Extracellular matrix dynamics during vertebrate axis formation

Developmental Biology 268 (2004) 111 – 122 www.elsevier.com/locate/ydbio

Extracellular matrix dynamics during vertebrate axis formation

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Andra´s Cziro´k, a,b Brenda J. Rongish, a and Charles D. Little a,* a

Department of Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, KS 66160, USA b Department of Biological Physics, Eo¨tvo¨s University, Pa´zma´ny Stny 1A, Budapest 1117, Hungary Received for publication 30 April 2003, revised form 29 September 2003, accepted 30 September 2003

The authors dedicate this article to the memory of John Phillip Trinkaus, Ph.D. (1918 – 2003). Trink helped us, and thousands of other developmental biologists, understand the ‘‘forces that shape the embryo.’’ While a postdoc, one of us (C.D.L.) had the great fortune to have a desk next to Trink during his visiting professorship in Jon Singer’s Lab (UCSD, 1981)—it was a glorious experience.

Abstract The first evidence for the dynamics of in vivo extracellular matrix (ECM) pattern formation during embryogenesis is presented below. Fibrillin 2 filaments were tracked for 12 h throughout the avian intraembryonic mesoderm using automated light microscopy and algorithms of our design. The data show that these ECM filaments have a reproducible morphogenic destiny that is characterized by directed transport. Fibrillin 2 particles initially deposited in the segmental plate mesoderm are translocated along an unexpected trajectory where they eventually polymerize into an intricate scaffold of cables parallel to the anterior – posterior axis. The cables coalesce near the midline before the appearance of the next-formed somite. Moreover, the ECM filaments define global tissue movements with high precision because the filaments act as passive motion tracers. Quantification of individual and collective filament ‘‘behaviors’’ establish fate maps, trajectories, and velocities. These data reveal a caudally propagating traveling wave pattern in the morphogenetic movements of early axis formation. We conjecture that within vertebrate embryos, long-range mechanical tension fields are coupled to both large-scale patterning and local organization of the ECM. Thus, physical forces or stress fields are essential requirements for executing an emergent developmental pattern— in this case, paraxial fibrillin cable assembly. D 2004 Elsevier Inc. All rights reserved. Keywords: Gastrulation; Fate map; Extracellular matrix; Time-lapse; Dynamics; Tissue deformation

Introduction Gastrulation and vertebral axis formation require extensive changes in the shape of the early vertebrate embryo. In avians, after developmental stage 4 (Hamburger and Hamilton, 1951), the primitive streak gradually regresses and Hensen node moves caudally. A concomitant process, somitogenesis, leads to the parsing of discrete segments of mesodermal tissue into small packets at regular intervals along the anterior – posterior axis, thereby establishing the segmented vertebrate body plan. Neurulation, which includes formation and extension of the notochord, is another

$ Supplementary data associated with this article can be found, in the online version, at doi: 10.1016/j.ydbio.2003.09.040. * Corresponding author. Department of Anatomy and Cell Biology, University of Kansas Medical Center, 3901 Rainbow Boulevard, Kansas City, KS, 66160. Fax: +913-588-2710. E-mail address: [email protected] (C.D. Little).

0012-1606/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.ydbio.2003.09.040

contemporaneous event in vertebrate development. The notochord is laid down as a ‘‘rod’’ of mesoderm by Hensen’s node, and elongates as the primitive streak regresses. The neural tube develops dorsal to the notochord; subsequently, the notochord and neural tube derivatives become encased in the vertebrae. Large numbers of cells move to new locations during these changes in embryonic anatomy. Recent cell fate mapping studies of gastrulation/neurulation – stage avian embryos established that mesodermal precursor cells first dislocate medially towards the primitive streak. Their ingression at the axis is followed by various degrees of lateral displacement, targeting the notochord, somites, or the segmental mesoderm depending on the position and time of ingression (Bortier and Vakaet, 1992; Catala et al., 1996; Psychoyos and Stern, 1996; Sawada and Aoyama, 1999; Schoenwolf et al., 1992; Yang et al., 2002). While the cellular fate map of gastrulation is well established, the mechanisms underlying the morphogenetic movements are

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still far from clear (for recent reviews, see Wallingford et al. (2002); Keller et al., 2003). It has been long understood that the extracellular matrix (ECM), which is known to provide signals to cells and affect cell behavior, plays an important role in morphogenesis. However, the ECM is not simply a static scaffold: cell motion involves cell traction, a mechanical interaction between cells and the ECM, which, in turn, results in a local realignment of the ECM (Oliver et al., 1995; Stoplak and Harris, 1982; Tranquillo, 1999). ECM fibers can also be reoriented by long-range (nonlocal) mechanical forces within the tissue (Olsen et al., 1999). Nevertheless, to the best of our knowledge, in vivo ‘‘fate maps’’ of ECM component movements have not been determined—nor has it been established whether ECM molecular complexes, like cells, ‘‘behave’’ in a predictable, developmental stage- and position-dependent fashion. Proteins of the fibrillin family are promising candidates for ECM tracking studies because they are ubiquitous and easily recognizable, discrete filamentous structures when made visible by immunolabeling methods (Burke et al., 2000; Gallagher et al., 1993; Rongish et al., 1998; Visconti et al., 2003; Wunsch et al., 1994). Fibrillin 1 and 2 are constituents of connective tissue microfibrils (Sakai et al., 1986; Zhang et al., 1994) and are both observed in the avian embryo before gastrulation in the form of slender filaments marking the future anterior – posterior axis (Gallagher et al., 1993; Wunsch et al., 1994). During gastrulation, the fibrillins continue to display a remarkable pattern widely distributed in association with the mesoderm (Burke et al., 2000; Rongish et al., 1998; Visconti et al., 2003). The importance of these proteins to the structural and functional properties of connective tissues has been demonstrated by the finding that mutations in the human FBN1 and FBN2 genes result in heritable disorders, with patients manifesting ocular, skeletal, and cardiovascular abnormalities (Dietz et al., 1994). In accord, fibrillin-deficient mouse models demonstrated that the absence of normal levels of either fibrillin 1 or 2 protein is incompatible with the proper function of mature elastic fibers (Artega-Solis et al., 2001; Pereira et al., 1997; Pereira et al., 1999). Deficiency in one type of fibrillin was found to be compatible with gastrulation of mouse embryos; nevertheless, exogenous expression of a truncated fibrillin in the presumptive dorsal mesoderm of gastrulation stage frog embryos disturbed endogenous fibrillin localization and blocked gastrulation (Skoglund, 1996). Motivated by the excellent tracking properties and the developmental importance of fibrillin fibers, we investigated the reorganization and assembly of this ECM component during gastrulation/neurulation. We demonstrate the coalescence of an extended, loose meshwork into larger cables. Our high-resolution map of filament displacements shows that morphogenesis-associated ECM movements essentially repeat the same basic pattern at progressively more caudal positions. These quantitative data open the door to future studies focusing on the mechanical forces that drive mor-

phogenesis. We conclude with the hypothesis that largescale ECM reorganization is primarily driven by changing mechanical tension fields.

Materials and methods Microinjection The location of fibrillin 2-containing ECM was monitored with Cy3 (Amersham Pharmacia Biotech Inc., Piscataway, NJ) fluorochrome-conjugated monoclonal JB3 IgG antibodies (Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA). Delivery of these nonperturbing (Rongish et al., 1998) marker antibodies was accomplished by injecting 25-nl volumes, one to four times per embryo, at 1 ng/nl concentrations (Pico-Injector, Harvard Apparatus Inc, Holliston, MA, mounted on a micromanipulator, Narishige Scientific Instrument Lab, Tokyo, Japan). Embryo culture As detailed in Little and Drake (2000), quail embryos (Coturnix coturnix japonica, John Smith Farms, Bucyrus, KS) at Hamburger and Hamilton (HH) stages 4 – 5 were encircled with paper rings that adhere to the embryo’s vitelline membrane and then excised from the egg. After removing the adherent yolk by washing with sterile embryo phosphate-buffered saline (EPBS), the embryos are placed on a solid agarose bed (5% agarose in EPBS) for microinjection. After introduction of the antibodies, the embryos are placed in a custom designed, microscope-attached incubation chamber (Czirok et al., 2002; Rupp et al., 2003). The wells of a suitably modified Costar polystyrene 6 well plate (Corning Inc., Corning, NY) are filled with 3 ml of CO2independent culture medium (Leibovitz-L15 medium supplemented with 2 mM L-glutamine, 10% chicken serum, and 1% penicillin – streptomycin, GibcoBrl, Grand Island, NY) in which a Millicell (Millipore Co., Bedford, MA) tissue insert is placed. The plastic filter material of the insert was replaced with a sparse array of polypropylene surgical suture fibers (7– 0 Surgipro, USSC, Norwalk, CT). Embryos are cultured under sterile conditions, at the medium –air interface, ventral side-up, with their vitelline membrane in contact with the suture mesh. Their normal development was established by the lack of morphological defects as well as by the timely addition of somites and elongation of the notochord (Rupp et al., 2003). Microscopy and image processing As described in detail in Czirok et al. (2002), an automated Leica DMR upright microscope is used in DIC and epifluorescence modes with a long working distance 10
(0.25 NA) objective. Fields of 875
688 Am are imaged with a cooled Quantix CCD camera (Photometrics,

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Tucson, AZ) at a spatial resolution of 658
517 pixels, 12 bit light intensity resolution, and typically 100 ms exposure times. In each period of the imaging cycle, 2– 4 embryos are sequentially positioned under the objective. For each embryo, a rectangular area of 1
4 or 2
4 slightly overlapping microscopic fields are scanned, and each of these fields is recorded at multiple (10 – 12) focal planes, separated by 20 Am, first in DIC and immediately thereafter in epifluorescence mode. The acquisition of the ‘‘z-stack’’ pairs is accomplished within a short period of time (typically a minute) ensuring the correct spatial registration of the corresponding DIC—epifluorescence image pairs. The practical result of this technology is that every position of the embryonic plate lies in one of the focal planes; thus, no feature can be lost. Embryos change their position considerably during development, and this motion must be compensated to keep the specimen centered in the field of view. This is accomplished in two stages: First, during recording, the stage positions are periodically (about every 2 h) revised either manually or using software-generated feedbacks. During the processing of the stored images, a second, automatic frameby-frame realignment is performed to eliminate the drift and sudden shifts of the object. These tasks, the merger of the neighboring microscopic fields into a composite image, and the local contrast enhancement of the immunofluorescence data were achieved using our algorithms (Czirok et al., 2002), which are available to download at http://www. kumc.edu/mic-utils. Laser scanning confocal microscopy was performed as described in Rongish et al. (1998). Tracking of image details A modified version of a program used in previous cell tracking studies (Hegedu¨s et al., 2000) allowed us to browse through the stored images (time-lapse ‘‘z-stacks’’) and follow various details, such as fluorescent filament segments, 5 –10 Am in size, through consecutive images. Although this tracking procedure is performed in three dimensions (using each optical section), due to the large depth of field of the microscope objective, the resolution along the vertical (z) direction is rather limited. Therefore, here we confine our studies to the two-dimensional (x – y) projections of the obtained position data. In the following, two-dimensional vector quantities are denoted by boldface letters. The data analyses are detailed in the Supplementary Material.

Results Fibrillin immunolabeling Extracellular fibrillin 2 was labeled in quail embryos by the microinjection of fluorochrome-conjugated JB3 monoclonal, bivalent, IgG antibodies. After the antibody micro-

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injection at HH stages 4 –5, embryos were recorded until HH stage 10 with scanning time-lapse microscopy (Table 1). Within 1 h, the JB3 antibodies reach a substantial portion (up to 50– 70%) of the area pellucida (Fig. 1). The fluorescence intensity profile, that is, the extent of areas with various fluorescence intensities, becomes stable after the initial diffusion period (data not shown). This stability indicates very low rates for photo bleaching or other antibody degradation, since these processes would be expected to eliminate brightly stained foci. The resulting labeling is specific, as demonstrated by experiments with microinjected control (non-ECM-binding) antibodies, including the avian endothelial cell marker QH1 (Czirok et al., 2002) and mouse mAb G that recognizes avian h1 integrins (Drake et al., 1991). Further, the labeling pattern observed in JB3-microinjected embryos was compared to that in postexperimental whole-mount immunostaining of fixed embryos (Fig. 2D) and the pattern appears identical (Rongish et al., 1998). In contrast, this pattern is strikingly different from that observed following microinjection of antibodies to other ECM proteins, including tropoelastin and fibulin 1 (Visconti et al., 2003), as well as fibronectin, collagen IIA, and latent TGF-h binding protein 1 (data not shown). Occasionally some highly fluorescent precipitates remain near microinjection sites, which exclude these areas from further analysis (star in Fig. 1). The perturbative effect of microinjected JB3 antibodies was assayed by comparing whole-mounted JB3-injected embryos to non-injected controls after 10 h of development. No observable differences were found either in the postfixation JB3 staining pattern or in the anatomy of the embryos. The timing of major morphogenic processes were also the same in JB3-injected embryos and non-injected controls (n > 20). In particular, the somites form on schedule, every 90 min, the heart loops and commences beating normally (data not shown). Changes in fibrillin organization During the recorded 8- to 12-h-long period, the fibrillin 2 immunolabeling pattern changes remarkably. As Fig. 1

Table 1 Summary of the experimental data Embryo

1. 2. 3. 4. 5. 6. 7. 8. 9.

# of somites at

Observation

Injection

Start

End

Duration

Frames

Fields

PS PS PS 5 4 PS PS PS PS

3+ 2+ 2 6+ 5 2 4 3+ 1

8 10 9 12 9 7+ 9+ 8+ 8

6 h 55V 10 h 30V 11 h 10V 8 h 40V 6 h 55V 9 h 30V 9 h 30V 9 h 30V 11 h 10V

26 42 108 30 26 33 33 33 108

4 4 4 4 4 4 4 4 4












1 2 2 2 1 2 2 2 2

# of features 1324 2665 3074 864 877 – – – –

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Fig. 1. Relevant developmental stages of a quail embryo. The images in panels (a, aV), (b, bV), and (c, cV) display corresponding epifluorescence-DIC image pairs, taken at 3-h intervals, following the microinjection of Cy3-JB3 anti-fibrillin 2 antibodies. The DIC images show the characteristic changes in embryonic anatomy, most notably the elongation of the notochord (nc), formation of the anterior intestinal portal (aip), neural folds (nf), and somites. The progressively longer red arrows demonstrate the elongation of the embryonic axis and addition of somites during the 6-h elapsed time. Positions of the first and the newest somites are marked on one side of the embryo with an asterisk and a caret, respectively. The green immunofluorescence patterns reveal substantial rearrangements of the fibrillin 2-containing ECM. Most notable are parallel cables just lateral to the notochord (brackets, panels a, b, c). These fibrillin cables enclose the somites and also extend into the cranial aspect of the segmental plate mesoderm (sp, panel cV); this is readily observed at the interval between the caret and the right bracket in panels b and c. Also, an array of curved cables connect the aip to the somitic field (parentheses, panels b, c). The meshwork of fibrillin 2 that ensheaths the notochord progressively diminishes at the level of the last two somites (white arrows). The overexposed area (black star, panels a, b, c) is precipitated material at the microinjection site.

demonstrates, a fibrillin 2-containing ECM meshwork encases the notochord. At HH stage 7, a bundle of fibrillin fibers connect the AIP to the somitic regions, reaching the first few somite pairs. Most notable are fibrillin 2-containing fibers that assemble parallel to the embryonic axis. The cranial portion of these bundles encloses the somites, while caudally they extend more than 200 Am into the segmental plate mesoderm and approach Hensen node. Caudal to the node, fibrillin 2 exhibits a different pattern, which consists of punctate, disconnected fluorescent foci. Fig. 2 shows the somitic array and the segmental plate mesoderm in greater detail. These images reveal an oriented, dense array of paraxial filaments, remarkably different from the loose meshwork-like organization characteristic of the more lateral regions. (See supplementary data, movie 1.) Filaments retain their identity Figure 3 demonstrates the progressive sequence of incremental changes that constitutes the tissue-scale ECM reorganization. Filament shapes and positions remain fairly similar between consecutive image pairs; these observations

form the basis of our tracking procedure. As multiple focal planes were recorded with our multi-field scanning timelapse (4D) microscopy system, no intraembryonic object is lost or rendered out of focus. Thus, one can unambiguously follow filaments, frame-by-frame, over the recorded time. Note that although only minor alterations are seen in the ECM meshwork between short (30 min) time intervals in Fig. 3,while tracking filaments over a several hours reveals substantial changes in ECM organization (see circled structures in Fig. 2). Filament trajectories reveal substantial relocation Five embryos were selected for a systematic tracking procedure in which filaments were identified and followed frame-by-frame, resulting in a database of spatial and temporal coordinates (Table 1). No disassembly was observed within the sample of more than 200 traced filaments. Changes in the obtained coordinates can reflect both genuine alterations in embryonic anatomy, as well as anatomically irrelevant movements of the whole specimen relative to the microscope objective. To eliminate the latter, intersomitic

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Fig. 2. Fibrillin 2 reorganization in the axial areas. The individual images in panels (a, aV), (b, bV), and (c, cV) depict corresponding areas outlined by the yellow boxes in Fig. 1. Selected filaments, marked with the colored circles, visibly alter their relationship with nearby filaments and the axis. Despite this motion, the filaments retain connectedness and shape to some degree. Note that a ‘‘new’’ filament (orange circle), absent in panel aV, appears in the next two images—and is shown to move medially. The blue boxes denote the formed somites. Panel d shows the somitic array at a higher magnification. This double-labeled image was collected at the termination of a time-lapse recording, when a second, whole-mounted JB3 immunostaining was performed. The red channel contains the labeling pattern generated by the microinjected Cy3-JB3, while the second, whole-mounted Cy2-JB3 staining yields the green channel. The yellow color indicates staining by both labeling methods. Panel e shows a high-resolution laser scanning confocal microscope (LSCM) image of a comparable area in a specimen prepared for ideal immuofluorescence quality. The LSCM image is a projection of 10 focal planes into a single field. Comparing our double-labeled whole-mount wide-field epifluorescence image with the LSCM micrograph reveals a surprising amount of similar detail, including fluorescent filaments on the order of 1 Am in size.

cleft positions were extracted from DIC images and used as anatomical reference points. For each filament, a mathematical procedure, described in the Supplementary Material, resulted in its trajectory, i.e., its position on each frame relative to the somitic array. Thus, changes in these positions indicate anatomically relevant filament displacements. Fibrillin 2 trajectories, obtained from multiple embryos, reveal that the ECM is substantially relocated during development. Filament motion seems to be ordered and its tendencies highly reproducible: trajectories of adjacent filaments are similar. The direction of motion is medial in the lateral plate mesoderm, while the direction is caudal in the segmental plate mesoderm near (and posterior to) Hensen node (Fig. 4). Craniolaterally to the first somite pair, filament trajectories reflect that ECM is pulled toward the forming AIP. However, the tendency of ECM motion is time-dependent: early trajectory segments can be perpendicular to later segments of the same and nearby filaments (boxed area in Fig. 4).

Velocities characterize anatomical changes Trajectory plots are tools commonly used to visualize rearrangements in a given dynamic sample. Nevertheless, the actual trajectory shapes strongly depend on the particular timing and duration of the analyzed recording, as well as on the locations of available tracer particles. Local tendencies of motion during a short period are much better suited for quantitative, statistical comparison of different embryos, or various developmental stages of the same embryo. In this study, we subdivided the observational period into 1-h-long segments and determined filament displacements during these time intervals. The obtained displacements give the distance between filament positions at the beginning and at the end of the time segment. The displacement during a short period is proportional to the velocity; thus, we use velocities instead of trajectories as the fundamental descriptor of embryonic rearrangements. As we noted in Fig. 4, displacements of nearby filaments

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Fig. 3. A time-lapse sequence of ECM remodeling at high resolution. The four panels show the movements and rearrangements of two selected fibrillin 2 filaments (circle and square) during a 90-min interval. Despite changes in overall ECM organization, filaments can be identified with high fidelity in the consecutive frames.

are similar. This local average filament velocity (both magnitude and direction) within embryo a at time t around a certain location (x, y) is denoted as the Va(x, y, t) velocity field (or snapshot). Traveling wave-like velocity pattern The velocity field V can be described as a traveling-wave V a ðx; y; tÞcUðx; y  Ya ðtÞÞ

ð1Þ

where Ya(t) is the position of the somitogenesis front along the cranial –caudal axis and U is a time-independent, axially symmetric (Ux(x,y) = Ux(x,y), Uy(x,y) = Uy(x,y)) velocity pattern. Eq. (1) summarizes the assumptions that filament motion is similar on both the left and right sides of the embryo, and the same pattern is repeated at gradually more caudal locations as the primitive streak regresses. In other words, if an observer’s motion is coupled to Hensen’s node, then that observer would witness a stationary velocity pattern: Similar velocities would be obtained at the same locations (relative to the node), regardless of time or developmental stage. This time-independent behavior was statistically verified by calculating C(s), the temporal correlation function of the velocity field. As described in the Supplementary Material, C(s) measures the average similarity of velocity snapshot pairs, in which the elements are separated by a time interval of length s. We found that the velocity field V becomes uncorrelated (C(s) = 0) within 4 h ( P > 0.2), while the same velocity data, when viewed from a reference system co-moving with the node, remained significantly ( P < 0.01) correlated (C(s) > 0) during the entire observational period. To extract the velocity pattern U, velocity snapshots were shifted so that the locations of Hensen’s node and the somitogenesis front were held stationary and subsequently averaged over time as U a ðx; yÞ ¼ hV a ðx; y þ Ya ðtÞ; tÞit : These patterns, characterizing the fibrillin 2 velocity pattern in individual embryos, are similar (Fig. 5A) and form the basis of our statistical sample-to-sample comparison. The velocity pattern, which is also averaged over the individual embryos (Fig. 5B), reveals tendencies more clearly than

trajectories do. The prevailing motion is caudally oriented around the node. This directionality changes to a medial orientation at lateral positions, while at the extra-embryonic boundary displacements are directed cranially—resulting in a vortex-like velocity field. The medial component of the velocity field thus substantially contributes to the accumulation of fibrillin-containing ECM between the node and the somitogenesis front. Within the same area, the gradually increasing cranial velocity component reflects an anterior – posterior axial strain of the tissue. The statistical errors of the data presented in Fig. 5B can be directly estimated from the scatter of velocities characterizing individual embryos (Fig. 5A). While the sample size and thus the error depends on the location, the estimated average statistical error is F20 Am/h for each velocity component.

Discussion In vivo ECM assembly The biological function of the ECM—its assembly, turnover, and remodeling—has been the focus of numerous studies (Dzamba et al., 2001; Hay, 1982; Ohashi et al., 1999; Schwarzbauer and Sechler, 1999). Recently a threedimensional structure of the fibrillins was proposed to explain its elastic properties in molecular terms (Baldock et al., 2001). However, very little is known about the dynamics of these molecules, particularly regarding the mechanisms by which the ECM undergoes large-scale (tissue level) remodeling. Yet, the distribution of the ECM can be as crucial as its molecular composition. ECM organization can determine tissue mechanical properties (Barocas and Tranquillo, 1997; Olsen et al., 1999), as well as serve as a guidance system for cells. For example, some cell types, such as fibroblasts (Dickinson et al., 1994; Stoplak and Harris, 1982), endothelial cells (Vernon et al., 1995), and neurons (Dubey et al., 2001), are known to preferentially follow oriented fibers. In addition to establishing a detailed time-dependent record of fibrillin localization, we also determined the dynamics of ECM redistribution. To achieve this, we traced filaments, that is,

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certain identifiable segments within the immunolabeled network. Although the ultrastructure of the early ECM is not yet determined at early developmental stages, it is highly likely that filaments are composed of multiple ECM molecules (Visconti et al., 2003). Because filaments retain their shape between consecutive frames, position changes were interpreted as physical displacements of intact objects rather than as a process involving filament disassembly followed by reassembly and antibody reattachment. If

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disassembly – reassembly were occurring, that would be manifested as the visible, frequent disappearance of entire filaments; however, this was never observed. Moreover, the finding that displacements of adjacent filaments are similar suggests that filaments are embedded in a viscoelastic continuum (most likely including attached cells), rather than existing as free-moving or quickly disconnecting and reconnecting polymers. Fibrillin 2 in somitogenesis Our data show that the fibrillin-containing ECM is substantially reorganized during development. Among the observed processes, the formation of fibrillin bundles in the somitic and pre-somitic mesoderm is of special interest. These fibers delineate existing and future somites and may contribute to the integrity of the somitic array by providing a suitable anchoring mechanism. Since somites are surrounded by an acellular ECM, such a scaffold is likely to be required before the physical separation of the cells from the pre-somatic mesoderm and at the intersomitic clefts. The spatial and temporal pattern of fibrillin 2 localization is thus fully compatible with a presumptive scaffolding role. The condensation of fibrillin-containing bundles in the medial segmental plate mesoderm is also an unsuspected process of pattern formation: The demarcation between the axial cables and the less organized lateral meshwork is fuzzy and does not coincide with preexisting anatomical structures. Moreover, a substantial portion of fiber components are not secreted locally, but collected from a large area (Fig. 2). This is in contrast, for example, with the ECM ensheathment of the notochord, where the creation of the notochord boundary clearly precedes the gradual ac-

Fig. 4. Trajectories of fibrillin filaments and reference points. Eighty filaments were selected randomly in the embryo shown in Fig. 1, and their respective trajectories plotted with colored lines. Each color represents positions within 2-h-long time intervals: red denotes the original position of each filament (at frame 1), followed by orange, green, cyan, and blue positions, respectively. Concurrent trajectories of the spatial reference system, the intersomitic clefts, are plotted with cyan lines (e.g., white circle). The DIC image, upon which the trajectory lines are superimposed, shows the embryo at an intermediate time point, taken 4 h after the start of the observation. The motion of the filaments within the lateral plate mesoderm (bracket) is strongly medialward. In contrast, filament trajectories near Hensen’s node show a caudalward directional bias (asterisk). Between these two regions, the filament trajectories gradually change from medialward to caudalward. This is in distinct contrast to the displacements of the intersomitic clefts or reference points that are typically very small— an indication that the intersomitic clefts are well suited to serve as an anatomical reference system. The yellow box encloses an area in which early (red) fibrillin trajectory segments are perpendicular to later (blue) trajectory segments. Thus, the prevalent directionality of a fibrillin 2 filament can be highly time-dependent at one region (yellow box), while very little change in directionality is exhibited at positions caudal to Hensen’s node (asterisk). These data demonstrate that fibrillin filaments are not moving in lockstep; rather, the trajectories appear as a part of a swirllike pattern (see Fig. 5).

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Fig. 5. Experimentally derived velocity patterns of fibrillin 2. Locally averaged velocities of Cy3-labeled fibrillin filaments are plotted at grid-points of a square mesh. For each grid-point, velocities are represented by scale bars giving the local average displacement in 1 h. The vertical and horizontal yellow lines show the positions of the embryonic axis and the somitogenesis front, respectively. Small and large red circles indicate the somitic array and Hensen’s node. The velocity patterns characterizing five individual embryos are superimposed in panel a; thus, at each grid-point, the different scale bars represent velocity data obtained in different embryos. The green box delineates an area, where most grid-points contain data from at least two embryos. For each grid-point, the average of the various velocities was calculated and plotted in panel b. The blue arrow depicts the prevailing filament motion lateral to the node.

cumulation of the fibrillin-containing meshwork (see Figs. 1 and 2; supplementary data movies 1 and 2). Thus, in the latter case, tissue boundaries can direct a local ECM

accumulation. In accord with this assumption, we found no evidence of fibrillin 2-containing filaments being transported to the notochord from distant sites.

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Cranial – caudal developmental gradient is reflected in the traveling velocity pattern The similarities between a caudal region of an older embryo and a corresponding cranial part of a younger embryo underlies the hypothesis that the velocity field reiterates a basic pattern at progressively more caudal locations—following the regression of the primitive streak. Since the distance between the newest somite and Hensen node is stable during these developmental stages, we chose to follow the somitogenesis front as a positional indicator that moves in concert with this cranial – caudal developmental gradient. Our observations are confined to the caudal region of the embryo, between HH stages 5 and 10. Later, or concomitant but more cranial events of ECM remodeling are further complicated by additional morphogenetic processes. An important example of these is vasculogenesis, since fibrillin fibers incorporate into, and appear to delineate the devel-

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oping vascular network (not shown). Moreover, as Hensen’s node approaches the caudal boundary of the area opaca, its linear motion slowly diminishes (Spratt, 1946, 1947). Therefore, we do not expect our scheme of an evenly shifting velocity pattern to hold for later stages (above HH 13). While the scope of the present investigation is limited to stages and locations in which early axis patterning is occurring, our method of visualization, tracking, and quantifying the motion of ECM filaments should be useful in studies of other, perhaps more complex, morphogenic processes (as evidenced by the data obtained from regions of AIP regression and bilateral heart tube fusion). Forces that shape the embryo Several investigators have drawn attention to the importance of physical forces in the shaping of an embryo (Oster et al., 1983; Trinkaus, 1984). This conceptual framework

Fig. 6. The conjectured interdependence of ECM and mesodermal cell motion. In a computer simulation, cells were assumed to engage in autonomous motility relative to the surrounding ECM (including fibrillin 2), while the ECM was also assumed to move—according to the experimentally obtained velocity field summarized in Eq. (1). The simulation was two-dimensional; only anterior – posterior and lateral – medial movements were considered. Six distinct cell populations were followed—three groups on each side of the axis (panel a). The drawing in panel b, modified from Hay (1982), indicates the postulated path (yellow arrow) of the involuting mesodermal cells at a position along the anterior – posterior axis, denoted by the blue line in panel a. In particular, cells were assumed to move laterally with a constant speed of 25 Am/h (yellow arrow). The empirically determined ECM motion is indicated by red arrows. As a result of these assumptions and empirical data, the computer simulation yielded a patterned array of cells dispersed along the anterior – posterior axis, depicted in panel as black dots. Thus, the combination of lateral cell locomotion and a caudalward passive motion can result in a pattern that bears a striking resemblance to cell fates described in Fig. 12 of Schoenwolf et al. (1992) and Fig. 7 of Psychoyos and Stern (1996). The red circles in panel a denote the somites and Hensen’s node, respectively, while the red lines indicate the notochordal axis.

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asserts that (1) as tissues are physical objects, their deformation inevitably involves mechanical forces. (2) The relevant forces are exerted internally within the embryo by cells. Morphogenetic movements are thus determined by the interplay between the spatio-temporal pattern of exerted forces and the mechanical properties of the responding tissues. Recently, viscoelastic parameters of embryonic cell aggregates were established (Forgacs et al., 1998) and gastrulation movements of sea urchin embryos have been analyzed using viscoelastic models (Davidson et al., 1995, 1999; Drasdo and Forgacs, 2000). These latter studies also shed light on the crucial role of the ECM in the mechanics of gastrulation. Mechanical tension fields within the avian embryo are also thought to play a significant role during avian gastrulation and axial morphogenesis. It has been postulated that at very early stages, the tension generated by epiboly, the expansion of the embryonic plate (epiblast) on the vitelline membrane, is partially responsible for the elongation of the primitive streak (reviewed in Stern and Canning, 1988; Trinkaus, 1984). Although no stress or strain data are available yet for avian embryo development, such experiments were carried out in Xenopus embryos (Moore et al., 1995). Unlike cells, ECM components cannot move themselves; thus, they are ideal ‘‘passive tracers’’ of tissue movements. Although cells are known to reorganize matrix molecules on their surface, displacements exceeding 100 Am clearly cannot be generated in that manner. Thus, we propose that (1) the observed displacements reflect—at least partially— large-scale tissue movements driven by mechanical forces, (2) an existing ECM structure can be substantially reorganized by a mechanical tension-driven meshwork-to-cable transition, and (3) formation of these structures is strongly dependent not only on local cellular activity, but also on the global developmental movements of the embryo. Based on the nonspecificity of mechanically transduced effects, we expect that fibrillin 2 is not unique and that other ECM constituents exhibit similar behaviors. Further experiments are underway to test these conjectures directly. Although our results conclude with the determination of a velocity field that characterizes the nodal region of an avian embryo, we expect that these data will eventually be used to estimate the distribution and origin of the forces acting on the embryonic plate. A similar approach was recently applied to determine the traction forces of cells embedded in collagen gels (Vanni et al., 2001). In our case, however, the viscoelastic properties of the tissue are more complex than those of a collagen gel undergoing small deformations, and the full determination of the stress field requires knowledge about all three components of the velocities (Helmke and Davies, 2002).

local environment, or large-scale tissue movements can passively transport them. For example, heart looping clearly involves tissue deformations that move cells—relative to the somites, but also relative to each other (Voronov and Taber, 2002). These active and passive modes of relocation are not mutually exclusive; in fact, both are expected to shape the embryo. Our data inevitably raise the question about the possible relation between the studied ECM motion and the reported features of mesodermal cell migration (Psychoyos and Stern, 1996; Schoenwolf et al., 1992). Based on the idea of superimposing the ECM and cell movements, we employed a computer simulation to model the lateral spreading of mesodermal cells (Fig. 6). In this hypothetical model, (a) the traced cell population is ingressing continuously along the primitive streak, (b) the ingression is followed by a steady, laterally directed active movement relative to the surrounding ECM, (c) the ECM moves according to the experimentally obtained pattern, summarized in Eq. (1). In Fig. 6, a subset of the cells is visualized as being both carried along with and moving relative to the ECM environment. Although we assumed a uniform locomotory activity, the combination of the lateral migration of the cells together with the experimentally determined nonuniform motion of the ECM results in a pattern recapitulating the experimental findings of Schoenwolf et al. (1992) and Psychoyos and Stern (1996). The validity of this picture needs to be established by comparing timeresolved experimental data on concurrent cell and ECM movements. Systems approach It has been evident for decades that morphogenesis involves an interplay of physical forces, cell behavior, and genetic regulation. Unfortunately, models encompassing pertinent developmental complexities are difficult to develop due to the paucity of nongenetic quantitative experimental data. Recent advances in optical microscopy, image processing, and computing power have extended the scope of timelapse investigations into the quantitative realm. One resulting advantage is the ability to characterize developmental processes with higher precision than earlier fate mapping methods. These precise quantitative data can be used to construct and test numerical models. Such models, used as part of a ‘‘systems’’ or integrative approach (Hove et al., 2003; Noble, 2002), will permit a new level of understanding of cell, tissue, and ECM dynamics during morphogenesis.

Acknowledgments Gastrulating cells are conjectured to move relative to the surrounding ECM Embryonic cells can be rearranged by at least two different means. Cells can actively migrate relative to their

We are grateful to Adrienn Gabor, Erica Perryn, and Paul Rupp for their valuable help in this project. We thank Tom Skalak (UVa) for critical comments on the manuscript. This work was supported by awards of NSF (NATO-NSF DGE-

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0075179, to A.C.), the Institutional Biomedical Research Training Program of KUMC (to A.C.), the Hungarian Research Fund (OTKA T034995, to A.C.), the Heartland Affiliate of the American Heart Association (0160369Z, to B.J.R.), the Mathers Charitable Foundation (to C.D.L.), and NIH (5P01 HL58213-08, to C.D.L.; 1R01 HL73700 to B.J.R.). The JB3 antibody was obtained from the developmental studies Hybridoma Bank at the Univ. of Iowa.

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