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First published online January 23, 2009
doi: 10.1242/10.1242/dev.026211
Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA 15260, USA.
* Author for correspondence (e-mail: ldavidson{at}engr.pitt.edu)
Accepted 1 December 2008
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: Gastrulation, Convergent extension, Axial elongation, Mechanics, Young's modulus, Viscoelasticity, Biomechanics, Extracellular matrix, Microfilaments, Xenopus
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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).
Additionally, anatomical structures within the embryo are themselves composed
of physical materials that exhibit molecular deformation, but their
contribution to the anatomical structure depends on how they are assembled
into the structure. To identify the relative contribution of these two aspects
of mechanical resistance requires the ability to measure tissue elasticity
experimentally after structures in the embryo or the molecular interactions in
materials the make up the embryo have been altered
(Koehl, 1990).
The notochord, a defining anatomical feature of chordate embryos, dominates
early morphogenesis as both a source of molecular signals and later as a
mechanical structure. The notochord first acts as an organizing
center-releasing growth factors to induce midline structures such as the
neural notoplate (later the floorplate) and hypochord
(De Robertis and Kuroda, 2004;
Stemple, 2005). The second
function of the notochord later in development is as a mechanical structure,
providing a stiff `backbone' for muscle attachment in the swimming tadpole
(Hoff and Wassersug, 2000;
Wassersug, 1989). The earliest
mechanical role of the notochord during morphogenesis occurs as notochord
cells change shape and undergo mediolateral intercalation during convergent
extension (Shih and Keller,
1992), and then extends faster than somitic mesoderm as the dorsal
axis elongates (Keller et al.,
1989). Experimental embryology and computer simulations suggest
that elongation of the notochord and the attached notoplate shape the
vertebrate neural plate (Jacobson and
Gordon, 1976). Other dorsal tissues, including the paraxial
somitic mesoderm (Wilson et al.,
1989) and the neural plate
(Elul and Keller, 2000;
Ezin et al., 2006), also
contribute to dorsal extension.
In contrast to the clear-cut role for the notochord in patterning, there is
qualitative support for and against a mechanical role of the notochord in
dorsal elongation and neural tube closure. Malacinski and co-workers
(Malacinski and Youn, 1981;
Youn and Malacinski, 1981)
demonstrated that Xenopus embryos without notochords could converge,
extend and elongate nearly as well as embryos with notochords. Evidence
supporting a mechanical role for the notochord in dorsal extension is found in
both amphibian embryos (Kitchin,
1949; Lehman and Ris,
1938) and zebrafish mutants
(Talbot et al., 1995), in
which notochord tissues have been ablated, which produces embryos with
shortened axes; however, these developmental defects may be due to losses of
other dorsal tissues and failure to establish dorsal tissue architecture. The
generation of force by the notochordal region is required to produce the
elongation in the amphibian neural plate
(Jacobson and Gordon, 1976).
Failure of dorsal tissue elongation is thought to be one of the principle
causes of neural tube defects in frog
(Wallingford and Harland,
2002), mouse (Copp et al.,
2003; Ybot-Gonzalez et al.,
2007) and humans (Kibar et
al., 2007). These studies suggest an important mechanical role for
dorsal mesodermal tissues, as they apply forces to the overlying neural plate,
extend the dorsal axis and the close the neural tube without failing or
buckling.
The extracellular matrix and the cytoskeleton are the most likely
contributors to the molecular basis of tissue mechanics relevant to
embryogenesis. ECM has been identified as a major mechanical contributor to
tissue stiffness in echinoderm embryos
(Davidson et al., 1999), adult
tissues (Levental et al.,
2007; Vincent,
1990) and tumors (Paszek et
al., 2005). Collagen type II fibers form some of the stiffest
fibers identified in animals (Levental et
al., 2007) and are a strong candidate for stiffening the notochord
in later dorsal tissues (Adams et al.,
1990). Likewise, fibrillin could also contribute to dorsal tissue
stiffness analogous to its contribute to tissue stiffness in the lung
(Waters et al., 2002), where
fibrillin and elastin undergo cyclic extension and contraction and prevent
viscoelastic creep (Findley et al.,
1989) that would otherwise lead to rapidly failing `stretched-out'
hearts and lungs. Laminin in its many heteromeric forms may also contribute to
dorsal tissue stiffness (Georges et al.,
2006). Each of these candidate ECM components, fibronectin
(Lee et al., 1984), fibrillin
(Skoglund et al., 2006) and
laminin (Wedlich et al., 1989)
are assembled into fibrils in frog dorsal tissues through these stages,
suggesting that ECM contributes to tissue stiffness from mid- to
late-gastrulation and beyond during neurulation.
Tissue mechanics crucially depend on factors such as Rho-GTPase that
regulate the actomyosin cytoskeleton to direct cell migration, cell mechanics
and assembly of the extracellular matrix
(Paszek et al., 2005).
Numerous studies have implicated the actin cytoskeleton as a major contributor
to the mechanical properties of single cells (for a review, see
Luby-Phelps, 2000). Additional
crosstalk may couple cell-cell adhesion to actomyosin contractility in frog
embryos (Kofron et al., 2002;
Tao et al., 2007), where
activated cadherin in cell-cell adhesions has been shown to control cortical
F-actin assembly. In contrast to a growing consensus on the molecular factors
that modulate dynamic F-actin assembly
(Pollard and Borisy, 2003) and
the mechanics of crosslinked F-actin gels in purified systems
(Gardel et al., 2006;
Janmey and Weitz, 2004),
little is understood about the contribution of the actin cytoskeleton to
mechanical properties of tissues.
In order to investigate the mechanical properties of embryonic tissues and
isolate the relative contribution of anatomical structures and molecular
composition to those properties, we use the frog embryo to microsurgically
isolate dorsal tissues, manipulating both tissue architecture and molecular
composition, and then measuring the time-varying stiffness (i.e. Young's
modulus or mechanical resistance) (Janmey
et al., 2007) along the anterior-posterior direction that results
from these manipulations. We use standard engineering measures of material
properties so that our results can be compared directly with theoretical and
empirical studies conducted on a variety of biomaterials, structures and
organisms (Koehl, 1990). In
agreement with a previous study of Xenopus embryonic tissues
(Moore et al., 1995), we find
that dorsal isolates also behave like a conventional viscoelastic material
capable of elastically supporting applied forces. We find dramatic increases
in tissue stiffness at later stages and equally large differences between the
stiffness of germ layers that form dorsal tissues. Using antisense morpholinos
to block fibronectin fibril assembly and acute-acting actomyosin disrupting
drugs, we find that embryonic tissue stiffness at these stages originates in
the actin cortex rather than in the fibronectin extracellular matrix. We
propose that dorsal tissues form an integrated composite structure that is
dependent on actomyosin contraction that generates force and resists
catastrophic failure to drive dorsal axis elongation and neurulation.
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MATERIALS AND METHODS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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) and
cultured to stage 16 (Nieuwkoop and Faber,
1967), at which time dorsal axial and paraxial tissues (dorsal
isolate) are microsurgically removed from the embryo
(Fig. 1A). Dorsal isolates can
be harvested from embryos from mid-gastrulation (stage 11.5) to neural tube
stages (stage 22) and beyond. The dorsal isolate can be reproducibly
subdivided even further along the mediolateral or dorsal ventral axis.
Histology, immunostaining, and confocal microscopy
To visualize ECM and pMLC, tissue explants and whole embryos were fixed in
either 3.7% formaldehyde or 3% TCA in PBS (see
Davidson et al., 2004).
Fibronectin fibrils were localized with mAb 4H2 directed against
Xenopus fibronectin (1:500), fibrillin-2 fibrils with mAb JB3 (1:500;
Developmental Studies Hybridoma Bank), laminin 1 with a rabbit pAb (1:100;
cat# L-9393, Sigma) and activated myosin light chain (pMLC) with a rabbit pAb
(1:200; ser19 phosphorylated; Cell Signaling Technology) and visualized with a
rhodamine-conjugated goat anti-mouse or anti-rabbit IgG (Jackson
ImmunoResearch Laboratory). To visualize F-actin in embryos, we fixed embryos
using two media: either a modified Dent's fixative [replacing methanol with
isopropanol (Dent et al.,
1989; Munro and Odell,
2002)] or 4% paraformaldehyde with 0.25% glutaraldehyde
(Tao et al., 2007) in PBS.
F-actin was visualized by incubating fixed tissues for 3 hours with bodipy-FL
phallacidin (5U/ml PBST; Invitrogen). Following immunofluorescence or
phallacidin staining whole embryos were bisected or cut into en face
fragments, dehydrated in methanol or isopropanol (to preserve phallacidin
binding to F-actin) and cleared in Murray's clear
(Davidson and Wallingford,
2005). Single optical sections and z-series of explants
and whole embryo fragments were collected with a confocal laser scan head
(SP5, Leica Microsystems) mounted on an inverted compound microscope (DMI6000,
Leica Microsystems) using image acquisition software (LASAF, Leica
Microsystems). Maximum projection and reslicing of z-series stacks,
and collection of intensity profiles were carried out with ImageJ (v. 1.38,
Wayne Rasband, NIH).
Stress-relaxation and the nanoNewton force measurement device
The nanoNewton force measurement device (nNFMD) has been described
previously (Davidson and Keller,
2007). Briefly, the device for measuring nanoNewton to microNewton
scale forces was based on the earlier `Histowiggler' design
(Moore et al., 1995). Prior to
the start of an experimental run, the anterior or posterior face of a tissue
explant is brought into contact with a sensitive force probe. An experimental
run begins as the explant is moved into contact with the force probe and
compressed by 15 to 20% of its original length. The explant is held in place
for 180 seconds as a record of resistive force is collected. The experimental
run concludes as the explant is moved away from the force probe and placed in
fixative [MEMFA (Sive et al.,
2000)]. Stiffness (i.e. Young's modulus) is calculated from the
resistive force measured during the stress-relaxation test, the
cross-sectional area of the explant measured from the fixed sample and the
compressive strain observed from a time-lapse sequence of the
stress-relaxation test.
Determining the viscoelastic properties of embryonic dorsal tissues from an unconstrained uniaxial compression test
Dorsal tissues isolated from the late gastrula stage embryo exhibit
viscoelastic behavior like most `soft' biological tissues
(Fig. 1D)
(Wainwright et al., 1976).
This tissue, referred to as `dorsal isolate', contains all three germ layers
in their proper laminar context from the spinal cord/hindbrain boundary to
approximately 200 µm from the blastopore. The transverse cross-section of
the dorsal isolate is consistent along its anterior-posterior length. Germ
layers are separated from each other by tissue interfaces that delineate axial
notochord from surrounding tissues, and paraxial pre-somitic mesoderm from
neural ectoderm and endoderm (Fig.
1B).
The first step in quantifying the viscous and elastic properties of these
tissues is to determine the time (t)-dependent Young's modulus, E(t).
Throughout this study, we chose to compare the Young's modulus at the 180
second time-point in our compression test as E(180) represents the static
mechanical properties of the embryo in vivo. We refer to E(180) throughout
this study as tissue `stiffness', as this value has been obtained from an
unconfined uniaxial compression test. The Young's modulus at earlier
time-points reflects the changing viscous response of the tissue to a suddenly
applied strain, as well as variable loading conditions such as stiction and
compliance of our force transducer. The stiffness immediately after strain is
applied, E(0), after 90 seconds, E(90), and after 180 seconds, E(180), along
with standard deviations are provided in supplementary tables for each tissue
measured [see Tables S1-S11 in the supplementary material; in addition we
provide the parameters k1 (Einf), k2 (Esp), and
(viscosity) representing the Standard Linear Solid (SLS) Model
(Findley et al., 1989) fitted
using nonlinear regression techniques (NLREG ver. 3.2; Brentwood TN) for each
tissue measured]. Statistical tests of significance of E(180) values were
carried out pair-wise for sets of explants from each clutch with the
non-parametric Mann-Whitney U-test
(Sokal and Rohlf, 1994) using
commercial software (SPSS v. 16; Chicago, IL). In the course of measuring the
E(t) of embryonic tissues, we found several complications to fitting the data
with the SLS model. First, the initial elastic response [E(t) for t<10
seconds] is significantly higher than the parameters describing the initial
elastic response (k1+k2) from a best-fit SLS model (see
Fig. S2B in the supplementary material). This high value may reflect static
friction (i.e. stiction) of the tissue explant with the holder, or may
indicate high frequency mechanical compliance of our force transducer. Second,
we occasionally observe spontaneous mechanical contractions during compression
tests (data not shown). These contractions appear as `bumps' 30 seconds to 1
minute in length in which the E(t) rises transiently and then falls back.
These contractions are similar to spontaneous contractions observed with other
mechanical test devices in our laboratory
(von Dassow and Davidson,
2009). Applying nonlinear regression to fit these traces results
in low-quality SLS parameters that consistently underestimate the observed
Young's modulus. In order to enable comparison of our data with those
collected by other laboratories and for comparison with other materials, we
include the nonlinear regression `best-fit' SLS model parameters of
k1, k2, and for all tests in Table S1-S11 (see
supplementary material). We chose to use the measured E(180) rather than the
SLS model-calculated k1, because, as discussed above, fitted SLS
model parameters introduce errors not found in measured data.
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):
Etotal x Atotal=
Ei x
Ai. The special case for the dorsal isolate (DI) is written:
EDI x ADI=Enoto x
Anoto + Emedial mesoderm x Amedial
mesoderm + Elateral mesoderm x Alateral
mesoderm + Eneural x Aneural +
Eendo x Aendo. This equation relates the
dependence of the stiffness of the dorsal isolate (DI) (see Fig. S3A in the
supplementary material) on its component parts: notochord (noto),
paraxial-medial mesoderm (medial mesoderm), paraxial-lateral mesoderm (lateral
mesoderm), neural plate (neural) and endoderm (endo) (see Fig. S3B in the
supplementary material). Measured stiffness and cross-sectional areas from
sets of explants can be written as sets of algebraic equations where we can
`solve' for the mechanical contribution of the tissue that has been
excised.
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Dorsal tissues are viscoelastic but behave elastically during elongation
As biological tissues can exhibit a wide range of mechanical behaviors from
rigid elastic to fluid-like and can actively change shape unpredictably in
response to applied forces, we carried out a series of mechanical tests to
determine whether standard techniques could be used to study dorsal isolates.
First, we assessed the amount of strain present in the embryo as large amounts
of pre-strain can significantly alter the interpretation of mechanical tests
of stiffness (Zamir and Taber,
2004b). To measure the degree of pre-strain in dorsal isolates, we
collected images of the epithelial surface of whole embryos before
microsurgery and compared those images with ones collected immediately after
microsurgery. We measured strain in five embryo-explant pairs and have found
the surface layer in dorsal isolates contract around 10% isometrically (see
Fig. S1A-D in the supplementary material). Thus, some small degree of
pre-strain is present in the embryo but we do not see large amounts of elastic
recoil as observed when biological tissues within hydrostat-like structures,
such as the embryonic heart, are removed from their micro-environment
(Zamir and Taber, 2004a). We
then determined the response of dorsal isolates to applied force, the
linearity of their elastic response to that force, whether the response
changes with repeated application of force and whether tissue became stiffer
in response to higher applied forces. Single dorsal isolates at stage 16 were
placed into the nanoNewton force measurement device (nNFMD)
(Fig. 1C) and subjected to a
nine-minute uniaxial compression test (see Fig. S1E in the supplementary
material) demonstrate that the elastic resistance of dorsal isolates reaches
steady-state by 180 seconds. Repeated series of five, 3-minute
compression/1-minute relaxation tests (see Fig. S1F in the supplementary
material) or five 1-minute steps of incrementally increasing strain (see Fig.
S1G in the supplementary material) on the same dorsal isolate reveal that
isolates do stiffen slightly with repeated cycles or increasing strain but
remain viscoelastic.
Taken together, these mechanical tests demonstrate that dorsal tissues behave like conventional viscoelastic materials with an immediate viscous response to applied strain. Dorsal isolates are able to maintain a long term residual stiffness and their residual Young's modulus (hereafter referred to as `stiffness') can be reliably and reproducibly measured at the conclusion of a single 180-second unconfined uniaxial compression test. The goal of the remainder of this paper is to characterize stage- and tissue-specific variation in these tissues, and to test the role of the ECM and actomyosin cytoskeleton as contributing factors.
Developmental changes in axial stiffness
Since a previous study (Moore et al.,
1995) found increasing stiffness in dorsal involuting marginal
zone tissues early in gastrulation (between stages 10+ and 11.5) coincident
with the onset of convergent extension, it is important to know whether
tissues continue to stiffen during the subsequent rapid phase of narrowing and
elongation that precedes neurulation. Dorsal isolates excised from whole
embryos converge and extend at the same rate as the identical tissues in whole
embryos and preserve the relationship between endoderm, mesoderm and neural
ectoderm, as seen in whole embryos (Fig.
1B). To ensure that the same tissues were tested at both early and
late stages, we prepared dorsal isolates at the end of gastrulation (stage 13)
and aged them until co-cultured control embryos reached the defined stage
(with the exception of the mid-gastrula stage isolate). Single explants were
placed into the nanoNewton force measurement device
(Fig. 1C) and subjected to a
180-second stress-relaxation protocol applied along the anteroposterior axis
of the isolate (Fig. 1D). In
preliminary studies, we found clutch-to-clutch variation in the mechanical
properties of embryonic frog tissues (von
Dassow and Davidson, 2009) (data not shown). To compensate for
this variance, we typically measured and compared tissue stiffness changes
with developmental stage within the same clutch. Previous measurements found a
threefold increase in axial mesodermal tissue stiffness from 3 to 10 Pa from
early gastrula to mid-gastrula stages
(Moore et al., 1995). With the
updated force measurement device, we found dorsal tissues continued to
increase stiffness along the anteroposterior axis from 13 Pa to 85 Pa
(Fig. 1E; representative data
shown), an increase of more than sixfold from late gastrula (stage 13) to
early neural tube stage (stage 22).
Midline region, but not notochord, contains stiffest tissue
As the notochord is a central structure during gastrulation and
neurulation, and undergoes substantial changes in architecture during this
period, we decided to test the contribution of the notochord to the stiffness
at early neurula stages (stage 16). To accomplish this, we microsurgically
made dorsal isolates with two-notochords and zero-notochord
(Fig. 2A). To make a dorsal
isolate with two notochords, we recombined a left-half explant with notochord
with a right-half explant with notochord. The medial edges of the left and
right explants were then held in apposition with a glass coverslip fragment
until they healed together, typically 30 to 60 minutes. Dorsal isolates
without notochord were assembled from the two remaining lateral fragments. To
control for the possibility that microsurgery altered the stiffness of dorsal
isolates we also made `sham-operated' control dorsal isolates that were split
axially and then re-combined. After healing, sham-operated controls,
two-notochord and zero-notochord isolates retain the capacity for elongation
and achieve rates similar to intact dorsal isolates
(Fig. 2B). To confirm that the
architecture of these explants contained properly positioned dorsal tissues,
we carried out immunofluorescence to reveal fibrillar fibronectin and
collected stacks of confocal sections of stained explants
(Fig. 2C).
Using the nNFMD, we found two-notochord isolates had a twofold greater
stiffness over zero-notochord dorsal isolates
(Fig. 2D). We can compare the
stiffness of these differently sized explants as stiffness is independent of
the shape and size of the tissue (Koehl,
1990). Close inspection of the tissue architecture of explants
with two notochords revealed that the two notochords were separated from each
other by a piece of notochord-adjacent paraxial mesoderm
(Fig. 2C, asterisk). To rule
out the effect of this small piece of notochord-adjacent paraxial mesoderm, we
again prepared two-notochord and zero-notochord dorsal isolates cutting closer
to the boundary between the notochord and somite
(Fig. 2A'). Staining of
these explants for fibronectin fibrils confirmed that we had completely
removed the medial notochord-adjacent paraxial mesoderm from the two notochord
isolates (Fig. 2C').
However, after removing the small amount of midline tissue, we found no
significant differences in the stiffness between isolates with only two
notochords and those without notochords
(Fig. 2D').
In order to rule out the effects of microsurgical recombination, we
compared the stiffness of dorsal isolates cut from dorsalized embryos. Our
strategy was to alter the early patterning of embryos to generate a larger
notochordal field and then use these embryos to microsurgically excise dorsal
isolates. We chose LiCl to dorsalize early embryos and measured the
consistency and degree of dorsalization with the Dorso-Anterior Index [DAI
(Stewart and Gerhart, 1990)].
By regulating the timing and limiting the dosage of LiCl to 0.3 M
(Fig. 2E), we could
consistently generate embryos with a DAI between 7 and 8
(Fig. 2F). DAI 7/8 embryos have
a single deep archenteron containing more than double the normal amount of
notochord, as judged both by chordin expression in dorsal isolates
(Fig. 2G), and fibronectin
fibril localization in transverse confocal sections
(Fig. 2H). Neither dorsalized
embryos nor dorsal isolates made from them undergo much elongation (data not
shown). Assured that dorsal isolates made from dorsalized embryos contained
large quantities of notochord, we measured their axial stiffness and found
that they did not differ significantly from control dorsal isolates made from
the same clutch (Fig. 2I)
(note: a single clutch from the first three clutches showed dorsalized tissue
was stiffer and prompted a more extensive test of two additional clutches,
none of which showed significant differences in stiffness). Thus, two
independent methods of producing excess notochord in dorsal isolates failed to
show any increase in stiffness. This surprising result prompted us to evaluate
paraxial somitic mesoderm as a potential source of the stiffness of the dorsal
isolate.
|
). As our first microsurgical attempts to make explants with
double notochords instead revealed a stiff material lodged between the two
notochords, we suspected the medial fragments from paraxial mesoderm made a
larger contribution to overall stiffness. To test this hypothesis, we again
prepared zero-notochord explants from stage 16 embryos (here referred to as
LMML for Lateral-Medial-Medial-Lateral isolates) consisting of two complete
arrays of paraxial mesoderm and compared them with isolates that contain only
lateral mesoderm (LL, Lateral-Lateral isolates)
(Fig. 3A). Both of these
tissues are similar in appearance when stained for fibronectin fibrils
(Fig. 3B); however, isolates
containing medial somitic mesoderm are clearly identified by their expression
of the prospective muscle marker XmyoD
(Hopwood et al., 1989) (left
and middle columns in Fig. 3C),
which is absent in LL explants (right-most column in
Fig. 3C). Furthermore, LL
explants retain the elongation capacity of intact dorsal isolates
(Fig. 3D). Confident of our
ability to microsurgically separate lateral from medial paraxial tissues we
made LL and medial-medial paraxial (MM) explants combining both left- and
right-hand tissues from the same embryo
(Fig. 3A). Measuring the axial
stiffness of these tissues we found LL explants were significantly less stiff
than MM explants (Fig. 3E). The
higher stiffness of isolates containing medial notochord-adjacent paraxial
mesoderm supports the hypothesis that it is not the notochord but rather
notochord-adjacent paraxial tissues that contribute most to the axial
stiffness of dorsal isolates at mid neural plate stages.
The MM explant consists of neural and endodermal tissues in addition to
medial paraxial mesoderm; each of these tissues could contribute to the
increased stiffness of the MM explant. To determine the contribution of these
other tissues to axial stiffness, we made explants from intact dorsal isolates
from which either endoderm or neural ectoderm had been removed
(Fig. 4A). Each of these
explants had been used in previous studies of segmentation
(Wilson et al., 1989) where
they were reported not to extend as well as intact dorsal isolates. We are
able to prepare mesoderm-only explants; however, these explants are too thin
to test in the nNFMD. Comparing the stiffness of no-neural dorsal isolates
with control explants, we found they were not significantly different
(Fig. 4B). By contrast, we
found no-endoderm isolates were significantly stiffer than control dorsal
isolates (Fig. 4C). This last
result indicates that the endoderm is the weakest tissue in the dorsal
isolate.
Resolving the stiffness of tissues that comprise the dorsal isolate
Stiffness measurements of reconstructed dorsal isolates reveal significant
differences between tissues but cannot directly quantify the stiffness of any
single tissue. By definition, the stress relaxation protocol reports a
representative bulk time-dependent stiffness that is independent of the
geometry of the dorsal isolate (Koehl,
1990; Vincent,
1990). Another limitation is due to sensitivity of the nNFMD;
smaller pieces of tissue do not produce sufficient compression-resistant
forces to determine their stiffness.
To determine the stiffness of each of the germ layers, we formulated a
simple architectural model of the dorsal isolate. Based on superposition
principles of composite structures
(Christensen, 1991), we
constructed a simple mechanical model of the dorsal isolate that allows an
estimate of the stiffness of the component tissues of the dorsal isolate (see
Materials and methods). In essence, each tissue contributes to the stiffness
of the whole tissue explant according to its individual stiffness and to its
contribution to the full transverse cross-sectional area of the tissue
explant. Each component tissue is analogous to a spring and the full explant
to a series of springs in parallel. Force applied to the anterior or posterior
ends of the explant is distributed to all the springs to bring about the same
degree of compression in each. In this way, we account for tissue mechanical
properties that cannot themselves be isolated from their surroundings or are
not compatible with the nNFMD. Qualitatively, we can consider the contribution
of the various tissues to the stiffness of recombinant explants: (1) when we
remove a relatively stiff tissue the resulting recombinant is less stiff; (2)
when we remove an easily deformed tissue the resulting recombinant is stiffer;
and (3) when we remove a tissue that is equal to the stiffness of the whole
isolate we see no difference in stiffness. Using the composite model and
comparing the stiffness of endoderm-free, neural-free, notochord-free,
two-notochord, LiCl-notochord and LL versus MM explants, we conclude that
there are three levels of tissue stiffness in the dorsal isolate. From this
analysis, we rank the relative stiffness of the components of the dorsal
isolate: (1) the endoderm is less stiff by an order of magnitude than the
intact dorsal isolate (with stiffness between 2 and 11 Pa; 6 to 22% stiffness
of the intact dorsal isolate); (2) the neural and notochord are equivalent to
the stiffness of the intact dorsal isolate (between 40 and 60 Pa), and the
paraxial mesoderm is stiffest (between 70 and 100 Pa; 140 to 170% of the
stiffness of the intact dorsal isolate) and nearly twofold stiffer than the
notochord or neural tissues. A more quantitative evaluation of specific tissue
stiffness will require development of more spatially sensitive methods either
by excising and testing smaller tissue explants, or by measuring spatial
variation within intact explants or embryos. With our measurements of
anatomical contributions to tissue stiffness, we turned to assessing the
molecular contribution to tissue stiffness.
|
) and
deposition of fibrillin (Skoglund et al.,
2006) around the paraxial mesoderm
(Fig. 5A). To reduce the
assembly of fibronectin ECM, we injected fertilized eggs with a mix of two
antisense morpholinos directed against the 5' start of the two
fibronectin pseudoalleles expressed at these stages (6 µM each; FNMO). At
these doses, FNMO has been previously shown to knock down fibronectin
synthesis and its subsequent assembly into fibrils in Xenopus embryos
(Davidson et al., 2006).
Dorsal isolates from injected embryos at stage 13 showed no difference in
stiffness from control uninjected embryos
(Fig. 5B, left-most bars);
furthermore, additional measurements of stiffness of stage 16 dorsal isolates
showed FNMO-injected explants increased in tissue stiffness, just as control
dorsal isolates (Fig. 5B,
right-most bars). Each tissue explant tested was stained for fibronectin,
demonstrating reduced assembly of fibronectin fibrils in FNMO-injected
explants (Fig. 5C, top panel).
Thus, dorsal isolate stiffness was unchanged after fibronectin fibrillogenesis
was severely blocked.
As it has been suggested that assembly of other ECM proteins might depend
on fibronectin fibril assembly (Sivakumar
et al., 2006), we investigated whether we could also assess the
contribution of fibrillin and laminin extracellular matrix to the stiffness of
the dorsal isolate by mid-gastrula stages. Staining FNMO injected dorsal
isolates revealed reduced fibrillin and laminin fibril assembly. Fibrillin
shows strong reduction in the assembly in FNMO injected dorsal isolates
(Fig. 5C, middle panel),
whereas laminin assembly is both reduced and disorganized
(Fig. 5C, bottom panel).
Quantitative analysis of staining across the somite-notochord boundary
demonstrates that, in addition to the 60% reduction of fibrillar fibronectin,
fibrillin and laminin are reduced by 22% and 29%, respectively, in
FNMO-injected dorsal isolates (Fig.
5C'). The collateral inhibition of fibrillin and laminin
assembly suggests that fibrillar ECM may not contribute to tissue stiffness
during mid-gastrulation and neurulation.
Reducing F-actin or myosin II contractility reduces dorsal tissue stiffness
To test the contribution of the actin-based cytoskeleton to tissue
stiffness, we prepared dorsal isolates and treated them with a panel of
acute-acting drugs. Previous studies have shown that latrunculin B (latB)
effectively de-polymerizes F-actin within whole embryos
(Fig. 6A)
(Benink and Bement, 2005;
Lee and Harland, 2007) and
Y27632, a Rho-kinase inhibitor, blocks myosin II activation
(Maekawa et al., 1999;
Narumiya et al., 2000). Both
latB (incubated 20 minutes) and Y27632 (incubated for 60 minutes) reduced
tissue stiffness by at least 50% in dose-dependent manners
(Fig. 6B,C, respectively).
Treatment with high doses of latB over long times causes tissue explants to
irreversibly dissociate. LatB, like another F-actin depolymerizing drug
(cytochalasin D), causes disassembly of the fibronectin matrix
(Davidson et al., 2008) (data
not shown). However, as reduced fibronectin knockdown does not alter tissue
stiffness, we propose that the reduction in stiffness after treatment of
dorsal isolates with 0.6 µM latB is due to reduced F-actin rather than to
disruption of fibronectin fibrils. Combinations of both latB and Y27632 show
that the effect of latB dominates and that reduced myosin II contractility
makes little additional contribution to tissue stiffness
(Fig. 6D). Two additional
compounds have been reported to stabilize F-actin [jasplakinolide
(Cramer, 1999)] or to increase
myosin II contractility [calyculin A (Yam
et al., 2007)]; however, neither jasplakinolide (up to 10 µM
for 60 minutes) nor calyculin A (40 nM for 20 minutes) produced significant
changes in tissue stiffness (see Table S8 in the supplementary material).
|
),
we were able to resolve stage specific and germ layer-specific changes in
actomyosin. Surprisingly, neither F-actin
(Fig. 7A,B) nor pMLC
(Fig. 7C) shows gross changes
in intensity between stages 13 and 16 (embryos were fixed and stained
together, and confocal sectioned with the identical settings). However, there
are considerable changes in the organization of pre-somitic mesoderm into a
dorsal and ventral leaflets. The interface between these two layers, known as
the pre-myocoel (Keller,
2000), shows increased levels of F-actin localization
(Fig. 7B,C, arrowheads) that
may indicate the assembly of a new load-bearing mechanical structure within
the pre-somitic mesoderm. In contrast to the lack of F-actin differences
between stages 13 and 16, the endoderm, the most compliant tissue in the
embryo, shows dramatically lower levels of F-actin. We suspect that this does
not simply reflect the larger size of endoderm cells (as large mesendoderm or
bottle cells show intense F-actin, data not shown) but instead reflects
repression of F-actin assembly within these cells. Thus, we may conclude that
some, but by no means all, of the spatial and temporal variation in stiffness
is due to absolute levels of F-actin or activation of myosin II. |
DISCUSSION |
|---|
|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
) to
determine the contribution of various regions of the dorsal isolate to its
stiffness. Through a process of microsurgical elimination, we discovered that
it is not the notochord but a notochord-adjacent domain of paraxial mesoderm
that is responsible for more than 60% of the mechanical stiffness of the
dorsal axis.
|
The classification of the dorsal isolate, or any material for that matter,
as viscoelastic, does not imply any particular mechanism is responsible for
its physical response to applied force. We have chosen to represent the
viscoelasticity by a spring and dashpot network for several reasons: (1) the
parameters from the fitted spring and dashpot network allow comparison of
embryonic tissues with synthetic materials and with biological tissues whose
resistive force increases with increasing applied strain; and (2) these
measured values can be used by theorists interested in simulating gastrulation
and neurulation. In contrast to the generalized springs and dashpots that make
up a viscoelastic model, it is not correct to think of the viscoelastic
response of a biological tissue as a `passive' response but instead as a
property of a living tissue. One could imagine a more complex response to
applied force in which the tissue mimics a viscoelastic response; however, we
think this interpretation is both unlikely and unnecessary. Viscoelastic
mimicry would require a complex mechano-sensory feedback network capable of
actively generating resistive forces to match applied forces. By contrast,
theoretical and experimental analyses of biological polymers predict
viscoelastic behaviors like those seen in frog embryonic explants
(Flory, 1953;
Vincent, 1990). Further
studies will be needed to test the predictions of polymer mechanics, to search
for mechano-sensory feedback, and to resolve the molecular, cellular and
architectural sources of tissue stiffness.
Stiffness increases from gastrula to neurula stages
Adult organisms simply cannot be supported by the extremely low stiffness
found in early Xenopus embryos [as low as 3 Pa in some cultures (our
results) (Moore et al.,
1995)]. This paper begins to resolve the striking disparity
between the highly deformable tissues of early embryos and the stiff tissues
found in adults [more than 1000-fold stiffer
(Levental et al., 2007)].
Early Xenopus embryos are some of the most deformable cellular
tissues yet measured, but increase their stiffness by 10- to 50-fold in as
little as 8 hours. Tissue stiffening occurs as the embryo establishes the
basic vertebrate body plan, shapes the neural plate and folds the neural tube
(Davidson and Keller,
1999).
A simple composite mechanical model of the dorsal isolate reveals the relative stiffness of different germ layers
In order to estimate the stiffness of individual tissues that make up the
dorsal axis, we devised a simple analytical model based on the principles of
superposition of composite materials
(Christensen, 1991) (see
Materials and methods). We find that endoderm lining the archenteron roof
plate has the lowest stiffness, nearly matching the stiffness of dorsal
involuting marginal zone explants at early gastrula stages
(Moore et al., 1995), and
indicates that increasing stiffness is not a necessary consequence of
development. The stiffness of microsurgically enhanced explants with two
notochords does not differ from the stiffness of explants where the notochord
had been removed (Fig.
2D'). Likewise, the stiffness of explants with expanded
notochords was no different from control explants
(Fig. 2I) and explants without
a neural plate did not differ in stiffness from intact dorsal isolates
(Fig. 4B). These results reveal
a complex architecture within the early embryo that sets the stage for the
initiation of organogenesis that follows neural tube closure.
Our finding that stiffness of dorsal axial tissues vary in time and
position suggests these mechanical properties may serve multiple roles during
embryogenesis: (1) allowing tissues to serve as a mechanical scaffold for the
action of cell-generated forces (Stern,
2004; Trinkaus,
1984); (2) provide positional cues to pattern cell identity; or
(3) may simply reflect ongoing steps in cell specification and
differentiation. At the present, we can speculate only: that deformable
tissues in the early gastrula are more compatible with large movements, such
as involution and epiboly; or that later convergence and extension or folding
movements of the neural epithelium may require stiffer tissues. It is also
intriguing that germ layers at this early stage exhibit different stiffnesses.
In principle, these spatial variations in the micro-environment may provide
positional information to embryonic cells by triggering alternative
differentiation pathways. Studies of cultured precursor cells grown on stiff
or deformable substrates differentiate according to the mechanical properties
of the substrate, e.g. soft substrates generate adipose cells and stiff
substrates osteocytes (McBeath et al.,
2004). Mechanical cues such as these can equal the strength of
growth factors in contributing to cell fate choices
(Engler et al., 2006). In the
case of the frog embryo, the outer layer of pre-somitic mesoderm faces stiff
neural tissues and may use that information to direct cells to a myotome fate,
while the inner layer faces much more deformable endoderm and could use that
information to initiate a sclerotome fate. The different mechanical
environments seen by the inner versus the outer layer of the presomitic
mesoderm may modulate the cell fate choices of these two adjacent tissues.
Future studies combining biomechanics, signal transduction and classical
embryology will be needed to address these prospective roles for spatial and
temporally regulated tissue mechanics.
|
) and may
be required for laminin polymerization
(Colognato and Yurchenco,
2000); and (4) laminin-1 and fibrillin are strongly localized to
surface of midline axial tissues such as the notochord
(Fig. 5) and removal of the
notochord or addition of a second notochord does not change the stiffness of
isolate from control isolates (Fig.
2). Of course, there may be other fibrillar ECM components that we
cannot yet visualize, but, if they follow similar patterns of assembly and
require a pre-existing fibronectin ECM, they too are unlikely to contribute to
the stiffness of dorsal tissues in the embryo.
The molecular pathways that control tissue stiffness may be the same as those that control actin dynamics during cell motility and tissue elongation
The pathways that regulate actin dynamics during cell movement also control
aspects of tissue stiffness. Contractility within the actomyosin cytoskeleton
can alter its apparent stiffness as it generates pre-stress (see
Wainwright et al., 1976).
Pre-stress can increase the apparent stiffness of a material (known as
`strain-hardening') and is often used by civil engineers in construction of
bridges and skyscrapers. Strain-hardening in response to pre-stress occurs in
both cells (Stamenovic, 2005)
and reconstituted actomyosin gels (Gardel
et al., 2006). In addition, complex regulatory networks can
control the organization and assembly of actin and myosin II within motile
cells. Xenopus embryos, like all other embryos yet studied, exhibit a
wide variety of different cell behaviors that are regulated by these control
networks. Our study, by depolymerizing F-actin or inhibiting myosin II
contractility, has found that these same networks may also control spatial and
temporal patterns of stiffness within the embryo. Surprisingly, although these
acute drug treatments reduce tissue stiffness by half, these changes are much
smaller than the six- to ten-fold spatiotemporal changes measured in the first
half of our study. Further work will be needed to identify both anatomical and
molecular factors responsible for patterning the large-scale changes in
stiffness as the embryo develops and the role that spatial and temporal
changes in stiffness plays in morphogenesis.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/4/677/DC1
|
Footnotes |
|---|
|
REFERENCES |
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TOP
SUMMARY
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RESULTS
DISCUSSION
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