for 10 minutes at 37 °C. 1.5 mls of Neurobasal media plus B27 was then added and the cultures were returned to 37°C.

For the bead in vitro cultures, neural tubes were prepared and plated as above. Heparin-acrylic beads previously soaked in 250–500 ug/ml of VEGF165 (293-VE-010, R&D systems, Minneapolis, MN), Semaphorin 3A (1250-S3, R&D systems), or left in PBS for controls were added and 150 ul of fresh 2 mg/ml of collagen (354236, BD Biosciences, San Jose, CA) was placed over the cultures. Once the collagen set, 1.5 mls of Neurobasal media plus B27 was added and the cultures were returned to 37°C.

For the in vitro proliferation experiments, neural tube cultures were prepared as above. Neurobasal media was added directly to the cultures after the neural tubes had adhered. 1 ug/ml of VEGF165 was added to the Neurobasal media and the cultures were incubated overnight at 37°C. These cultures were then exposed to BrdU (1:100, 00-0103, Zymed, San Francisco, CA) for 1 hour, washed with PBS and fixed for 20 minutes in 4% paraformaldehyde.

In vivo injections and implantations

Soluble VEGFR1 (sVEGFR1) (400 ug/ml) (a gift from Charlie Little) was injected into the growing tissue adjacent to the r4 region of DiI-labeled HH Stage 11 embryos. 18 hours later, the embryos were harvested and fixed. The control and injected sides were both imaged using the same parameters.

For implantation experiments, DiI–labeled embryos were incubated until HH Stage 11. At this point, VEGF165, Sema3A or PBS soaked beads were implanted adjacent to the neural tube at the r3–r5 axial level using fine glass pipettes and tungsten needles. Alternatively, clumps of DiI-labeled endothelial cells (control (CRL-2279, ATCC, Manassas, VA) or VEGF-expressing (CRL-2460, ATCC)) were implanted to the same regions of unlabeled embryos. 18 hours later, the embryos were harvested and fixed. The embryos containing beads were cut in half down the midline and the intensity of DiI at the bead site was compared to the intensity of the DiI at the same site on the control side. Embryos containing clumps of cells were cut into 100 um vibratome transverse sections, stained for HNK-1 (see Immunohistochemistry section) and imaged.

In situ hybridization

In situ hybridization was performed on 25 um cryostat sections using Ventana Discovery Automated ISH System and a RiboMap Kit (Ventana). After, probes were incubated overnight, anti-digoxigenin-AP fragments (11093274910, Roche, Mannheim, Germany) diluted 1:5000 was applied to each slide for 2 hours and washed. The following day slides were rinsed in warm water with a drop of dish soap and washed twice in water. Slides were put into NBT/BCIP (11681451001, Roche) for color reaction to develop. After development, slides were washed with PBS and processed for HNK-1 staining before being coverslipped and imaged. Semaphorin-3A sense and antisense probes were used (kind gift from Elizabeth Jones and Anne Eichmann).

Immunohistochemistry

VEGF, HNK-1 and BrdU immunohistochemistry were performed on 15–25 um cryostat sections, at different stages of development, and NCC cultures using standard immunohistochemistry procedures. Anti-VEGF (SC-507, Santa Cruz Biotechnology, Inc., Santa Cruz, CA) was used at 1:50 dilution, HNK-1 (TIB-200, ATCC) at 1:500 dilution, and anti-BrdU (03-3900, Zymed) at 1:300. Alexa Fluor secondaries (Invitrogen) were used at 1:500. For the VEGF immunohistochemistry, the fluorescent signal was amplified by using a second secondary specific to the first. After staining, the slides were coverslipped with Vectashield Hard Set Mounting Medium with DAPI (Vector Laboratories, Inc, Burlingame, CA). The NCC cultures were incubated with 10 ul/ml DAPI (D9542, Sigma-Aldrich) for 5 minutes before being imaged.

Static and Time-lapse 3D Confocal Imaging

For static analysis, sagittal sections or half hindbrain mounts were prepared and imaged as previously described (McLennan and Kulesa, 2007). For in vitro time-lapse confocal imaging, cultures were chosen based on quality of tissue and distance from bead (optimally 150–300 um, average distance was 211 um). An LSM5 Pascal (Carl Zeiss Microimaging, Jena, Germany) was used to collect single plane images collected every 4 minutes. Time-lapse confocal imaging of whole embryo cultures were prepared and imaged as previously described (Kulesa and Fraser, 1998). Z-stack images were collected every 5 minutes.

Quantitative Measurements

All cell tracking and morphometric analysis were performed using Imaris (Bitplane, Saint Paul, MN) and AIM software (Carl Zeiss Microimaging). The cell tracking was performed on the first 10% of NCCs to migrate from the neural tubes. The cell directionality was determined using Imaris software, which was calculated for each track by dividing the total displacement by the total distance traveled. Statistical analyses were performed using the standard Student’s t-test, ANOVA or Duncan’s test.

Results

R4 NCCs express VEGFR2 and certain plexins

Neuropilin receptors do not contain known intracellular signal transduction domains; plexins and VEGF receptors serve as co-receptors to transduce semaphorin and VEGF signaling, respectively, through neuropilin (Nakamura et al, 1998; Tamagnone and Comoglio, 2000). We isolated migratory NCCs from HH Stage 14 embryos and examined gene expression by RT-PCR. Using this technique, we found that, apart from the neuropilins, VEGFR2 was the only VEGF signal-transducing Nrp1 co-receptor expressed by r4 NCCs (Fig. 1A). VEGFR1, VEGFR3 and L1-CAM were not expressed by r4 NCCs using this method. Plexins A1, B1 and C1 were also expressed by migratory r4 NCCs; Plexins A2, B2 and D1 were not (Fig. 1A).

Figure 1. r4 NCCs express VEGFR2 and PlexinA1 while the overlying ectoderm of the growing ba2 expresses VEGF.

Figure 1

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(A, B) RT-PCR expression analysis of VEGF receptors and plexins within r4 NCCs and VEGF within ba2 cells. For this experiment, r4 NCCs were isolated via FACS. VEGFR1, VEGFR3, L1-CAM, and Plexins A2, B2 and D1 were not expressed by r4 NCCs. (C–F) Transverse sections through different developmental stages covering r4 NCC migration with VEGF expression in the ba2 ectoderm. The scale bar is 100 um. R4 CNCC, rhombomere 4 cranial neural crest cells; ba2, branchial arch 2; NT, neural tube.

VEGF is expressed in the vicinity of r4 NCCs, in the ectoderm directly overlying the NCC migratory pathway

Expression of plexins and VEGFR2 by r4 NCCs indicated either semaphorin or VEGF signaling could be involved in 2nd branchial arch invasion. To determine which signaling pathway was active, the sema3A and VEGF ligands were examined. Using RT-PCR on isolated ba2 tissue from HH Stage 14 embryos, we showed that VEGF was expressed by ba2 at the transcript level (Fig. 1B). Next, we examined VEGF protein expression levels from HH Stage 9 through HH Stage 15, the entire period of r4 NCC migration from the neural tube into ba2. Analysis of the VEGF expression pattern at the multiple stages revealed that VEGF was expressed by the surface ectoderm directly overlaying and along the r4 NCC migratory route (Fig. 1C–F). At HH Stage 9, prior to r4 NCC emigration, VEGF was expressed by the surrounding ectoderm and endoderm tissues (Fig. 1C, green). VEGF expression was strong in the ectoderm along nearly the entire r4 NCC migratory route at HH Stages 10–11, but sparse at the distal portion (Fig. 1D, green). VEGF expression within the ectoderm increased through HH Stages 12 and 13 (Fig. 1E, green). During HH Stages 10–13, r4 NCCs undergo an epithelial to mesenchymal transition (EMT) and migrate along the migratory route adjacent to the VEGF-expressing surface ectoderm (Fig 1D, compare red and green expression patterns). VEGF expression was greatly reduced throughout the ectoderm overlying the r4 NCC migratory route by HH Stages 14 and 15, with the highest expression at the distal portion of ba2 (Fig. 1F, green). This corresponded in time to when r4 NCC migration and invasion of ba2 was nearly completed. VEGF was also expressed by the ectoderm at other rhombomere levels (Fig. S1,G–L). We performed in situ hybridizations using Sema3A antisense and sense probes on sectioned embryos at different stages, but no Sema3A expression was seen in or directly around the developing r4 NCC migratory stream during the stages of r4 NCC migration and ba2 target invasion (Fig. S1A–C). As a positive control, we examined the expression of Sema3A in the trunk region (Fig. S1,D–F) where it was expressed by the dermamyotome as previously described (Wright et al, 1995; Shepherd et al, 1996; Anderson et al, 2003; Masuda et al, 2003). Thus, expression of both the VEGF ligand and co-receptor presented the possibility for an essential role of neuropilin-1 during r4 NCC entry into ba2.

VEGF increases proliferation of cranial NCCs in vitro

To examine the influence of VEGF on cranial NCC proliferative activity, cranial NCCs were cultured in vitro to eliminate unknown proliferative agents in the environment (Fig. 2G). Isolated cranial neural tubes (r3–r5), from which NCCs emerged, were cultured overnight with either normal Neurobasal media, or Neurobasal media containing 1 ug/ml of VEGF. After a 1-hour BrdU pulse, the cultures were fixed, stained and the number of proliferating cells was counted. We found there was a significant increase in the number of proliferating NCCs from 37.6% (± 1.6%) to 44.5% (± 2.2%) when VEGF was added to the Neurobasal media (Fig. 2G).

Figure 2. Cranial NCCs are attracted to both ba2 tissue and VEGF-soaked beads in vitro.

Figure 2

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(A–D’) Selected images from confocal time-lapse imaging of in vitro cultures (n=7 static and 14 time-lapse imaging sessions for control bead, n=10 static and 8 time-lapse imaging sessions for ba2 tissue, n=13 static and 13 time-lapse imaging sessions for VEGF-soaked beads, n=19 static and 6 time-lapse imaging sessions for Sema3A-soaked beads). Black circles represent positions of a few of the first emerging NCCs at beginning of each time-lapse imaging session and colored circles represent their final positions. Both ba2 tissue and VEGF-soaked beads were attractive for NCCs while Sema3A-soaked beads were repulsive. (E) Measurements of the time for the first 10% of emerging NCCs to reach the edge of the bead/tissue on average from multiple time-lapse imaging sessions (n=71 cells for control bead, n=64 cells for ba2 tissue, n=87 cells for VEGF-soaked bead, n=21 for Sema3A-soaked bead). (F) Cell directionality calculated from in vitro cultures. *, p<0.001 for BA2 compared to control, p<0.001 for VEGF compared to control and p=0.008 for Sema3A compared to control. (G) Percentage of proliferating cells exposed in vitro to Neurobasal media only or Neurobasal media containing 1 ug/ml of VEGF (n=1823 control cells, n=1353 cells exposed to VEGF). *, p=0.013. The scale bar is 100 um. Error bars show s.e.m. NT, neural tube; t, time; hrs, hours; ba2, branchial arch 2; VEGF, vascular endothelial growth factor; Sema3A, semaphorin 3A.

Cranial NCCs are attracted to ba2 tissue in vitro

To determine whether cranial NCCs were attracted to specific target tissue in the absence of the endogenous microenvironments through which NCCs migrate, we cultured cranial neural tubes adjacent to ba2 tissue in vitro (Fig. 2B–B’). Based on the expression patterns and prior knowledge of VEGF and neuropilin-1, we anticipated that ba2 tissue actively attracted r4 NCCs. When neural tube cultures were examined 18 hours after plating, NCCs had migrated towards the ba2 tissue as opposed to NCCs uniformly spreading into any available space as observed in control cultures (neural tubes only or neural tubes plated with control PBS-soaked beads). Time-lapse analysis revealed the NCCs migrated in a highly directed manner towards the ba2 tissue versus the uniform spreading of NCCs in multiple directions observed in control cultures (compare Fig. 2A–A’ with Fig. 2B–B’). Specifically, the lead 10% of migratory NCCs reached the edge of the ba2 tissue on average after 15 hours (Fig. 2E, asterisk) whereas at the same time point, NCCs migrating in the presence of a control bead migrated only 50% of the distance between the neural tube and the bead (Fig. 2E, dashed line). Furthermore, NCCs reached the edge of the ba2 tissue with significantly higher directionality (0.477±0.02; greater than 100% increase) when compared to control cultures (0.250±0.02) (Fig. 2F).

To further test the attractive quality of the ba2 microenvironment, we plated half of a neural tube adjacent to the ba2 tissue, with the ventral side of the neural tube facing the ba2 tissue (Fig. 3A). Typically, both sides of the neural tube are plated together, resulting in migration of NCCs from either side. In this experiment, because only half a neural tube explant was plated, NCCs migrated from the dorsal neural tube, which in this assay was the side furthest from the ba2 tissue. Even under these conditions, the NCCs (DiI-labeled) migrated around the end of the cultured neural tube directly towards the ba2 tissue (Fig. 3B–F, S2B; Movie 1). Initially no NCCs were seen in the culture field of view, but after collecting time-lapse data for 6 hours, the first NCCs could be seen migrating around the end of the neural tube, from the dorsal side to the ventral side (Fig. 3B; Movie 1). As time progressed, the NCCs migrated in a highly directed manner towards the ba2 tissue in a very dense pack (Fig. 3C–E, S2B; Movie 1) and ultimately surrounded the ba2 tissue (Fig. 3F; Movie 1). When this experiment was performed without ba2 tissue, very few NCCs migrated around to the ventral side of the neural tube (Fig. S2E).

Figure 3. Cranial NCCs migrate in a highly directed manner towards ba2 tissue in vitro.

Figure 3

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(A) Schematic overview of experiment. Note, with this configuration, NCCs will only migrate from the dorsal side of the neural tube. (B–F) Selected images from a time-lapse imaging session in which the DiI-labeled NCCs diverted from the dorsal side of the neural tube and migrated directly towards the ba2 tissue. NT, neural tube; NCCs, neural crest cells; ba2, branchial arch 2; hrs, hours.

Cranial NCCs are attracted to VEGF-soaked beads and repelled by Sema3A-soaked beads in vitro

We next explored the specific molecular mechanisms underlying the attraction response of NCCs to ba2 by focusing on the role of VEGF in r4 NCC ba2 invasion. We exposed NCCs to VEGF-soaked beads in vitro (Fig. 2C–C’; Movie 3). The goal of this in vitro assay was to remove endogenous influences present in the embryo and allowed us to focus solely on the relationship between cranial NCCs and VEGF signaling. NCCs migrated towards the VEGF-soaked beads in a highly directed manner (Fig. 2C–C’, S2C; 0.366±0.01), compared to the random migration of NCCs exposed to control beads (Fig. 2A–A’, F, S2A; Movie 2). Additionally, NCCs reached the edge of the VEGF bead in less time than when they were exposed to control beads, that is, after 15 hours, NCCs were within 20% of the distance to the VEGF bead as opposed to 50% of the distance to the control bead on average (Fig. 2E). As a positive control, NCCs were also exposed to Sema3A-soaked beads (Fig. 2D–D’; Movie 4). Cranial NCCs are known to be repelled by Sema3A in vitro (Eickholt et al, 1999). As expected, NCCs actively avoided Sema3A-soaked beads and did not migrate closer than 80% of the distance to the bead on average (that is, they only moved 20% of the distance from the neural tube towards the bead; Fig. 2E). Additionally, NCCs exposed to the Sema3A-soaked bead had very poor directionality (0.130±0.03) (Fig. 2F, S2D).

NCCs fail to properly invade ba2 when VEGF signaling is disrupted in vivo

We have shown that r4 NCCs express neuropilins and fail to enter into the 2nd branchial arch when neuropilin-1 expression is knocked down (McLennan and Kulesa, 2007). However, we did not know whether this phenotype was caused by a loss of neuropilin-1-VEGF or neuropilin-1-Sema3A signaling. To investigate the in vivo role of neuropilin-1-VEGF signaling in r4 NCC migration, we microinjected sVEGFR1 into the mesoderm lateral to r4 of the developing ba2 microenvironment of HH Stage 11–12 DiI-labeled embryos (Fig. 4A–C). Introduction of sVEGFR1 bound to endogenous VEGF and inhibited neuropilin-1-VEGF signaling (Drake et al, 2000); neuropilin-1-semaphorin signaling remained intact. After 18 hours, we measured the intensity of DiI (DiI marked the migratory NCCs) in the injected ba2 and compared this to the intensity of DiI in the ba2 on the control side of the same embryo (Fig. 4B, C). Both the area covered by DiI and the distance the DiI-labeled NCCs had migrated into ba2 were significantly reduced when the tissue was injected with sVEGFR1 (Fig. 4D, E). Specifically, the area of ba2 the DiI covered was reduced to 53.1% (± 5.3%) when injected with sVEGFR1 as opposed to 66.3% (± 5.1%) and 66.2% (± 3.9%) on the control side of the same embryo and when injected with Ringers solution, respectively (Fig. 4D). Furthermore, r4 NCCs migrated 86% (± 2.9%) of the distance from the neural tube to the distal tip of ba2 when injected with sVEGFR1 as opposed to 97.6% (± 0.8%) and 99.6% (± 0.4%) the distance from the neural tube to the distal tip of ba2 on the control side of the same embryo and when injected with Ringers solution, respectively (Fig. 4E).

Figure 4. Cranial NCCs fail to enter ba2 appropriately when VEGF signaling is disrupted.

Figure 4

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(A) Schematic overview of experiment. (B, C) Static confocal images of control and sVEGFR1-injected side of same embryo, *=site of sVEGFR1 injection. DiI-fluorescence intensity is lower in the ba2 injected with sVEGFR1 when compared to the control uninjected side. (D) Quantitative measurements of the area the DiI-positive NCCs covered within ba2, 24 hours after injection (n=18 sVEGFR1 injections and n=13 control Ringers solution injections). (E) Quantitative measurements of the distance NCCs migrated from the neural tube into ba2. *, p<0.001 for distance, p=0.045 for area. P values were calculated comparing the injected to control sides of the same embryos. The scale bar is 100 um. Error bars show s.e.m. St, Hamburger and Hamilton stage; r4, rhombomere 4; OV, otic vesicle; ba2, branchial arch 2; ba3, branchial arch 3; ba4, branchial arch 4.

NCCs are attracted to VEGF in vivo

To investigate whether VEGF can attract cranial NCCs in vivo, two different approaches were taken. First, we implanted control, VEGF-or Sema3A-soaked beads lateral to r3, r4 or r5 in HH Stage 11–12, DiI-labeled embryos (Fig. 5A). We examined the resulting positions of NCCs in 3D confocal z-stacks 18 hours later and calculated the difference between the fluorescence intensity around the beads and the fluorescence intensity at the same location on the control side of the embryo. When VEGF-soaked beads were placed lateral to r3 or r5 (typical NCC-free zones), DiI-labeled NCCs were found in these regions in 31 of 37 embryos (Fig. 5C, E). Embryos implanted with VEGF-soaked beads lateral to r3 or r5 showed an average difference in fluorescence intensity between the region around the bead and the same region on the control side of the embryo to be significantly higher (23.2 ± 3.9%) than the average difference in fluorescence intensity with control beads (1.3 ± 1.0%) (Fig. 5B, D, F). That is, there is a 23% increase in DiI intensity around the VEGF-soaked beads when compared to the DiI intensity in the equivalent area on the control side of the same embryos. As a positive control, we also calculated the average difference in fluorescence intensity with Sema3A-soaked beads (−0.5 ± 0.4%), which was statistically equivalent to the control beads (Fig. 5F). For embryos where the VEGF-soaked bead was placed adjacent to r4, the average difference in DiI intensity was also significantly higher; 9 out of 15 embryos showed an increase in DiI intensity around the VEGF-soaked beads (19.2 ±4.5%) when compared to control beads (3.86 ±1.8%) (Fig. 5F).

Figure 5. Cranial NCCs are attracted to VEGF-soaked beads in vivo.

Figure 5

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(A) Schematic overview of experiment. (B, C) Static confocal images of control side and side which received a VEGF-soaked bead adjacent to r3 of the same embryo. (D, E) Higher magnification views of the control side and the side containing the VEGF-soaked bead. White circle represents placement of bead, gray circle represents area on control side which the bead side was compared to. DiI-labeled cells are present around the bead where as the same area on the control side is void of DiI-labeled cells. (F) Quantitative measurements of the difference in arbitrary fluorescence units (AFU) (the different in DiI fluorescence intensity) between control and bead side (n=20 control beads adjacent to r3/r5, n=31 out of 37 VEGF-soaked beads adjacent to r3/r5, n=8 out of 11 Sema3A-soaked beads adjacent to r3/r5, n=4 control beads adjacent to r4, n=9 out of 15 VEGF-soaked beads adjacent to r4). *, p<0.001 for VEGF-soaked beads adjacent to r3/r5, p=0.051 for VEGF-soaked beads adjacent to r4. The scale bar is 100 um. Error bars show s.e.m. St, Hamburger and Hamilton stage; r4, rhombomere 4; r3, rhombomere 3; OV, otic vesicle; AFU, arbitrary fluorescence units.

In our alternative method, we transplanted clumps of VEGF-expressing cells or control cells into similar locations in HH Stage 11–12 embryos as the bead experiments (Fig. 6A). Specifically, we transplanted VEGF-expressing cells lateral to the r3/r4 boundary and examined the resulting NCC migratory stream pattern 18 hours later in tissue sections stained with HNK-1 to label NCCs (Fig. 6B–F). Using this method, we observed an irregular NCC migratory pattern with NCCs mixed in and around the transplanted VEGF-expressing cells in 15 out of 20 embryos (Fig. 6C, E). Out of the 5 embryos that did not show a phenotype, we judged the position of the transplanted VEGF-expressing cells to be far from the r4 NCC migratory stream. In contrast, the r4 NCC migratory pattern was normal on the control side (Fig. 6C, right side of image, 6F). Clumps of control cells transplanted into the same regions showed no abnormal r4 NCC migratory patterns (Fig. 6B, D). Prior to this experiment, VEGF-expressing cells were tested for their ability to attract NCCs in vitro (data not shown).

Figure 6. Cranial NCCs are attracted to VEGF-expressing cells in vivo.

Figure 6

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(A) Schematic overview of experiment. (B–F) Static confocal images of vibratome sections of control or VEGF-expressing cells (red) transplanted rostral to the r4 NCC stream (n=12 control cells, n=20 VEGF-expressing cells). NCCs (green) are present around the VEGF-expressing cells (C, E, arrowhead) but not on the control side (C, F, asterisk) or the control cells (B, D, arrow). The scale bars are 100 um for B, C and 50 um for D–F. St, Hamburger and Hamilton stage; NT, neural tube; r3, rhombomere 3.

Confocal time-lapse analysis reveals r4 NCCs divert from stereotypical pathways to migrate towards an ectopic VEGF source in vivo

To examine how NCCs behaviors responded to sources of VEGF placed into the chick embryo microenvironment, we performed time-lapse confocal imaging on embryos in which clumps of VEGF-expressing cells were placed into the mesoderm adjacent to r3. R4 NCCs exposed to an ectopic VEGF source behaved noticeably different from control NCCs (Fig. 7; Movie 5). As the r4 NCCs underwent EMT and began their migration, some r4 NCCs were immediately attracted to the VEGF-expressing cells and diverted towards the ectopic VEGF source (Fig. 7, top row; Movie 5). The diverted NCCs then resided at the site of ectopic VEGF. Interestingly, the lead cells r4 NCCs within the rostral portion of the steam (closest to the ectopic VEGF source) migrated around the ectopic VEGF source, but then when that source was no longer immediately adjacent to the r4 NCC migratory pathway, they rerouted to stay in the vicinity of the VEGF-expressing cells (Fig. 7, asterisk, t=10 hours; Movie 5). The diverted lead cells were followed by trailing cells which populated the area surrounding the ectopic VEGF source. This behavior resulted in the single, discrete r4 NCC migratory stream being expanded to follow two pathways (Fig. 7, asterisk and arrow, t=12 hours; Movie 5). Within the first pathway, r4 NCCs followed the stereotypical migratory route (Fig. 7, arrow; Movie 5). The other pathway consisted of diverted r4 NCCs positioned adjacent to or mingled in with the VEGF-expressing cells (Fig. 7, asterisk; Movie 5). Finally, NCCs from the r1/r2 stream were also attracted to the ectopic VEGF source and migrated caudally to reach the source (Fig. 7, arrowhead; Movie 5). All of these novel behaviors resulted in cranial NCCs surrounding the ectopic VEGF source. R4 NCCs on the control side of the embryo migrated to form the stereotypical r4 NCC migratory stream (Fig. 7, bottom row; Movie 5). Furthermore, the r4 NCCs on the control side migrated further towards the target ba2 microenvironment (with some NCCs leaving the field of view) than the r4 NCC stream that was exposed to the ectopic VEGF source (Fig. 7, compare top row to bottom row; Movie 5).

Figure 7. r4 NCCs actively reroute towards an ectopic VEGF source in vivo.

Figure 7

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Selected images from a typical time-lapse imaging session showing cranial NCCs in the r4 migratory stream (green). A clump of VEGF-expressing cells (red) was transplanted into the mesoderm adjacent to rhombomere3. (Top row) The majority of DiO-labeled r4 NCCs initially undergo an EMT and migrate towards the forming ba2 in a normal manner. However some r4 NCCs (asterisk), as well as NCCs from the rhombomere 1/2 stream (arrowhead) migrate towards the VEGF source. By t=10 hours, r4 NCCs have actively changed direction and turned to go towards the implanted VEGF-cells. At t=14 hours, a bifurcated r4 stream is visualized (asterisk), with a large portion of the stream now surrounding the VEGF-cells. (Bottom row) DiO-labeled r4 NCCs are in the same embryo as the top row, but are not exposed to a new source of VEGF. The unaffected r4 NCCs migrate in a more directed manner towards their target site and at each time-point, they have migrated further away from the neural tube than on the experimental side. The scale bars are 100 um. r2, rhombomere 2; r3, rhombomere 3; r4, rhombomere 4; hrs, hours.

To confirm that cranial NCCs were attracted to VEGF via neuropilin-1 signaling, we knocked down the expression of neuropilin-1 in the cranial NCCs and repeated the above experiment. When cranial NCCs were transfected with Np-1 siRNA, they no longer sensed the ectopic source of VEGF and their stereotypical pathways were maintained (Fig. S2F). The lead Np-1 siRNA transfected r4 NCCs did not divert from the original path and remained the lead NCCs throughout the time-lapse (Fig. S2F, asterisk). Furthermore, r2 NCCs migrated directly past the ectopic VEGF source (Fig. S2F, arrowhead). Thus, this phenotype was not only true for the r4 NCCs but also the NCCs in the r1/r2 stream (Fig. S2F).

Discussion

We used the chick as a model system to study cranial neural crest migration into the branchial arches. Our results suggest specific guidance signals regulate neural crest cell behaviors to direct the r4 migratory stream into the 2nd branchial arch. First, we show plexins and VEGFR2 are expressed by r4 NCCs (Fig. 1), indicating either semaphorin or VEGF signaling was involved in 2nd branchial arch invasion. To determine which signaling pathway was active, the Sema3A and VEGF ligands were examined. We show that VEGF is expressed in the vicinity of r4 NCCs, in the ectoderm directly overlying the r4 NCC migratory pathway (Fig. 1). Expression of both the VEGF ligand and co-receptor, and absence of Sema3A expression in the vicinity of ba2, presented the possibility for an essential role of neuropilin-1 during r4 NCC migration into ba2. This reflected a role for VEGF signaling through neuropilin-1. Injections of sVEGFR1, to block endogenous VEGF signaling, made distal to the invading r4 stream resulted in a decrease in the ability of r4 NCCs to invade ba2 (Fig. 4). In both in vitro and in vivo experiments, VEGF attracted neuropilin-1 expressing cranial NCCs (Figs. 2, 3) and caused cranial NCCs to divert from stereotypical migratory pathways (Figs. 5, 6, 7), respectively.

R4 NCCs not only expressed neuropilin-1, but also a number of neuropilin-1 co-receptors, indicating that r4 NCCs could interact with VEGF via VEGFR2 or class 3 semaphorins via plexin A1 (Fig. 1A). Previous studies have demonstrated that VEGFR2 mediates the majority of VEGF responses by cells (Oosthuyse et al, 2001; Robinson and Stringer, 2001; Murga et al, 2005; Olsson et al, 2006). Therefore, r4 NCC expression of VEGFR2 was consistent with VEGF-mediated migration. Plexins are the co-receptors for semaphorins, but only plexin As and plexin D1 interact with the secreted semaphorin-3 class (Kruger et al, 2005), and plexin A1 has been shown to be necessary to achieve Sema3A-mediated repulsion of growth cones (Giger et al, 2000). With this in mind, plexin A1 was the most likely candidate to facilitate neuropilin-1-semaphorin3 interactions in the r4 NCCs. Our expression analysis was consistent with previous studies that showed expression of plexin A1 transcript by r4 NCCs in St 11 (HH) chick (Osborne et al, 2005). This result highlighted the differences between NCC expression at distinct axial levels; chick cardiac NCCs express plexins A2 and D1, but not plexin A1 (Toyofuku et al, 2008). Together, this led us to consider that r4 NCC entry into ba2 might rely on neuropilin-1-VEGFR2-VEGF or neuropilin-1-plexin A1-semaphorin interactions.

VEGF protein was expressed in the surface ectoderm overlying the cranial NCC migratory pathways, placing VEGF in close proximity to r4 NCCs all along their migratory route. The pattern of VEGF expression was distributed homogeneously in the ectoderm, throughout the timeframe when r4 NCCs migrated towards ba2 (Fig 1). However, VEGF expression became localized to the ba2 region when lead r4 NCCs reached ba2, and was decreased within the ectoderm in the proximal direction back towards the dorsal neural tube (Fig. 1, 8). It has been shown that cranial NCC contact with the surface ectoderm plays an important role in inhibiting cell invasion into typical NCC-free zones (Golding et al., 2002). Although our expression analysis was consistent with a previous study of vasculogenesis that reported VEGF expression by the chick head ectoderm (Anderson-Berry et al, 2005), it was not clear whether the tissue directly overlying the r4 NCC migratory stream played a role for permissive guidance. We found the pattern of expression of VEGF correlated with the advancement of the r4 NCC migratory front, but whether NCCs would respond in a directional manner to either ba2 tissue or VEGF in the absence of endogeneous microenvironments through which the cells migrate was not clear.

Figure 8. Ba2 invasion of r4 neural crest cells is directed by neuropilin-1-VEGF interactions.

Figure 8

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(A) Transverse schematic of the r4 NCC migratory stream with the corresponding neuropilin-1 (blue) and VEGF expression (purple). VEGF expression plays a key role at the ba2 microenvironment entrance to facilitate correct NCC colonization of the ba2 microenvironment. (B) A sagittal view of a wildtype r4 NCC migratory stream. (C) When VEGF function is perturbed via soluble VEGFR1 or neuropilin-1 expression is knocked down via Np-1 siRNA perturbed, NCCs fail to properly enter the ba2 microenvironment. (D) When an ectopic source of VEGF (orange) is transplanted into the area adjacent to r3, NCCs are rerouted towards it. Abbreviations: r4, rhombomere 4, NT, neural tube.

VEGF increased proliferation and attracted cranial NCCs in vitro, suggesting a role for VEGF in both NCC proliferative activity and chemotaxis behavior. We found that although cranial NCCs invaded in vitro, there was a clear difference in cell behaviors in the presence of ba2 tissue, VEGF-, sema3A-, or control-soaked beads (Fig. 2; Movies 1–4). Cranial NCCs displayed high directionality (Fig. 2F; distance from start to finish/total distance traveled), reached the vicinity of a VEGF-soaked bead or ba2 tissue much faster than control or sema3A soaked beads, and sustained polarized directional migration to interact with the VEGF-soaked bead or ba2 tissue (Fig. 2A–D’; Fig. 3; Movies 1–4). Consistent with our observations of an increase in NCC number, VEGF has been shown to act as a proliferative agent for other cell types, including endothelial cells, Schwann cells, neural progenitor cells, as well as cortical neurons, astrocytes and microglia, neural stem cells and medulloblastomas (Neufeld et al, 1999; Murga et al, 2005; Sondell, et al, 1999; Bagnard et al, 2001; Zachary, 2005). Interestingly, we have shown in vivo that lead r4 NCCs proliferate three times as much as trailing NCCs (Kulesa et al, 2008). It will be productive to determine whether microenvironmental factors, such as VEGF, stimulate NCC proliferative activity or whether these differences are intrinsic to the lead NCC subpopulation. Our results were consistent with previous work demonstrating VEGF as a chemoattractant for various migratory cell types, such as neural progenitor cells, facial branchiomotor somata, and endothelial cells (Bagnard et al, 2001; Zhang et al, 2003; Schwarz et al, 2004; Barkefors et al, 2008). Therefore, our data implied that without any other influences, VEGF plays a dual role to facilitate chemotactic migration and proliferation of cranial NCCs in vitro. The next step was to test whether loss of VEGF function would affect r4 NCC migration and entry into ba2.

NCCs failed to migrate properly into ba2 when VEGF signaling was disrupted by microinjection of sVEGFR1 (soluble) into the mesoderm lateral to r4 (Fig. 4C–E). This phenotype was strikingly similar to the NCC phenotype when neuropilin-1-ligand (VEGF and semaphorins) signaling was disrupted via Np-1-Fc injection (McLennan and Kulesa, 2007). The phenotype in each case was less severe than reduction of neuropilin-1 function by siRNA (McLennan and Kulesa, 2007), however this may have been due to diffusion of the sVEFGR1 and growth of the tissue that act to dilute the local VEGF influence.

VEGF was expressed in the ectoderm directly overlying the r4 NCC migratory pathway, however no significant phenotype was seen when r4 NCCs were transfected with Np-1 siRNA (McLennan and Kulesa, 2007), or Np-1-VEGF signaling was blocked with sVEGFR1, until NCCs encountered the ba2 microenvironment. There are at least two possible explanations for this. First, the proper position of migratory r4 NCCs near the neural tube, and migration in a discrete stream, involves multiple processes (Trainor et al., 2002; Golding et al., 2002), including semaphorin/neuropilin signaling (Osbourne et al., 2005; Gammill et al., 2007). Cranial NCCs contact and receive guidance cues from the surface ectoderm near the neural tube (Golding et al., 2002), so VEGF expression in the ectoderm overlying the r4 migratory pathway may simply reinforce the permissive nature of the local microenvironment and other cues provide directional bias. Alternatively, the growing ba2 forms a pouch-like structure into which the r4 NCCs migrate, and as this structure forms, the surrounding ectoderm expresses VEGF (Fig. 1). Proximal to ba2, the ectoderm overlies the r4 NCC migratory pathway and expression of VEGF is homogeneously distributed. Expression of VEGF is not down-regulated in the proximal ectoderm until HH St 15, when the NCC migratory front is near the ba2 entrance. Therefore, in order for VEGF signals to direct early NCC migration, cells in direct contact with VEGF expression would have to sense a gradient and communicate direction information to more ventrally located NCCs.

Our results did not rule out a role for VEGF as a NCC chemoattractant from the neural tube to the 2nd branchial arch entrance, however in the absence of a proximal-to-distal gradient of VEGF, we suggest the mechanisms that underlie early neural crest migration may be more complex. That is, how might migratory cells respond in a chemotactic manner to a guidance molecule expressed in a homogeneous distribution? Recent hypotheses in comparable embryonic cell migratory populations (primordial germ cells and lateral line) both suggest that although the ligand (SDF-1) is expressed in a homogeneous distribution, a source-sink mechanism (and chemotactic response) is achieved by the differential response of cells to SDF-1. Work in the Raz lab has recently demonstrated that somatic cells expressing a second SDF-1/CXCL12 receptor, CXCR7, show enhanced internalization of the chemokine suggesting that CXCR7 acts as a sink for SDF-1a (Boldajipour et al., 2008). Whether this type of model mechanism is involved in cranial neural crest migration is not clear and for future study. However, it does provide a possible explanation for how migratory cells respond in a directed manner to a homogeneously distributed guidance signal.

VEGF positively influenced NCC behaviors by directing cells towards ectopic VEGF sources and away from stereotypical migratory routes. When VEGF-expressing cells or VEGF-soaked beads were placed in the embryo, there was an increase in the number of NCCs found around the VEGF source when compared to controls (Fig. 5–7). Time-lapse analysis of NCC behaviors revealed cells changed direction away from stereotypical migratory pathways adjacent to r2 and r4 (Fig. 7; Movie 5). Diverted NCCs moved towards and encountered ectopic VEGF sources (Fig. 5–7), suggesting VEGF chemoattraction could overcome local inhibitory cues. However, when neuropilin-1 expression in the cranial NCCs was knocked down, the migration of NCCs was no longer influenced by the presence of an ectopic VEGF source (Fig. S2F). Together, this suggested that the chemotactic response by r4 NCCs to a VEGF source was due to neuropilin-1-VEGF interactions.

In summary, our findings implicate VEGF as the ligand to neuropilin-1 that permits proper entry and migration of NCCs into the 2nd branchial arch (Fig. 8). We suggest that this is the first in vivo evidence in support of a chemoattraction mechanism to direct cranial NCC entry into and invasion of the branchial arches. These results compliment previous studies that show other factors influence the early r4 NCC migratory stream shape and maintenance, and help us to better understand how NCCs invade precise targets. NCCs respond to multiple guidance signals from the microenvironment and each other. Our findings do not rule out mechanisms of contact inhibition of movement (Carmona-Fontaine et al, 2008) and population pressure (Newgreen et al, 1996) to stimulate NCC movement. However, we suggest an underlying molecular mechanism of chemoattraction provides a directional bias for NCCs at the entrance into the 2nd branchial arch. It will now be important to investigate how molecular signals regulate NCC behaviors within different microenvironments through which the cells migrate and how local cell behaviors give rise to the global cell migratory stream pattern.

Supplementary Material

01

Acknowledgements

The authors would like to thank Clement Colleau for providing the initial methodology for the cell behavior analysis and Dr. Elizabeth Jones and Dr. Anne Eichmann for in situ probes. We kindly thank Cameron Cooper, Katie Hollander and Danny Stark for imaging assistance, Jeff Haug and his colleagues in Cytometry, Teri Johnson and Beth DeGarmo in Histology and Hua Li in Bioinformatics at the Stowers Institute. This work was funded by NIH Grant 1R01HD057922 (PMK) and the Stowers Institute for Medical Research.

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