* Department of Pathology, Anatomy, and Cell Biology, Jefferson Medical College, Thomas Jefferson University, Philadelphia,
Pennsylvania 19107; The Burnham Institute, La Jolla, California 92037; § Wellcome Trust Centre for Cell-Matrix Research,
School of Biological Sciences, University of Manchester, Manchester M13 9PT, United Kingdom; and
Kimmel Cancer Center,
Jefferson Medical College, Thomas Jefferson University, Philadelphia, Pennsylvania 19107
Decorin is a member of the expanding group of widely distributed small leucine-rich proteoglycans that are expected to play important functions in tissue assembly. We report that mice harboring a targeted disruption of the decorin gene are viable but have fragile skin with markedly reduced tensile strength. Ultrastructural analysis revealed abnormal collagen morphology in skin and tendon, with coarser and irregular fiber outlines. Quantitative scanning transmission EM of individual collagen fibrils showed abrupt increases and decreases in mass along their axes, thereby accounting for the irregular outlines and size variability observed in cross-sections. The data indicate uncontrolled lateral fusion of collagen fibrils in the decorindeficient mice and provide an explanation for the reduced tensile strength of the skin. These findings demonstrate a fundamental role for decorin in regulating collagen fiber formation in vivo.
Small leucine-rich proteoglycans (SLRPs)1 belong to
an expanding family of secreted proteoglycans that
comprise structurally related but genetically distinct
products, including decorin, biglycan, fibromodulin, lumican, epiphycan, and keratocan (28). The SLRPs share a
common structural architecture that can be divided into
three domains. The amino-terminal domain contains the
negatively charged glycosaminoglycan chains, dermatan or
chondroitin sulfate, or tyrosine sulfate. This region of the
molecule, which contains four conserved cysteine residues,
might be involved in binding to cationic domains of cell
surfaces and extracellular matrix proteins. The cysteinefree central domain comprises ~80% of the protein and is
composed of eight to ten tandem repeats of leucine-rich regions. This region is perhaps the best studied insofar as it has been recently shown that specific amino acid sequences located between repeats 4 and 6 are responsible
for binding to type I collagen (51, 59). If the SLRPs fold in
a manner similar to the leucine-rich ribonuclease inhibitor,
the concave face of the molecule could interact with other
proteins as the ribonuclease inhibitor does with its substrate (33). The carboxyl end domain contains two cysteine
residues, and its function still remains to be elucidated.
The evidence favoring protein-protein interaction as
the major function of SLRPs is persuasive. For example,
decorin binds noncovalently to the surface of fibrillar collagen, primarily type I (53), and retards the rate and degree of collagen fibrillogenesis in vitro (66). This specific
interaction is mediated by the protein core (42), whereas
the glycosaminoglycan chain of decorin extends laterally
from adjacent collagen fibrils, thereby maintaining interfibrillar spacing (52). This lateral orientation has also been
demonstrated in collagen fibrils reconstituted in vitro in the presence of decorin (55). Thus, coordinated expression
of decorin and associated collagens may regulate an orderly matrix assembly.
Decorin purportedly binds to collagen types II (66), III
(60), and VI (6), fibronectin (48), C1q (34), and transforming growth factor- To gain further insights into the functional role of decorin and to explore tissue specificity and functional redundancy during development, we generated mice disrupted
at the decorin gene locus. The nullizygous animals were viable but showed skin fragility with marked reduction in
tensile strength. Compared with normal skin, the collagen
fiber network was more loosely packed with abnormal collagen fibers varying in diameter along their shafts. These
observations provide the first genetic evidence that decorin is essential for proper collagen fibrillogenesis and demonstrate an important role for this proteoglycan in a process fundamental to animal development. Our results also
provide insights into how disruption of collagen fibrillogenesis might result in pathology and predict the potential
existence of a human genetic disease caused by mutations
in the decorin gene locus.
Targeting Vector and Identification of Targeting Events
A 5.5 XbaI decorin genomic fragment isolated from a Germ-Line Transmission, Husbandry,
and Genotype Determination
Targeted ES cells were introduced into mouse blastocysts according to
standard procedures. The resulting chimeras were bred to B1/Swiss females. Tail DNA from B1/Swiss female agouti progeny was tested for the
presence of the targeted allele by Southern blot analysis. Heterozygous
mice from 129Sv X B1/Swiss mixed background were mated to homozygosity. The genotype of each mouse was determined by Southern blotting
and/or PCR analysis of mouse tail DNA. Genomic DNA from mouse tail
fragments was digested with EcoRI and separated on 0.8% agarose gels.
A 310-bp probe encompassing a sequence just 5 Hematological Analysis, RNA Extraction, Proteoglycan
Purification, and Immunoblotting
Approximately 0.5 ml of blood was retrieved by cardiac puncture of homozygous animals and their respective control litter mates (n = 6). A full
body profile, including hematological and clinical chemistry, was performed as described before (4). Differential white blood cell counts were
determined manually on blood smears after Wright staining. RNA was
isolated from mouse tissues using a "Tri" reagent (Molecular Research
Center, Cincinnati, OH), and separated on agarose formaldehyde gels.
Proteoglycans were extracted from ~50 mg of adult mouse tail tissue in 4 M
guanidine HCl, 100 mM sodium acetate buffer, pH 6.0, containing the following protease inhibitors: 1 mM PMSF, 5 mM benzamidine HCl, 5 mM
iodoacetamide, 5 µg/ml leupeptin, 5 µg/ml pepstatin, and 10 mM EDTA.
After a 64-h extraction at 4°C with gentle rotation, the unextracted residue was removed by centrifugation, while the supernatant was dialyzed against 0.1 M NaCl, 0.1 M Tris HCl, pH 7.3, for 72 h at 4°C. An aliquot of
100 µl of the dialysate was incubated with protease-free chondroitinase
ABC (165 mU/ml) overnight at 37°C in the presence of chondroitin 4-S
and 6-S (0.1 µg/ml) as carrier. Total protein was precipitated with 10 vol of
cold 100% ethanol and collected by centrifugation. The resulting pellet
was air dried and resuspended in 40 µl SDS-PAGE buffer. Proteins were
separated on 8-10% polyacrylamide gels and transferred to nitrocellulose
(Hybond ECL; Amersham Corp., Arlington Heights, IL) for 1 h at 700 mA.
Immunoblots were incubated overnight at 4°C in a blocking solution containing TBS-T (20 mM Tris-HCl, pH 7.6, 137 mM NaCl, 0.1% Tween 20)
and 5% nonfat dry milk. After washing the membrane with TBS-T, it was
incubated for 1 h in a 1:5,000 dilution of a rabbit anti-mouse decorin antiserum (LF-113), which was generated against a synthetic decorin peptide from the amino-terminal region spanning residues 36-49 of the mouse
protein (16). The membrane was extensively washed with TBS-T, and then
incubated for 1 h with goat anti-rabbit IgG conjugated to HRP (Sigma
Chemical Co., St. Louis, MO) diluted to 1:10,000 in TBS-T. After further
washing, the antigen/antibody complexes were detected with ECL detection reagents (Amersham Corp.) according to the manufacturer's instructions. Exposure of the immunoblot to XAR5 film (Eastman Kodak Co.,
Rochester, NY) was for 2-10 s.
Morphological and Immunohistochemical Studies
Tissues were fixed in 10% buffered formaldehyde and processed for light
microscopy (27). For immunohistochemistry, paraffin sections were
warmed to 50°C for 30 min, dewaxed in xylene, and rehydrated through
decreasing concentrations of ethanol. Endogenous peroxidase activity was
blocked by a 15-min preincubation with methanolic H2O2 (0.5% vol/vol).
After washing in TBS, pH 7.5, the sections were incubated with normal
goat serum (1:200) for 30 min, washed extensively with TBS, and incubated with the rabbit polyclonal LF-113 anti-decorin or the LF-106 antibiglycan antisera (16) at 1:1,000 dilution for 18 h at 4°C. The LF-106 is directed against a murine biglycan synthetic peptide, amino acids 50-64, conjugated to horseshoe crab hemocyanin (16). An HRP-conjugated goat
anti-rabbit IgG (Sigma Chemical Co.) was applied at 1:200 dilution for 45 min. A 3-amino-ethyl carbazole substrate kit (AEC; Vector Laboratories
Inc., Burlingame, CA) was used to visualize the specific peroxidase activity. The sections were washed in water, counterstained with 0.2% methylene blue, and mounted with Gel/Mount aqueous medium (Biomeda Co.,
Foster City, CA) before photography. For EM, small portions of skin, tail
tendon, or cornea were fixed in 3% glutaraldehyde, 25 mM sodium acetate buffer, pH 5.7, containing 0.3 M MgCl2 and 0.05% cuprolinic blue
(53). The unosmicated tissues were rinsed three times in buffer containing MgCl2, en bloc stained with 1% sodium tungstate, dehydrated in graded
alcohols, and embedded in Spurr's epoxy resin. Thin-sections were observed with a transmission electron microscope 100B (JEOL USA, Peabody, MA), with or without further staining with uranyl acetate (27) or sodium tungstate (53). For quantitative studies, collagen fibril diameters
were measured on photographic prints with a calibrated final magnification of 90,000. Several hundred micrographs were taken from the skin,
tendon, and cornea of five homozygous, two heterozygous and three wildtype animals. A total of 2,803 collagen profiles from wild-type, heterozygous, and homozygous skin was measured, and histograms were generated. For scanning transmission EM (STEM) analysis, 4-mm2 pieces of
skin were finely minced, suspended in 1 ml Tris-buffered (pH 7.4) saline
supplemented with 50 mM EDTA and 100 mM sucrose, and subjected to
mild disruption in a hand-held Dounce homogenizer. The supernatants
were then sampled for EM. Fibrils were adsorbed to 400-mesh carbonfilmed grids, washed with ultrapure water, and air dried. The unstained
fibrils were examined by STEM and mass mapping procedures (21, 22, 23).
Biomechanical Test of Skin Tensile Strength
Dorsal skin was carefully excised and pinned out on a styrofoam support.
Using a plastic template, dumbbell-shaped skin specimens (4 cm × 2 cm)
oriented parallel to the spine were dissected with the aid of a scalpel and
placed between two microscope slides. The specimens were kept moist
with cold saline and transported to the bioengineering laboratory on ice.
The overall thickness of the specimens was measured with a micrometer
while the skin was between glass slides and found not be significantly different among animals. A tensile machine displacement controller manufactured by Comten Industries Inc. (St. Petersburg, FL) was used to measure tensile strength of the skin specimens, which were gripped between specially designed clamps equipped with Velcro-covered faces to prevent
slippage of the tissue. The skin specimens were kept continuously moist
with room temperature saline and stretched at a low speed rate of 1 mm/s
(10%/s) until failure. Tensile strength was computed from maximal load
or force in newton (N) at failure divided by the initial cross-sectional area
of the specimen in the reduced test area (32, 46).
Generation of the Decorin Null Mice
The genomic locus encoding the murine decorin gene was
targeted with a replacement construct containing ~5.5 kb
of DNA sequence interrupted in exon 2 by the neomycin
resistance cassette driven by the Pgk promoter (Fig. 1 A).
We made use of a positive/negative selection scheme by
using neomycin resistance for positive selection and diphtheria toxin-A driven by the thymidine kinase promoter for negative selection of nonhomologous recombinants
(70). 110 G418-resistant ES clones were isolated and expanded. Homologous recombination was detected by Southern blotting with a single genomic probe flanking the construct. This probe detected a 6.5-kb EcoRI fragment
specific for the targeting vector (the neo cassette has an internal EcoRI site), and a 7-kb EcoRI fragment specific for
the wild-type allele. Among the first 51 clones that were screened by Southern blotting, 13 (~25% targeting frequency) had the predicted 6.5-kb EcoRI band corresponding to the targeted allele (not shown). Two independent
ES clones with a normal karyotype were used to produce
male chimeras that were able to affect germ-line transmission of the disrupted allele when bred to C57BL/6 females. Crosses of heterozygous parents resulted in the birth of
mice that were homozygous for the mutant Dcn gene
based on either Southern blotting (Fig. 1 B) or PCR (Fig.
1 C) of tail biopsy DNA.
Homozygous Mutant Mice Are Viable
Mice heterozygous for the decorin knockout mutation
(129Sv X Bl/Swiss mixed background) were mated to homozygosity. The resulting progeny (n = 69) showed a standard Mendelian transmission: 16 homozygotes (23%), 37 heterozygotes (54%), and 16 wild type (23%). This Mendelian distribution was subsequently observed in several progenies derived from mating heterozygous animals. To
confirm the disruption of both decorin alleles, the absence
of decorin-specific transcript was checked by Northern
blotting analysis. Total RNA from several major tissues,
including manubrium sterni (which contains cartilage,
bone, and bone marrow), heart, liver, kidney, and skin, an
organ that is known to express high levels of decorin message (49), showed no detectable decorin-specific transcript (Fig. 2 A). Thus, both copies of the decorin gene were successfully disrupted by homologous recombination initiated
by our targeting vector. As expected, the levels of decorin
were slightly reduced in the Dcn+/
Lack of Decorin Leads to Thinning and Fragility
of the Skin
Initial analysis of the Dcn
The rupture of the skin similar to that shown in Fig. 3 A
had a considerable penetrance, insofar as it was observable in >50% of all the Dcn Skin of Decorin-deficient Animals Exhibits Reduced
Tensile Strength
It is generally accepted that skin strength correlates directly with the overall organization, content, and physical
properties of the collagen fibril network (15). Therefore,
to achieve a more objective measurement of skin fragility,
we determined the tensile strength in the physiological
range of loading (low rate of loading) in portions of dorsal
skin from six animals, three wild-type and three Dcn
A Structural Basis for Skin Fragility in the
Decorin-deficient Mice: Abnormal Morphology of
Collagen Fibrils and Reduction of
Collagen-bound Proteoglycans
Ultrastructural analysis of dermal collagen from the skin
of 3-mo-old Dcn
Ultrastructural analysis of tail tendon collagen revealed
even more bizarre profiles and shapes. Individual fibrils
exhibited very irregular, ragged outlines in cross-section
compared with normal skin. The collagen fibrils reached
660 nm in diameter (Fig. 6, A and B) in contrast with the
wild-type animals where the average was about 200 nm
(Fig. 6 C). The aberrant fibrils appeared to be due to multiple and concurrent lateral fusions of their central thick
segments to thinner fibrils (Fig. 6 B). Moreover, intermingled with these gigantic fibrils were numerous thin fibrils
with an average diameter of 40-60 nm. These thin fibrils
were less frequently detected in tendons of age-matched
controls. Since decorin is also expressed in the cornea, we
analyzed the ultrastructure of the corneal collagen in wildtype, heterozygous, and homozygous animals. We found
no significant changes in either packing or overage size of
corneal collagen in the Dcn
To investigate further the role of proteoglycans in collagen fibrillogenesis, we processed skin and tendon in the
presence of cuprolinic blue, a cationic dye that preserves
proteoglycans in an extended configuration in the presence of 0.3 M MgCl2 (53). Under such high salt concentration, binding of the dye is specific for proteoglycans with
sulfate esters. Using this method, decorin has been localized to the d and e bands in the D-gap zone of collagen quarter-stagger in tendon, cornea, and pericardium (52,
53, 56), a localization confirmed by immunoelectron microscopy (43, 56) and in situ staining of isolated molecules
(35). When collagen fibrils from tail tendons were fixed in
the presence of cuprolinic blue and stained with uranyl acetate to visualize the banding pattern, it was evident that
the collagen of affected animals (Fig. 6 E) exhibited the
67-nm periodicity characteristic of type I collagen. As expected, in the Dcn+/+ animals, all the d bands were occupied (Fig. 6 D, rows of arrowheads). In contrast, in the
Dcn Mass Mapping of Dermal Fibrils from
Decorin-deficient Animals Reveals Pronounced
Nonuniformity in the Axial Mass Distribution
Next, we wished to determine whether the collagen fibrils
from the skin of Dcn
Collagen provides the principal source of mechanical
strength in connective tissues and is often implicated in
diseases where such mechanical support is defective (44).
The molecular interactions that take place among various
types of fibril-forming collagens and members of the
SLRP gene family constitute a fundamental regulatory mechanism for the assembly of heterotypic fibrils. The results of the present investigation clearly show the developmental significance of decorin as a key regulator of collagen fibrillogenesis and validate previous observations
that the core protein of decorin plays a central role in regulating fibrillogenesis (10, 66, 67). Given the existence of
several members of the SLRP gene family that could conceivably compensate for the lack of decorin function, the
observed phenotype was not entirely predictable. We observed skin fragility and abnormal fibrillogenesis in dermal and tendon collagen, but not in corneal collagen, indicating tissue specificity and a requirement for decorin for
proper skin and tendon development and maintenance of
normal tensile strength.
Decorin Gene Knockout Causes Abnormal Collagen
Structure and Skin Fragility
The cross-sections of fibrils in skin and tendon showed
regular, uniform fibrils in control animals. In contrast, in
the decorin-deficient animals, the collagen fibrils were
coarser and irregular in size and shape. Furthermore, they
were often haphazardly arranged with increased interfibrillar spaces. These structural changes were associated
with a reduction in collagen-bound proteoglycans in both
dermal and tendon collagen. Of note, the STEM data
showed that the fibrils in null mice were not uniform in diameter but had bulges along their shafts. These data thus
explain the variability in size and shape seen in cross-sections of the skin and tendon collagen. There are two possible mechanisms to explain the formation of bulges along the
axis of individual collagen fibrils. First, the bulges (abrupt
increases in mass per unit length) could occur when early
fibrils fuse laterally onto an existing larger fibril. Our
quantitative data (Fig. 7 H) show that, as the fibril diameter increases in control animals, the bulge factor decreases.
Thus, with increasing fibril diameter there is less possibility of fibril fusion. This is the reverse in the decorin-deficient mice; with increasing fibril diameter, the bulge factor
increases. Interestingly, at a fixed decorin/collagen ratio,
with increasing fibril diameter, the surface area/volume ratio decreases and the concentration of decorin at the fibril
surface increases. Therefore, in control mice, as the fibril
diameter increases, the concentration of decorin at the
fibril surface increases and fibril fusion is inhibited. This
mechanism of restricting fibril fusion to small diameter fibrils is lost in the decorin-deficient mice. The second possible explanation is that collagen fibrils might have a natural tendency to be irregular. Decorin, in some unknown
way, would keep the fibrils regular in outline. There is in
vitro evidence against this interpretation because fibrils
formed from purified collagen solutions tend to be fairly
uniform in cross-section (31). Furthermore, collagen fibrils
assembled in vitro from purified acid-soluble calf skin collagen participate in lateral association and fusion in the
absence of decorin (unpublished observations). The most severely affected tissue in decorin-deficient mice is skin,
which cannot withstand sudden tensile strain as its normal
counterpart does. The abnormal packing of collagen together with the nonuniformity of fiber diameter may have
a considerable influence on the biomechanics of this tensile tissue. Skin fragility in these mutant animals could be
ascribed to this anomalous collagen network, which would allow full body development and viability, but would lead
to a reduced tensile strength with potential complications
such as increased incidence of injury, infection, and scarring. Biomechanical measurements using physiological stress
rates (i.e., slow stretching rates of 1 mm/s) revealed at least
a threefold reduction in tensile strength of dorsal skin
samples with a concurrent 35-40% reduction in ductility.
Of note, using similar low speed stress rates, it has been recently shown that the tensile strength of rat skin correlates
with insoluble collagen and is decreased when cross-linking of collagen is blocked by lathyrogen treatment (15).
This is similar to the decrease in mechanical strength detected in lathyritic rat tail tendon (19).
The presence of small proteoglycans occupying the d
band of collagen fibrils in the decorin knockout animals
suggests that other members of the SLRP gene family may
bind the same sites. However, neither biglycan nor lumican were upregulated at the mRNA level, thus suggesting
that there is no obligatory compensation for the lack of
decorin by at least two members of the SLRP gene family.
The absence of overt abnormalities in the cornea of
Dcn Decorin: A Key Regulator of Collagen Fibrillogenesis
Connective tissue interactions, including those between proteoglycans and collagen, are evolutionarily conserved and
physiologically relevant. High affinity interaction of dermatan sulfate proteoglycan with collagen was inferred three
decades ago when extraction with 6 M urea or digestion
with collagenase was required for efficient isolation of this
proteoglycan from the skin (61). Moreover, it is firmly established that (a) fibrillogenesis is a multistep assembly
process (64); (b) collagen molecules incubated in vitro at
neutral pH and physiologic temperatures aggregate and
develop into insoluble fibrils of various sizes and lengths
(31); (c) unipolar and bipolar fibrils occur in vivo (23); (d)
collagen fibrils grow from paraboloidal tips and this mechanism is highly regulated (23, 30); (e) the size and shape
of fibrils are enzymatically controlled by the procollagen
C-proteinase (26); and (f) this process can be modulated
by skin proteoglycans (62). When the kinetics of in vitro
fibrillogenesis were monitored by turbidimetry, it was discovered that decorin from tendon (66) or cornea (45) significantly delayed collagen fibrillogenesis. The lateral assembly of triple-helical collagen molecules was also delayed by
decorin resulting in thinner fibrils (8, 9, 65). During development, the ratio of dermatan sulfate to hydroxyproline decreases as the fibril cross-sectional areas increase (53), an observation corroborated by the finding that fibril
growth is associated with a significant decrease in fibril-
associated decorin (8). The corollary, that thicker fibrils
are associated with less dermatan sulfate/unit weight of
collagen, can be inferred from the fact that the upper dermis of calf skin containing finer fibrils has twice the dermatan sulfate/collagen ratio of deeper dermis containing
coarser fibrils (53). Of note, in human skin, the papillary
(upper) dermis expresses higher levels of decorin than
the reticular (deeper) layer (50). Collectively, these findings favor a role of decorin in permitting lateral association of fibril segments, a concept also supported by the
three-dimensional molecular model of decorin (69). Binding of decorin to collagen would interfere with further lateral accretion of collagen molecules and retard/stop the
growth of fibrils. Therefore, decorin would play a pivotal
role in regulating the orderly assembly and growth of collagen fibrils, which in turn would affect the tensile strength of connective tissues. The abnormal collagen morphology
and the STEM data fully support this interpretation.
Relation of the Decorin-deficient Phenotype to
Dermatosparaxis and Ehlers-Danlos Syndrome
Dermatosparaxis, "torn skin" syndrome, is a recessively
inherited connective tissue disorder that results from the
lack of activity of type I procollagen N-proteinase, the enzyme that removes the amino-terminal propeptide from
type I procollagen (36). Initially identified in cattle more
than 25 years ago, this disorder has now been reported in
sheep, cats, and dogs (5, 13, 18). Affected animals have
skin fragility, joint laxity, and often succumb prematurely
to sepsis after avulsion of portions of the skin (18). The
hallmark of the disease is the presence of abnormal collagen fibrils that are thinner and haphazardly arrayed, and often in cross-section resemble hieroglyphs. A similar syndrome has been now reported in three patients with skin
fragility and twisted, ribbon-like collagen fibrils (40, 41, 57).
The human syndrome, designated Ehlers-Danlos syndrome
(EDS) VIIC, differs from the EDS VIIA and VIIB insofar
as the latter two are caused by mutations of the NH2-terminal propeptide of collagen Transgenic mice homozygous for an in-frame deletion
of exon 6 of the collagen This study has been fruitful in establishing a functional
role for a secreted proteoglycan in maintaining the structural integrity of the cutis and perhaps tendon sheaths.
Tensile strength is certainly a major property of connective tissues, and our data demonstrate that decorin stabilizes the fibrillar matrix in vivo. Decorin governs collagen
fibril growth and could influence higher order matrix assembly. Understanding how changes in decorin alter the biological properties of collagen and disrupt the finely balanced network of molecular interactions that govern fibril
assembly and architecture may lead to the discovery of
novel genetic disorders caused by members of the SLRP
gene family, and could potentially lead to the development
of medical strategies for their treatment. In addition to
helping to elucidate the biological role of decorin, the
decorin-deficient mouse could represent a useful animal
model to investigate matrix assembly, wound healing, and
tumorigenesis. On the basis of our animal phenotype, we
predict the existence of a human skin fragility syndrome,
to be included in the EDS category, which could be caused
by either a recessive deletion or a mutation in the collagen
binding domain of the decorin gene at the 12q23 locus.
(20). Moreover, decorin has been implicated in the control of cell proliferation by inducing arrest of tumor cells in the G1 phase of the cell cycle (14, 47).
Materials and Methods
FIX II genomic
mouse 129Sv library (49), isogenic to the embryonic stem (ES) cells, was
used to construct the decorin targeting vector. This genomic fragment, encompassing exons 1 and 2, was ligated into pBluescript KS with a deleted
EcoRV site, and the resulting fragment was designated pMD. A plasmid
containing Pgk-neo (58) was digested with XhoI and EcoRV, and the
XhoI site was made blunt-ended with Klenow polymerase. The resulting
Pgk-neo fragment of 1.6 kb was ligated into a unique EcoRV site of exon
2, thereby dividing the genomic fragment into two arms of homology of
~3.8 and ~1.7 kb to the targeted locus, respectively. To enrich for targeting events, the diphtheria toxin-A cassette driven by the thymidine kinase
promoter (70) was added downstream of the targeting vector into the
XhoI site of the multiple cloning site of pMD. Linearization of this targeting vector was done with NotI before electroporation into the ES cells.
The R1-ES cells (39) were cultured in standard ES cell culture conditions
(2), in DME supplemented with 15% FCS, 0.1 mM
-mercaptoethanol,
and 1,000 U/ml of human lymphocyte inhibitory factor (h-LIF) on neomycin-resistant primary cultures of embryonic fibroblast feeders. LIF was
produced from the CHO cell line stably transfected with the h-LIF cDNA.
Cells were electroporated with 40 µg/ml of decorin targeting vector. The
resulting G418-resistant clones were isolated by ring-cloning and expanded (3). Approximately 100 clones were analyzed by Southern blot
analysis for the detection of the correct targeting events (see below).
to the targeting vector
was prepared by PCR using the T7 primer from a subcloned 5.5-kb genomic fragment of mouse decorin phage clone F 11-1 (49) and an antisense primer specific to exon 1. This probe detects a 7- or 6.5-kb sequence
of DNA digested with EcoRI derived from normal wild-type and targeted
animals, respectively. Alternatively, PCR was used to detect the presence
of homologous recombination of the decorin gene. Sense and antisense
primers corresponding to exon 2 of murine decorin (5
-CCTTCTGGCACAAGTCTCTTGG-3
and 5
-TCGAAGATGACACTGGCATCGG-3
)
were used to detect a 161-bp fragment, indicating lack of homologous recombination. An additional primer corresponding to the Pgk promoter of
the Pgk-neo cassette (5
-TGGATGTGGAATGTGTGCGAG-3
) was
used to detect a 250-bp fragment, indicating the presence of homologous
recombination. The reaction mix of 28 µl consisted of 100-500 ng genomic
DNA, 100 ng each of the three primers, 1.8 mM MgCl2, 1× buffer, 360 µm
dinucleotide triphosphates, and 2.5 U of Taq polymerase. Reaction conditions were: 1 min at 95°C, 20 s at 57°C, and 30 s at 72°C for 35 cycles.
Results
Fig. 1.
Disruption of the
decorin gene locus in mouse
ES cells and generation of
decorin-deficient mice. (A)
Targeting strategy. A 5.5-kb
XbaI genomic fragment encoding exons 1 and 2 (filled
boxes, not in scale) was used
to construct the targeting
vector. The predicted structure of the disrupted allele
(bottom panel) shows the 5
probe used to detect the diagnostic allele of 6.5 kb in contrast with the wild-type
allele of 7.0 kb. This was due
to the presence of a new
EcoRI site in the Pgk-neo
cassette. Abbreviations for
restriction endonucleases: E,
EcoRI; X, XbaI; N, NotI; Xh, XhoI. (B) Southern blot
analysis of tail DNA isolated
from two separate litters of
mice including wild-type (+/
+), heterozygous (+/
),
and homozygous (
/
) animals. Lanes 1-3 are from
3-mo-old animals derived
from the breeding of two heterozygous mice, while lanes
4-9 are from newborn animals derived from the breeding of a heterozygous male
and a homozygous female.
The targeted allele of 6.5 kb
is labeled by an asterisk. The
DNA was separated on a
0.75% agarose gel, transferred to a nitrocellulose filter, and hybridized under
high stringency to a PCRgenerated probe 5
to the
targeting vector. The size of
molecular weight markers is indicated in the left margin in
kb. (C) PCR detection of the
targeted allele using primers specific for exon 2 (a and c, top scheme) or Pgk-neo (primer b). By using primer a and c, a fragment of 161 bp was identified in the wild-type (lane 1) and heterozygous animal (lane 2). However, the combination of primer b and c gave rise to a
larger fragment of 250 bp, encompassing the 3
end of the neo cassette and the 3
end of exon 2, which was detected only in the heterozygous (lane 2) and homozygous (lane 3) animals. The products were separated on a 6% nonreducing polyacrylamide gel and
stained with ethidium bromide. The size in bp is indicated in the left margin.
[View Larger Versions of these Images (14 + 32K GIF file)]
animals when normalized on the housekeeping gene GAPDH. Moreover,
when the Northern blots were hybridized with either a
full-length cDNA probe encoding murine biglycan or a reverse transcriptase-PCR-generated probe encoding a portion of murine lumican, no significant change in either
transcript was observed (not shown). These findings indicate that neither biglycan nor lumican (murine fibromodulin probe was not available) is upregulated to compensate
for the lack of decorin in any of the tissues examined so
far. The results obtained at the mRNA levels were confirmed by the absence of the immunoreactive decorin proteoglycan from whole tail extracts (Fig. 2 B). When partially purified proteoglycans were separated by 7.5% SDS
gel electrophoresis and immunoblotted, no decorin immunoreactivity was observed by using a specific antipeptide antibody (16) either before (Fig. 2 B, lane 3) or after (Fig. 2 B, lane 6) chondroitinase ABC treatment, in spite of the
fact that approximately equal amounts of protein were
loaded on the gel. Collectively, these results indicate that
disruption of both alleles of the mouse decorin gene results in abrogation of decorin message and protein.
Fig. 2.
Absence of decorin
mRNA and protein in animals harboring a homozygous disruption of the decorin gene. (A) Northern blot
analysis using total RNA
prepared from selected tissues as indicated of wild-type
(+/+), heterozygous (+/),
and homozygous (
/
) adult animals. High stringency hybridization was performed
using cDNAs encoding the
full-length decorin or the
housekeeping gene GAPDH as 32P-labeled probes (49).
Notice the lack of decorin transcripts in the Dcn
/
animals (lanes 9-12, 15, and 16). (B) Immunoblot analysis of guanidine HCl extracts of tail tissues before or after chondroitinase ABC (+ABCase) digestion using an anti-decorin antibody. Notice the presence of a
proteoglycan centering around 94 kD and the presence of a 45-kD unprocessed protein core in the wild-type (lane 1) and heterozygous
animal (lane 2); no immunoreactive material was detected in the homozygous animal (lane 3). After chondroitinase ABC digestion, the
high molecular mass proteoglycan was converted into two broad bands of 45-50 kD (lanes 4 and 5). Decorin-specific epitopes were detected with a polyclonal antibody (LF-113) directed against a synthetic decorin peptide from the amino-terminal region spanning residues 36-49 of the mouse protein (16).
[View Larger Version of this Image (24K GIF file)]
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mice did not reveal any
gross anatomical abnormalities. The animals grew to normal size, were fertile, and did not exhibit obvious behavioral deficiencies. A complete serum chemistry and hematological profile of 2-mo-old homozygous and control mice
did not show any significant difference in the levels of albumin, globulin, electrolytes, sodium, and chloride, in the
hepatic enzymes, ALT and AST, or in the number of
white and red blood cells, hematocrit and hemoglobin. Radiographical studies of whole animals did not reveal any
overt bone abnormalities in the decorin-deficient animals
(not shown). However, closer analysis of the Dcn
/
mice
revealed skin laxity and significant fragility. When 3-moold mice were sacrificed by cervical dislocation, the simple
application of pressure to the cervical region caused an
abrupt rupture of the skin (Fig. 3 A, arrows). In addition the skin of the tail was detached in its entirety from the underlying soft tissues (Fig. 3 A, arrowheads). Light microscopic observation of the skin from the tail showed a sharp
detachment of the skin between the deeper dermis and the
fascia, with clean, sharp edges along the dissection (Fig. 3 B,
arrowheads). Random sampling of abdominal skin from
several Dcn
/
mice revealed dermal thinning and loose
connective tissue in the hypodermal layer (Fig. 3 D). Thus,
skin laxity observed in the decorin-deficient animals could be due at least in part to accumulation of loose connective
tissue in the dermal and hypodermal layers. In contrast,
age-matched Dcn+/+ mice littermates (Fig. 3 C) or Dcn+/
animals (not shown) showed no such structural changes.
To confirm the absence of decorin, immunohistochemical
analysis using the anti-decorin antibody showed no detectable epitopes in the skin of Dcn
/
animals (Fig. 3 D). In
addition, all the tissues tested, including skeletal muscle, adipose tissue, and uterus (Fig. 3, D and F), as well as
Fallopian tubes and ovaries (not shown), revealed no decorin immunoreactivity. In contrast, the wild-type animals
showed the expected immunoreactivity in the dermis (Fig.
3 C, asterisks), in the myoepithelial layer and connective
tissues surrounding mammary ducts (Fig. 3 E, arrows), and
in the adventitia of small blood vessels (Fig. 3 E, arrowheads). In the Dcn
/
animals, staining with an antibody
against biglycan showed the presence of this epitope in the
endometrium and intramural blood vessels (Fig. 3 G), as
well as in the epidermal layer and follicular epithelium
(Fig. 3 H).
Fig. 3.
The phenotype of decorin-deficient mice reveals thinning and fragility of the skin. (A) Gross photograph of wild-type (+/+), heterozygous (+/), and homozygous (
/
) littermates. Notice the sharp rupture of the back skin in the
/
animal (arrows) and the
detachment of the tail skin (arrowheads) that occurred during cervical dislocation. The skin fragility was never observed in either +/+
or +/
animals (n > 300). B is a cross-section of the detached tail skin. Notice the sharp and bloodless line of rupture (arrowheads)
along the deeper dermis. Immunohistochemical analysis of skin and skeletal muscle (C and D), mammary gland (E), and uterus (F)
from wild-type (+/+) and decorin-deficient (
/
) animals using the LF-113 anti-decorin antibody. Notice the intense immunoreactivity in the dermis of a wild-type animal (C, asterisks) and the fine immunoreactivity on either side of the skeletal muscle (Mu). In contrast,
the
/
dermis (D, asterisk), the subcutaneous and perimysial connective tissues, are totally unreactive as are the uterine wall and mucosa (F). As expected, the anti-decorin antibody labeled specifically the adventitia of small blood vessels (E, arrowheads) and the myoepithelial cells and fine connective tissue surrounding mammary ducts (E, arrows). Immunodetection of biglycan using an anti-peptide
(LF-106) antibody in uterus (G) and skin (H) from Dcn
/
mice. Notice the "normal" expression of biglycan in the endometrium (G,
arrows) and in the intramural small blood vessels (G, arrowheads). As expected, in skin, both the epidermis (H, arrowheads) and the follicular epithelium (H, arrow) were labeled by the anti-biglycan antibodies. Sections were reacted with LF-113 or LF-106 antisera at 1:
1,000 dilution, and then visualized with peroxidase-conjugated IgG (1:200) followed by counterstaining with 0.2% methylene blue. Bar,
100 µm.
[View Larger Version of this Image (114K GIF file)]
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mice. In contrast, skin
fragility was never observed in the heterozygous, wild-type
animals; nor was it encountered in other animals (n > 300)
with similar genetic background but harboring a wild-type
allele. These data indicated that, to obtain the skin fragility phenotype, both decorin alleles had to be inactivated.
/
mice. Two fresh specimens per animal of identical dumbbell-shape were generated on a plastic template, and load/
displacement curves were determined by subjecting them to a stress rate of 1 mm/s (10%/s) with constant recording
by a Comten tensile machine displacement controller. Tensile strength was expressed as the maximal load or force in
newton (N) before skin failure divided by the initial crosssectional area (1 cm2) of the specimen in the central test
area (46). A typical experiment derived from stretching
fresh dorsal skin from two age-matched animals is shown
in Fig. 4. The skin from the decorin-deficient animals exhibited a premature failure at ~7 N, in contrast with the
skin from the wild type that ruptured at ~27 N. The overall values of tensile strength measurements were significantly reduced in Dcn
/
animals (7 N ± 2) as compared
with the wild-type animals (21 N ± 5). The data further
suggest that the decorin-deficient mice exhibited a reduction in ductility, i.e., the percentage of deformation before
failure (calculated from the displacement values in Fig. 4).
The normal values centered around 170% vs 110% for the
decorin-deficient animals. These independent biomechanical parameters thus confirm the observation of skin fragility reported above and indicate that the decorin-deficient
phenotype is due to a decline in the intrinsic tensile
strength of the skin.
Fig. 4.
The skin of decorin-deficient mice has reduced tensile
strength. Freshly excised, dumbbell-shaped samples (4 × 2 cm) of
dorsal skin oriented parallel to spine from +/+ () or
/
(
)
animals were generated on a plastic template, thereby producing
skin specimens with a narrow central region (1 cm2) where failure
would occur by avoiding stress concentration at the jaws of the
grip. The samples were gripped into a tensile testing machine
(model 4Z387A; Comten Industries) and stretched to tensile failure at a constant strain rate of 1 mm/s, equivalent to 10%/s. A
data collecting system linked to a Macintosh Quadra computer
(Apple Computer Inc., Cupertino, CA) was used to generate load/displacement curves. Peak of curve represents point of failure. Tensile strength was computed from maximal load at failure
divided by the initial cross-sectional area of the specimen in the
reduced test area. Six age-matched animals were analyzed with
two specimens per animal. Data points represent a typical experiment from one animal each.
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animals revealed aberrant organization of collagen fibrils with several distinct structural abnormalities. First, dermal collagen was less orderly packed
(Fig. 5 A) than age-matched wild type (Fig. 5 C). Second,
there was great variability in shape and size (Fig. 5, A and
B); individual fibrils exhibited irregular outlines in crosssection compared with the circular outline of controls. In
some instances, the cross-sectional profiles of collagen fibrils in affected dermis reached very large diameters up
to 240-260 nm (Fig. 5 A, asterisks) and showed scalloped
edges (Fig. 5 A, arrowhead) or focal lateral fusion with
thinner fibrils (Fig. 5 B, arrowheads). In contrast, heterozygous (not shown) or wild-type skin (Fig. 5 C) showed
collagen fibrils with fairly uniform, circular cross-sectional
profiles. Third, there was the coexistence of thin collagen
fibrils (Fig. 5 A, circles) intermingled with large ones,
whereas in the wild type the average size was quite uniform. As a result of the overall round shape of the collagen fibrils, it was possible to measure the average diameter of several hundred cross-sectional profiles. The results
demonstrated that the average collagen diameter did not
significantly vary between wild-type (Fig. 5 D) and homozygous animals (Fig. 5 E), 116 ± 18 (n = 788) vs 119 ± 35 (n = 795), respectively. However, the Dcn
/
animals exhibited a wider range with profiles varying between 40 and 260 nm, confirming the qualitative data shown above.
In contrast, the wild-type animals contained profiles ranging between 40 and 180 nm. The heterozygous animals had
a distribution and an average collagen diameter (118 ± 14;
n = 1,220) nearly identical to the wild type (not shown).
Fig. 5.
Ultrastructural analysis of skin from the decorin-deficient mice reveals abnormal collagen fibrillogenesis. (A-C) Transmission electron micrographs of dermal collagen from Dcn/
(A and B) and Dcn+/+ (C) animals. Notice the presence of
larger (>200 nm) and irregular fibrils (A and B, asterisks) and the
coexistence of smaller (30-40 nm) fibrils (A, circles) in the Dcn
/
.
Note also the presence of coarser fibrils exhibiting lateral fusion to an adjacent tapered segment (B, arrowheads). In contrast, collagen from the wild-type mouse (C) showed a more compact and
uniform pattern of fibril diameter and distribution. The Dcn+/
collagen pattern was identical to the Dcn+/+ animals (not
shown). (D and E) Distribution of collagen fibril diameter in dermal collagen from Dcn+/+ (D) and Dcn
/
(E) animals. Notice
that, although the mean diameter is not significantly different between the groups, the range and distribution is quite different in
the Dcn
/
animals. Heterozygous animals had a distribution
and an average collagen diameter (118 ± 14; n = 1,220) nearly
identical to the wild type (not shown). Bar, 0.2 µm.
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animals (not shown).
Fig. 6.
Decorin-deficient
mice reveal abnormal collagen in tail tendon and a decrease in collagen-associated
proteoglycans. (A) Lower power view of cross-sectional
area of adult tail tendon collagen from Dcn/
reveals
abnormal shape and multiple
lateral fusions. Higher
power view of Dcn
/
tendons shows bizarre, gigantic
fibrils (>660 nm in largest diameter) with multiple lateral
fusions (B, arrowheads), in
contrast with the more uniform collagen fibrils of the
wild-type animals (C) with
an average diameter of ~200 nm. Transmission electron
micrographs of longitudinally sectioned collagen
fibrils from Dcn+/+ (D)
and Dcn
/
(E) tendon, after fixation in the presence of
0.05% cuprolinic blue and
0.3 M MgCl2, and staining
with uranyl acetate to visualize the collagen binding pattern. Notice that the typical,
67-nm periodicity of the type I collagen is maintained in
the Dcn
/
mice (E). However, numerous d bands are
not occupied by proteoglycan granules (E, spaces between arrowheads and unfilled arrowheads), in
contrast with the wild type
where nearly every d band is
occupied (D). Bar 0.2 µm.
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/
animals, several d bands contained no orthogonally arrayed proteoglycan granules (Fig. 6 E, spaces between arrowheads and also unfilled arrows). Moreover, we
found a significant reduction of proteoglycan granules and
filaments around dermal collagen fibrils of homozygous
animals when compared with wild type (not shown). The
proteoglycans appeared smaller in average length in the
Dcn
/
, likely reflecting the presence of keratan-sulfate- containing proteoglycans, such as fibromodulin and lumican, that are known to be expressed in both tendon and
skin (28). Collectively, these results provide a structural
basis for the phenotype described above and further indicate that the abnormal collagen morphology is associated
with a decrease in collagen-bound proteoglycans. The loss
of such important protein/protein interaction sites could significantly contribute to the abnormal collagen fibrillogenesis described above.
/
animals were uniform along
their axes. This was investigated by determining the axial
mass distributions of unstained, isolated fibrils using STEM.
Dispersed fibrils of up to ~55 µm (~800 D-periods; 1 D = 67 nm) in length were prepared, and montages of STEM
images were recorded along the entire lengths of each
fibril. Consecutive mass per unit length measurements (averages over two D-periods) were made along each fibril.
Typical results are shown in Fig. 7. Fibrils from Dcn
/
animals showed pronounced nonuniformity along their axes
compared with the Dcn+/+ control fibrils. A typical unstained fibril from the homozygous animals showed repeated bulges along the axis (Fig. 7 A). Axial mass distributions of these fibrils revealed a series of mass peaks,
often with a factor of two or three times the mass per unit
length of the adjacent parts of the fibril, and each typically extending 50-80 D-periods along the fibril (Fig. 7, D-F).
In contrast, fibrils from the wild-type animals appeared
uniform in STEM images (Fig. 7 C), and axial mass measurements confirmed that these fibrils lacked the mass
peaks seen regularly in the homozygous animals (Fig. 7 G).
A degree of nonuniformity existed in control fibrils (Fig.
7 G), although this was much less pronounced than that
seen in Dcn
/
mice (Fig. 7, D-F). To quantify the
changes in mass per unit length along individual fibrils, a
simple measure of fibril uniformity was defined as the
nonuniformity index, which was equal to the range of mass
per unit length for each fibril divided by the mean mass
per unit length (Fig. 7 H). Analysis of fibrils from Dcn+/+
mice showed that the nonuniformity index decreased with
increasing fibril mass per unit length (i.e., increasing fibril
diameter). Therefore, the fibrils with the most relatively uniform shafts were the larger diameter fibrils. In contrast, analysis of fibrils from Dcn
/
mice showed that the index increased with increasing fibril mass per unit length.
The lumpy nature of axial mass distribution profiles of the
fibrils from the Dcn
/
animals is consistent with a lateral fusion of short fibril segments with a longer fibril. The
occurrence of lateral associations of long fibrils is also indicated by the observation of twisted pairs of fibrils (Fig.
7 B). These results strongly indicate that the larger and irregular skin fibrils seen in transverse sections are likely to
be a consequence of lateral fusion, producing a fibril whose
morphology changes continuously along its axial length.
Fig. 7.
Mass mapping of collagen
fibrils isolated from the skin of decorin-deficient animals reveals pronounced nonuniformity in the axial
mass distributions with repeated
bulges. (A-C) Typical dark-field STEM images of isolate unstained
fibrils from mouse skin. A and B show
typical images for fibrils from a decorin-deficient mouse. A shows a single
fibril with distinct bulging at two locations over 60 D-periods (1 D = 67 nm)
length. B is consistent with two fibrils
twisted together in close contact but
not fused. C shows a typical fibril from
control skin with uniform diameter
along its length. (D-G) Axial mass distributions of single fibrils obtained by
measurements from STEM images. A
measurement of mass per unit length
was made every two D-periods along
the entire available length of each
fibril. Missing points occur where parts
of a fibril were clearly contaminated.
(D-F) Typical axial mass distribution profiles from the decorin-deficient
sample or (G) from wild-type animals.
The decorin-deficient sample shows
pronounced mass peaks in the fibrial
axial mass distribution, which are not
found in the control. (H) Scatter plot
of the relative variation in M/L along
single fibrils against the corresponding
mean M/L. First order regression lines
are shown superimposed on each set of
data. The data from the decorin-deficient samples show an overall increase in the relative intrafibrillar variation in
M/L with increasing lateral size (Mean
M/L); this contrasts with the control
sample that shows the opposite trend.
Bar, 200 µm.
[View Larger Version of this Image (36K GIF file)]
Discussion
/
animals may be explained by the larger requirement for keratan sulfate in this transparent structure and
by the presence of at least five members of the SLRP gene
family, which could control more tightly the fibril diameter
necessary for transparency. Of note,
-d-xyloside, a xylose
analogue that interferes with the synthesis of chondroitin/
dermatan sulfate but not keratan sulfate proteoglycans,
causes no alterations in fibril diameter but induces disruption of lamellar organization and fibril packing in developing avian cornea (17). These findings therefore suggest that
decorin may play a less prominent role in the regulation of
collagen fibril diameter in cornea, but is important in maintaining the interfibrillar spacing and lamellar cohesiveness.
1(I) and
2(I) genes, respectively (12). The mutations in EDS types VIIA and B
result in the skipping of exon 6, which contains the cleavage sites for the N-proteinase, thereby leading to accumulation of unprocessed collagen. Intuitively, it might have
been expected that a deficiency in N-proteinase activity
and the lack of the cleavage site for the enzyme would
have given rise to the same disease. However, EDS types
VIIA and B are distinct from dermatosparaxis in that (a)
joint hypermobility, rather than skin fragility, is the principal clinical feature, and (b) ultrastructural analysis of the
dermal collagen shows rough-bordered collagen fibrils
rather than hieroglyphs (24, 68). A contribution factor to
the skin fragility in dermatosparaxis may be the steric
blocking of a factor to the gap zone of the collagen fibrils
insofar as the amino-terminal propeptide would interfere
with this region by forming a hairpin (29). In dermatosparactic skin, several proteoglycan filaments are not closely
bound to collagen fibrils but are floating rather freely in
the interfibrillar spaces (54). Thus, dermatosparactic animals (by blocking access to decorin) and decorin null animals (by lacking decorin) could share a common pathogenetic mechanism, i.e., the obstruction/deficiency of a key
regulatory molecule in collagen fibrillogenesis. Further
support for this concept is provided by the fact that the
dermatan sulfate/hydroxyproline ratio in dermatosparactic skin is reduced from 4.7 to 3.6 (38). Since the surface of
the abnormal hieroglyphic collagen is two to four times
higher than that of cylindrical fibrils of normal skin, dermatosparactic animals should also have a significant decrease in collagen-associated decorin.
2(V) gene exhibit phenotypic
and ultrastructural features resembling EDS and some aspects of the decorin-deficient phenotype (1). The mutant
animals develop skin fragility with disorganized dermal
fibrils of variable diameter; however, the affected animals
show various degrees of kypho-lordosis and corneal abnormalities, not observed in the decorin null mice. Several independent studies have now confirmed the linkage of
1(V) collagen gene to EDS I and II (11, 37, 63). It is
known that type V collagen localizes at the periphery of
the coarser collagen fibrils, composed predominantly of
type I, and it has been proposed that the interaction between collagen types I and V regulated fibril diameter (7).
Thus, genetic defects in type V collagen could explain the
observation that collagen fibrils in skin of EDS I and II patients are larger than normal and irregular in outline.
These observations indicate that "modifiers" of collagen fibrillogenesis, such as type V collagen and decorin, might
have profound biological consequences in vivo.
Conclusions
Address all correspondence to Renato V. Iozzo, Department of Pathology, Anatomy, and Cell Biology, Thomas Jefferson University, 1020 Locust Street, Room 249, Jefferson Alumni Hall, Philadelphia, PA 19107. Tel.: (215) 503-2208. Fax: (215) 923-7969. e-mail: iozzo{at}lac.jci.tju.edu
Received for publication 26 September 1996 and in revised form 8 November 1996.
K.G. Danielson and H. Baribault contributed equally to this work.We thank T. Scholzen for help in the initial stages of this work; C. Clark and R. Trelstad for their critical reading of the manuscript; M. Mathiak, I. Eichstetter, and B. Tuma for valuable assistance; S. Kalidindi for help with biomechanical measurements; M. Pisano for valuable discussion and advice; L. Fisher for the generous gift of antisera; K. Soderberg for ES cell work; J. Avis for embryo manipulation; M. Verloop and R. Jenkins for animal procedures; M. Wilson-Heiner for genotyping; T. Doetschman for the gift of the neomycin-resistant mouse colony; A. Nagy for the R1 cells; and the Genetics Institute for the gift of CHO-LIF cell line.
This work was supported by National Institutes of Health grants RO1 CA39481 and RO1 CA47282 (R.V. Iozzo), AR41816 and CA30199 (H. Baribault), and by Wellcome Trust 019512 and the Medical Research Council ICS-95-14 (K.E. Kadler).
EDS, Ehlers-Danlos syndrome; ES, embryonic stem; SLRP, small leucine-rich proteoglycan; STEM, scanning transmission EM.