Comparative equilibrium mechanical properties of bovine and lamprey cartilaginous tissues
1 Department of Biomedical Sciences, UPEI Atlantic Veterinary College,
Charlottetown, Prince Edward Island, C1A 4P3, Canada
2 Department of Mathematics, Lafayette College, Easton, PA 18042,
USA
3 Department of Biology, St Francis Xavier University, Antigonish, Nova
Scotia, B2G 2W5, Canada
* Author for correspondence (e-mail: edemont{at}stfx.ca)
Accepted 27 January 2003
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Summary |
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Key words: cartilage, collagen, tension, compression, lamprey, Petromyzon marinus, stiffness, viscoelastic properties
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Introduction |
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Biochemical characterizations have greatly enhanced our morphological
understanding of lamprey cartilages. The resistance of annular cartilage to
CNBr and NaOH digestion confirmed that the major structural protein of lamprey
cartilages is not composed of collagen. Instead, a unique insoluble matrix
protein, named lamprin, was found to constitute 44-51% of the dry mass of the
annular cartilage (Wright et al.,
1983; Robson et al.,
1993
). In the case of lamprey pericardial cartilage, amino acid
composition analysis indicates significantly higher levels of hydroxyproline
compared with annular cartilage (Robson et
al., 1997
). Morphological, biochemical and molecular biological
analysis has demonstrated that the major matrix protein(s) of pericardial
cartilage is not lamprin but yet another noncollagenous, elastin-like protein,
despite it also being resistant to CNBr/NaOH digestion
(Wright et al., 2001
).
Taken together, previously established research and recent biochemical studies indicate that the skeleton of the sea lamprey consists of a family of related, but non-identical, cartilages characterized by non-collagenous, elastin-like matrix proteins. Currently, we know nothing of the innate mechanical properties of lamprey cartilages nor of how their mechanical properties compare with those of mammalian cartilages. Consequently, the lamprey skeleton presents itself as a unique model from which we may broaden the current understanding of cartilage structurefunction behavior. In conducting this investigation, we addressed the following questions: (1) do lamprey annular and pericardial cartilages exhibit a viscoelastic response to static mechanical loading; (2) do lamprey annular and pericardial cartilages demonstrate different equilibrium mechanical stiffnesses; (3) do adult lamprey cartilages stiffen with age and (4) do lamprey annular and pericardial cartilages differ from mammalian cartilages in their equilibrium mechanical stiffness? To answer these questions, we employed a series of equilibrium mechanical tests and morphological characterizations on two lamprey cartilages and one non-hyaline mammalian cartilage.
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Materials and methods |
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Morphology and histochemistry
Water content was determined for young adult annular (N=8), young
adult pericardial (N=8), aged annular (N=8), aged
pericardial (N=5) and bovine auricular (N=8) cartilages.
First, cartilage samples were thawed and cut as if for mechanical testing.
Specimens were then equilibrated in lamprey or lactated Ringer's solution for
20 min, lightly blotted with Kimwipe® tissue paper and weighed on an
analytical balance (accuracy 0.001 g). Finally, samples were dried in an oven
at 80°C for 48 h. The wet mass minus the dry mass yielded the water
content of the cartilage.
Annular (5 young and 5 aged each) and pericardial (5 young and 5 aged each)
cartilage samples were taken from each of the two lamprey sample batches
(before and after testing) in order to examine tissue morphology. Four bovine
auricular cartilage samples were also taken from four separate animals for
morphological analysis. Cartilage samples were fixed in either 80% ethanol for
assessment of calcium deposition (Presnell
and Schreibman, 1997) or 10% neutral buffered formalin to assess
any disruption of tissue morphology resulting from the mechanical tests. All
fixed tissues were then dehydrated through an ascending ethanol series,
cleared in xylene and embedded in paraffin. Sections, 5-6 µm thick, were
cut and mounted on glass slides previously coated with a 2%
silaneacetone solution. For calcium assessment, the sections were
stained using alizarin red S and alizarin red S after a 0.5 mol l-1
EDTA soak. Sections of formalin-fixed bovine muscle (calf skeletal muscle)
containing a calcified lesion (white muscle disease) served as positive
controls for these histochemical stains, and replicate sections treated with
0.5 mol l-1 EDTA prior to staining served as negative controls.
Sections of the formalin-fixed and ethanol-fixed lamprey and bovine cartilages
were also stained according to Harris' hematoxylin and eosin procedure
(Prophet et al., 1992
).
Measurements of tissue structures at the light microscopic level were made
using a calibrated ocular scale.
Isolated annular (aged; N=5), pericardial (aged; N=5), and bovine auricular cartilages (N=2) were prepared for high-resolution light microscopy and transmission electron microscopical analysis. Individual cartilage pieces were fixed in 2% glutaraldehyde in 0.1 mol l-1 phosphate buffer, pH 7.3-7.5 for 24 h at 4°C and post fixed (1% OsO4 for 1 h at room temperature). Samples were then washed (0.1 mol l-1 phosphate buffer), dehydrated through an ethanol series and embedded in Spurr's resin. Thick sections (1 µm) were cut and stained with 1% toluidine blue in 1% sodium borate solution. Ultrathin sections (85-100 nm) were stained with uranyl acetate and Sato's lead stain before being examined using Hitachi H-600 and H-7500 electron microscopes operated at 75 kV and 80 kV, respectively.
Stree-relaxation equipment and sample preparation
Stress-relaxation experiments were conducted using a custom-built uniaxial
lever system (Fig. 2) that
accommodated two sets of adjustable, custom-engineered gripping devices. For
tensile experiments, two stainless steel clamps with beveled ridges and metal
spikes were created to hold cartilage samples firmly at both ends. Once
clamped, the cartilage was trimmed to match the size of the clamp heads,
gripping approximately 18 mm2 of cartilage. Between the clamps,
specimens were trimmed to yield a thin gauge region with relatively constant
dimensions. For compression experiments, cartilage samples were held between
two smooth, impermeable stainless steel plates. Clamped specimens were encased
in a transparent bathing chamber filled with the appropriate Ringer's
solution. Bovine cartilage runs were performed at room temperature
(27±3°C), while lamprey cartilage runs were performed at
10±2°C. Before each experiment, the height of the top clamp/plate
was adjusted to the position where the sample was just straight. In this
position, the cartilage experienced negligible strain and an initial tare load
of 0.1-0.2 mV (0.003-0.006 N). This initial `pre-stress' was applied to ensure
total contact over the specimen surface and to minimize the effects of surface
irregularities.
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Analysis of cartilage sample dimensions was made possible through the use of a CCD high-resolution video camera (Sanyo, Inc., Concord, ON, Canada; 2x magnification), which was manually positioned to take separate images from the front (length, width) and side (thickness) of each sample. For all acquired images, the camera was oriented 20 mm from the bathing chamber and perpendicular to the sample's axis of loading (Fig. 2). The length, width and thickness of each specimen were measured at five equally spaced positions along each dimension. Measurements of specimen length, width and thickness were manually computed using standard pixel analysis tools in LabView® (v.5.0, National Instruments, Inc., Austin, TX, USA). Using a variety of calibration objects, the measurement accuracy of this photo-digitizing method was estimated to be within 4%. All samples of a given cartilage type were of uniform shape and similar in size (Table 1).
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Stress-relaxation experiments
The experiments conducted were dependent upon the morphology of the
cartilage tissues and the ability of the testing machinery to clamp specimens
adequately. With these considerations in mind, uniaxial tensile tests were
performed on lamprey pericardial cartilage samples (with attached
perichondria) sectioned from the animal's dorsal plane and oriented laterally
around the tissue's caudal apex (Fig.
3A). In this fashion, the loading axis was parallel to the plane
of the perichondria. Uniaxial compressive tests were performed on
dorso-ventrally oriented samples (without perichondria) taken from the
anterior-most portion of the lamprey annular cartilage ring
(Fig. 3B). For bovine auricular
cartilage, uniaxial tensile tests were performed on lengthwise samples
(distal-proximal to the head) without perichondria, and uniaxial compressive
tests were performed on dorso-ventrally oriented samples without perichondria
(Fig. 3C,D).
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All stress-relaxation experiments consisted of unconfined uniaxial
deformations enabling the quantification of equilibrium stresses after fluid
loss and shape changes along the tissue's longitudinal and lateral dimensions.
Prior to each stress-relaxation run, cartilage samples were allowed to
equilibrate for 20-30 min in the bathing chamber. Then, a multifunction
synthesizer (8904A DC-600 MHz, Hewlett Packard, Inc., Palo Alto, CA, USA)
directed the motorized lever (Model 300 Servo System Lever; Cambridge
Technology, Inc., Watertown, MA, USA) to impose near instantaneous (<1 s)
fixed deformations (fixed strains) on the cartilage samples. Each sample's
resistance to deformation was interpreted as a voltage output, which was read
by a digitizing oscilloscope (54501A 100MHz; Hewlett Packard, Inc.).
LabView® (v.5.0) was used to automate the entire process, storing voltage
data every 4 s until equilibrium. Equilibrium was initially determined by
visual inspection and subsequently quantified by calculation of the time where
V/
t=0 (V is voltage, t is
time) for ±5 min. The resulting equilibrium voltage required to
maintain the applied strain deformation in tension (or compression) was
recorded and this process repeated 3-5 times, allowing recovery of the
cartilage between each trial, defined as each sample's return to its initial
shape and initial pre-stress load.
Stress-strain analysis
According to isolation, the dominant dimension was considered the length
for each cartilage sample, and tensile/compressive deformations were imposed
along this longitudinal axis. The resting length (L0) and
resting stress (0) of each cartilage sample's gauge region
were measured prior to testing. In addition, the width and thickness of each
gauge region were measured at five equidistant points along the length of the
sample after each instance of deformation. From these values, the mean
thickness (th) and width (w) were calculated, the product of
which was the mean cross-sectional area (A) for the cartilage sample.
After deformation, the extension or compression of each sample
L was calculated using:
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Statistical analysis
Determination of cartilage modulus values was performed using a standard
least-squares regression fit to our experimentally determined stress-strain
data, and in this manner all stress-strain plots yielded regression fits with
r2 values of 0.80. To evaluate differences in mean
modulus values and mean water contents of different cartilages, single-factor
analysis of variance (ANOVA) was performed after confirming the normality of
residuals and using Minitab® statistical software (v.12, Minitab, Inc.,
State College, PA, USA). If an ANOVA indicated the presence of a significant
difference, Tukey multiple comparison tests were performed to determine
specific mean differences. The abovementioned statistical tests were all
performed with an upper limit of significance at P=0.05.
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Results |
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While extensive light microscopical analyses of lamprey annular and
pericardial cartilages have been reported
(Wright and Youson, 1983;
Wright et al., 1988
;
Robson et al., 1997
), several
previously unpublished structural features were observed during this
investigation. The perichondria of the pericardial cartilage (young and aged)
give boundaries, as the cartilage forms the pericardium enclosing the heart
and supports it within the body cavity
(Fig. 1). The outer
perichondrium (exposed to the body cavity) was found to be thin (
20-40
µm) with densely packed collagen fibers. The inner perichondrium (lining
the pericardial cavity) is 2-4 times as thick (
80-120 µm) as the outer
perichondrium, largely due to the presence of what appears to be a looser
arrangement of collagen fibers containing large adipose cells and blood
vessels (Fig. 4A). The central
positioning of these cells and blood vessels allows for a delineation of the
inner perichondrium into two layers, one contacting the cartilage and the
other contacting the pericardial cavity. In annular cartilage (young or aged),
there is no obvious boundary for perichondria. Instead, the perichondria tend
to blend with a variety of adjacent connecting tissues
(Fig. 4B). Depending on
location, the perichondria can be seen merging with overlying layers of muscle
and even adipose tissue. The dense collagenous perichondria of bovine
auricular cartilage were similar to that of the annular cartilage in that they
blend with adjacent connective tissues
(Fig. 4C).
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Hematoxylin/eosin staining gave identical results for young adult and aged lamprey cartilages, although staining differences between the two lamprey cartilage types were observed. `Hard' annular cartilage ECM and perichondria were found to stain bright pink/red (acidophilic) with eosin (Fig. 4B). Bovine auricular cartilage ECMs and perichondria also stained pink/red with eosin (Fig. 4C) although, in the case of the ECM, eosin staining was mainly associated with elastic fibers while the remainder of the ECM was found to stain a light gray/blue (weakly basophilic). The `soft' pericardial cartilage ECM demonstrated a staining pattern opposite to that of annular cartilage, as evidenced by blue staining (basophilic) with hematoxylin, but the outer and inner perichondria stained pink/red (Fig. 4A). A slight difference in subperichondrial ECM concentration was noted between the outer and inner perichondria of lamprey pericardial cartilages. Specifically, ECM adjacent to the outer perichondrium appeared to form thicker seams between smaller chondrocytes than in the inner perichondrium, where thin seams of ECM surrounded large chondrocytes.
Alizarin red S stains were used for histological assessment of calcium deposition at three pH levels. The positive control gave light, moderate and high intensity red staining within a known calcified lesion for pH levels of 9, 6.4 and 4.2, respectively. Alizarin red staining at a pH of 4.2 gave the most vibrant stain colorations in cartilage samples. The ECM and cellular matrix (CM) regions of all pericardial cartilages exhibited a strong orange/red tint, while all annular cartilage ECMs gave a strong yellow coloration. Perichondria of all cartilages demonstrated a proportional increase in color intensity (as compared with staining of the pure cartilage tissue) and appeared dark red and/or orange. Replicate sections of young and aged lamprey annular and pericardial cartilage pre-treated with 0.5 mol l-1 EDTA were all found to stain a pale red with alizarin red pH 4.2. Thus, the high intensity of staining, particularly in the perichondria of pericardial and annular cartilages, was not apparent for staining after pretreatment with EDTA. Bovine auricular cartilage resembled the EDTA-pre-treated negative control by staining pale red with alizarin red pH 4.2.
Analysis of aged and young cartilage ultrastructure gave no evidence of
mineralization. The only notable difference between aged and young lamprey
cartilages was an apparent degeneration of aged pericardial cartilage outer
perichondria, which were found lacking their epithelial cell layer. The
epithelial cell layers of inner perichondria appeared intact in both young and
aged lamprey pericardial cartilages. Ultrastructural examination of lamprey
cartilages and bovine auricular cartilages showed no other differences from
previous publications on these tissues
(Wright and Youson, 1983;
Serafini-Fracassini and Smith,
1974b
), confirming that our mechanical tests did not alter the
structural integrity of the tissues.
Lamprey pericardial cartilage sections cut parallel to the plane of the perichondrium revealed that collagen fibers of both the inner and outer perichondria were not aligned along a specific line of orientation but existed in a staggered array (Fig. 5). Within this array, collagen fibers were rarely oriented parallel to each other. Instead, fibers typically demonstrated inter-fiber angles ranging from 30° to 90°. There were no noticeable differences in perichondrial fiber orientation between inner and outer perichondria or between young and aged cartilages.
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Both young and aged lamprey annular cartilages were found to contain canal-like structures at all tissue depths (Fig. 6). These canals ranged from 80 µm to 600 µm in diameter, and the cartilage immediately surrounding the canals consisted of dense ECM with many flattened chondrocytes. The canals typically contained loose connective tissue with appreciable quantities of an amorphous ground substance, which contained randomly arranged, widely dispersed fibers and blood vessels. Microscopical examination of annular cartilage sections revealed that canals were sufficiently abundant throughout the tissue to render the testing of regions without canals impossible. No canal structures were observed in lamprey pericardial (young or aged) cartilages or bovine auricular cartilages.
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Viscoelasticity
Characteristic water contents (% wet mass) of lamprey and bovine cartilages
are listed in Table 2. All
cartilages were found to possess a water content constituting >60% of the
cartilage's wet mass. The mean water content for bovine auricular cartilage
was found to be significantly lower than those of lamprey annular and
pericardial cartilage (young adult and aged), although this is probably not
physically significant. Lamprey annular and pericardial cartilages gave mean
water contents that were statistically indistinguishable and there were also
no differences in mean water content between young and aged samples of lamprey
annular and pericardial cartilage.
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All tested cartilages demonstrated a time-dependent stress response to
loading (Fig. 7). That is,
there was a characteristic stress-decay behavior and characteristic length of
time (equilibrium time, te) required for cartilage samples
to reach e. Bovine auricular cartilage, gave
te that were identical for both tensile and compressive
loading. There were, however, noticeable differences in te
among the different cartilage tissues
(Table 2). Both young and aged
lamprey annular and pericardial cartilages required a period of at least 120
min to reach equilibrium after the initial strain was applied. Bovine
auricular cartilage differed in its te, requiring only 30
min to reach equilibrium. After reaching te, all cartilage
samples exhibited complete recovery upon removal of the imposed strain
deformations as demonstrated by 100% recovery to both their initial shape and
0. Still, variations in recovery time
(tr) were found (Table
2) and, although semi-quantitative, they afford clear
distinctions. Lamprey pericardial cartilage (both young and aged) required the
shortest tr of all the cartilages (<0.5 min). Bovine
auricular (tension and compression) and lamprey annular (both young and aged)
cartilages gave substantially longer tr values of 30 min
and 120 min, respectively. Note that tr is identical to
te for these two cartilages. However, while bovine
auricular cartilage required 30 min to return to its initial
0, it required <0.5 min to recover to its initial shape;
upon removal of the deformation, the recovery of
0 lagged
behind the near instantaneous recovery of shape.
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Preliminary work on modeling the viscoelastic properties of lamprey and
bovine cartilages has been previously set forth
(Courtland, 2001). In
accordance with this approach, all cartilages studied in this investigation
exhibited non-exponential (non-linear) decay behavior as supported by two
observations. First, there were demonstrable nonlinear variations of the
centered difference approximation to the decay rate
(
n-1
n+1)/(tn+1tn+1)
versus
n for these cartilages. Second, using
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Equilibrium stiffness
In all cartilages, the stress-strain data were best described with linear
regressions (there was no indication of curvilinear behavior through the
tested range of strains; Fig.
8). Mean equilibrium moduli, or equilibrium stiffnesses
(E), are listed in Table
3 and ranged from 0.71 MPa to 4.85 MPa. An ANOVA indicated the
presence of at least one significant difference among the reported mean
E, and a series of Tukey comparison tests yielded a total of nine
significant differences among them. As can be seen from
Table 3, there are clear
differences in mean E between lamprey cartilages. Young pericardial
cartilage possessed a larger mean E than young annular cartilage. In
addition, aged pericardial cartilages demonstrated a significantly larger mean
E in comparison with all annular cartilages. Aged lamprey pericardial
cartilage also had a larger mean E than that of young pericardial
cartilages. However, comparison of aged versus young annular
cartilage demonstrated no significant difference in mean E. There was
no difference between the mean E values of bovine auricular
cartilages tested in tension and compression. In addition, both young and aged
annular cartilages gave mean E that were not significantly different
to the tensile and compressive mean E of bovine auricular cartilages.
The mean E of young adult lamprey pericardial cartilages was also not
significantly different to either of the bovine mean E; however, aged
pericardial cartilage possessed a mean E significantly larger than
that of either bovine group.
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Discussion |
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The te of lamprey annular cartilage was approximately
120 min. This is the longest relaxation time reported for any vertebrate
cartilage; the typical te for human and bovine articular
cartilage is 15 min (Akizuki et al.,
1986; Schmidt et al.,
1990
). Such a long te for lamprey annular
cartilage is suggestive of an extremely low permeability for this tissue, and
there are three factors that may contribute significantly to this behavior:
canal structures, ECM proteoglycans (PGs) and ECM fibrils. The presence of
canals is not unique to lamprey annular cartilage but is a feature common to
`thick' mammalian cartilages such as rib and epiphyseal growth cartilage
(Shingleton et al., 1997
;
Craatz et al., 1999
). In
lamprey annular cartilage, the combination of the canal walls and the
surrounding dense ECM region may serve to decrease permeability of the
cartilage by creating a surface around which drag forces arise. In terms of
PGs, lamprey annular cartilage contains mostly N-acetyl
galactosamine-containing glycosaminoglycans (GAGs;
Wright et al., 1983
). Since
the majority of characterized N-acetyl galactosamine-containing GAGs
are found in chondroitin sulphate, keratin sulphate and hyaluronate PGs
(Serafini-Fracassini and Smith,
1974a
), this would suggest that the PGs of lamprey annular
cartilage are similar to the low conductivity PGs of mammalian articular
cartilages in that they too incorporate a substantial number of alternating
GAG ß1,4 and ß1,3 linkages. This would help to explain the low
permeability of lamprey annular cartilage as a function of increased flow
resistance around PG GAG side chains
(Comper and Zamparo, 1990
;
Comper and Lyons, 1993
). Still,
one must remember that lamprey annular cartilage has a te
that is twice that of rabbit articular cartilage
(Mow et al., 1989
). As an
additional factor, we propose a functional significance for the lamprin-based
ECM fibrils that, at the ultrastructural level, exist as a highly branched
three-dimensional network (Wright et al.,
1983
; Wright and Youson,
1983
) quite unlike the non-branched arrangement of collagen
fibrils in mammalian articular cartilages (see
Oloyede and Broom, 1996
for a
review). The effect of branched fibrils may be to permit a significant surface
area for hindering the rate of fluid flow from annular cartilage. This would
give rise to a lower permeability for this tissue and explain the longer
te as compared with mammalian cartilages. The
tr for annular cartilage was also approximately 120 min,
suggesting that the recovery process, in which extruded water is drawn back
into the tissue, encounters the same drag-induced forces as were experienced
during stress-relaxation, despite the influence of Donnan osmotic forces that
would encourage fluid flow back into the cartilage. Equal resistance to
extrusion and imbibition of water is in line with drag-induced low
permeability possibly arising from canal structures, low conductivity PGs and
branched ECM fibrils.
When considering the te of lamprey pericardial
cartilage, which was also found to be approximately 120 min, the
aforementioned contributions of PGs and ECM fibrils to reduced permeability
may be valid. However, the extent of their contributions is less clear due to
a lack of knowledge regarding the chemistry of pericardial cartilage PGs and
also due to the substantially reduced quantity of ECM fibrils. There is also
reason to believe that other features, such as chondrocytes and adipose cells,
may influence this tissue's viscoelastic properties. The chondrocytes of
lamprey pericardial cartilage occupy a considerably larger volume fraction
than those of the annular cartilage (compare
Fig. 4A and
Fig. 4B) and, as a result, they
account for most of the tissue's total volume. Since lamprey annular and
pericardial cartilages were found to have identical water contents and most of
the pericardial volume exists as chondrocytes, it can be assumed that most of
the bound water in pericardial cartilage is intracellular. Work by Freeman et
al. (1994) demonstrated that
chondrocytes impart a substantial viscoelastic effect to agarose gels that
they were cultured in, and other research has shown that chondrocytes exhibit
viscoelastic behavior common even to whole cartilage (for a review, see
Guilak, 2000
). It is therefore
conceivable that the viscoelastic behavior of pericardial cartilage is largely
a function of chondrocyte viscoelasticity. Given that the
te for pericardial cartilage is 120 min, it seems likely
that this effect would result from chondrocyte fluid exudation. Adipose cells
may also contribute to the viscoelastic characteristics of lamprey pericardial
cartilage, but further investigation is required.
If viscoelastic effects (e.g. drag and fluid redistribution) were truly the
only determinants of lamprey pericardial cartilage's permeability, then a
te of 120 min could not accommodate a
tr <<120 min. Yet, all pericardial cartilages recovered
fully to their initial shape and 0 in <30 s. This
phenomenon can best be explained as a result of testing the pericardial
cartilages with their attached perichondria. The perichondria are composed
mainly of collagen fibers (see Fig.
5). Since the tensile E for pericardial cartilages was
two orders of magnitude smaller than that for pure collagen,
stress-minimization via intrafibrillar contraction of collagen seems
an unlikely determinant of pericardial cartilage's short
tr. A more likely explanation is the potential
crosslinking of perichondrial collagen with the surrounding electron-dense
elastic-like fibers (see Wright et al.,
1988
), which could permit tensile loading-induced fiber
reorientation. If this were true, then release of the tensile load after
equilibrium would permit a rapid minimization of this high-energy
conformation, thereby explaining the observed small tr.
Regardless of the exact mechanism behind tr<30 s, it is
clear that the initial shape and
0 of pericardial cartilages
are determined by the tissue's collagenous perichondria alone, and it follows
that any tr based on these parameters will not necessarily
indicate the true recovery behavior of the tissue's CM and ECM. Thus, the
tr reported here for lamprey pericardial cartilages is
best viewed as an apparent recovery time, one that is partially
independent of and hides the tissue's true viscoelasticity.
Bovine auricular cartilage gave a te of 30 min for both
tensile and compressive deformations. This is substantially shorter than the
te of both lamprey annular and pericardial cartilages,
suggesting that bovine auricular cartilage has a comparatively larger
permeability. Possible explanations for this behavior are the presence of more
PGs, a higher percentage of high-conductivity PGs and a less-connected
arrangement of structural proteins in bovine auricular cartilage. The
tr of bovine auricular cartilage for both tension and
compression was found to be identical to the te
(approximately 30 min), suggesting a certain degree of structural homogeneity
for this tissue. In addition, this suggests that in auricular cartilage, the
0 load is borne not by the elastic or collagenous structural
fibers but by the cartilage's Donnan osmotic pressure, the recovery of which
requires a time period equivalent to that of the stress-relaxation experiment
(about 30 min). It must be noted, though, that auricular cartilage's recovery
to its initial shape required a time substantially less than 30 min (<0.5
min) and this is probably due to recoil of elastic fibers and/or an overall
reorientation of the fibrous network. We must then define the true
tr for bovine auricular cartilage as one indicated not by the
recovery of shape alone but by the recovery of
0
and shape. Given the differential recovery of both lamprey
pericardial and bovine auricular cartilages, it seems that assessing the
actual recovery of loaded cartilages can be quite difficult.
The above discussion has suggested that lamprey cartilages have markedly
lower hydraulic permeabilities than mammalian cartilages and, if so, this must
be related to the mechanical function of these tissues. In lamprey pericardial
cartilage, the role of low permeability is probably to prevent relaxation of
the tissue during short intervals of loading. Since the cartilage is very thin
and the majority of fluid originates from its chondrocytes and/or adipose
cells, even a small relaxation of these components would decrease cartilage
Donnan osmotic pressure and the cartilage as a whole would be less flexible
and less resilient. That is, the pericardial cartilage would be virtually
incapable of absorbing/releasing any expansion/recoil forces imparted to it by
the heart and branchial basket. In lamprey annular cartilage, the low
permeability is more likely to be a mechanism for the dissipation of energy
over longer intervals of time (i.e. feeding). The highly branched lamprin
fibril network probably affords a greater surface area (per fibril volume)
over which forces are distributed, thus minimizing the load borne by
individual fibrils and transferred through the aqueous phase of the ECM. It is
important to note that while lamprey cartilages appear to differ from
mammalian cartilages in their permeabilities, they each exhibit a non-linear
stress-decay behavior that is also characteristic of mammalian articular
cartilages (Mansour and Mow,
1976; for a review, see
Maroudas, 1979
).
Stiffnesses
For the intrinsic, static parameter E, charge effects are normally
the major contributing factors in the ability of cartilage to resist
compressive deformations. A list of compressive E can be found in
Table 4 and from these it is
clear that the compressive E of young adult lamprey annular cartilage
is nearly identical to the compressive E reported for bovine
articular cartilage (hyaline cartilage) as well as bovine meniscus
(fibrocartilage). The reported E for lamprey annular cartilage is
somewhat smaller than for human articular cartilage and bovine auricular
cartilage but is nonetheless within one order of magnitude of both. Possible
determinants of the similarities in compressive stiffness between lamprey
annular cartilage and mammalian cartilages include the incorporation of
similarly charged PGs and similar ionic contents. In addition, the lamprin ECM
fibrils may contribute to the tissue's compressive stiffness through extensive
fibril interactions (i.e. flow resistance).
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When considering the ability of connective tissues to resist tensile
forces, structural proteins are usually of fundamental importance. The fact
that lamprey pericardial cartilage easily disintegrates in the absence of its
perichondria suggests that the pure cartilage has a very low tensile
E compared with the intact tissue with both perichondria attached.
The ECM fibrils, scarce in comparison to the tissue's fraction of
chondrocytes, must therefore play only a minor role in the tensile E
of the cartilage. Consequently, in order to resist loads while at the same
time preserving the viscoelastic properties of the tissue, the pericardial
cartilage employs perichondrial layers composed of a dense network of collagen
fibrils. This gives rise to a tensile E that is significantly larger
than the compressive E of lamprey annular cartilage. A listing of
reported tensile E can be found in
Table 5 and shows that young
adult lamprey pericardial cartilage is comparable with mammalian cartilages in
its tensile E but clearly falls in the low end of reported cartilage
E. Since research has also shown that the number of hydroxypyridinium
cross-links within collagen fibrils directly influences the tensile E
and strength of the intact cartilage
(Schmidt et al., 1987), the
relatively low volume fraction content of collagen fibers in the perichondria
(Fig. 4A) and the non-parallel
orientation of these fibers (Fig.
5) may be particularly relevant to the lower E of lamprey
pericardial cartilage.
|
The fact that both young pericardial and bovine auricular cartilages give
comparable tensile E suggests that they may share common features.
Indeed, both cartilages have a low density of collagen fibers/fibrils when
compared with hyaline and fibrocartilages, and both contain ECM fibrils
demonstrating biochemical similarities to elastin. Still, in pericardial
cartilage, the collagen fibers are fairly well organized in the perichondria
but are a small fraction of the total tissue volume. When compared with
auricular cartilage, where collagen fibrils constitute up to 80% of the
fibrous content (Stockwell,
1979) and are distributed throughout the tissue, the similarity in
tensile E of these two cartilages is startling. Of course, with such
a low sample size for bovine auricular cartilage, the mean E may not
be truly representative of a population, but, assuming the similarity is real,
lamprey pericardial cartilage may be circumventing a dearth of collagen by
incorporating a select number of fibers at high density in critical regions
(perichondria), by orienting fibers in an optimal way or by developing
more-extensive fibril cross-links within fiber fibrils. Further biochemical
analyses are necessary to validate these possibilities.
Similar to the classification reported here for lamprey, the cartilaginous
elements of the hagfish have been characterized as either hard or soft
structures (Cole, 1905). Hard
hagfish cartilages such as the nasal and auditory capsule were found to have
an ECM that stained orange/red with eosin, while soft cartilages such as the
branchial arches had an ECM that stained blue with hematoxylin
(Robson et al., 2000
). Results
from the present investigation demonstrated that hematoxylin/eosin staining of
lamprey cartilages gave an identical hardsoft staining pattern. Thus,
while mammalian cartilage ECMs (bovine auricular cartilage) typically
displayed both hematoxylin and eosin staining
(Fig. 4C), the cartilages from
jawless craniates stain differently, suggesting that variation at the
molecular level gives rise to differentiation at the structural level. It
turns out that an underlying similarity in ECM fibril primary structure can be
associated with hard and soft cartilage groups. Acidophilic hard cartilages
(orange/red stain) contain a larger proportion of basic amino acids, while
basophilic soft cartilages (blue stain) contain a larger proportion of acidic
amino acids. For example, hagfish contain more basic amino acids (e.g.
histidine and tyrosine) in their hard cartilages than in their soft cartilages
and this is also true for hard and soft lamprey cartilages (Robson et al.,
1997
,
2000
).
Although lamprey annular cartilage was not found to stiffen with age,
lamprey pericardial cartilage clearly does. The mean E of aged
pericardial cartilage was found to be statistically larger than the mean
E of young and aged annular cartilage, young pericardial cartilage,
as well as bovine auricular cartilage tested in tension and compression. Since
lamprey cartilage has demonstrated the ability to calcify in vitro
(Langille and Hall, 1993), the
`crisp' consistency of aged lamprey pericardial cartilage suggested a possible
deposition of calcium within the cartilage ECM, within the chondrocytes or,
possibly, within the inter-fibrillar spaces of perichondrial collagen fibers.
However, staining with alizarin red was inconclusive; young and aged
pericardial cartilages gave staining patterns that were virtually identical
(both with and without EDTA treatment). In addition, examination of lamprey
pericardial cartilage perichondria via transmission electron
microscopy also failed to uncover evidence of mineralization, as no
electron-dense aggregations were found within collagen inter- or
intrafibrillar spaces of perichondria. Thus, while we cannot completely rule
out the possibility of mineralization in aged pericardial cartilage (i.e. free
ion mineralization), we cannot substantiate its contribution to an increase in
the tissue's stiffness. Nevertheless, it does seem likely that the
determinant(s) of increased stiffness is in some way associated with the
condition of the perichondrial tissue. Electron microscopical observations
revealed degeneration in the mesothelium of the aged pericardial cartilage's
outer perichondrium and this is probably linked to the age-associated
degradation of the liver, which in turn results in a large portion of
metabolic waste being released to the surrounding environment (i.e. the
pericardial cartilage's outer perichondrium).
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Conclusions |
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Acknowledgments |
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References |
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Aitken, G. K., Bourne, R. B., Finlay, J. B., Rorabeck, C. H. and Andreae, P. R. (1985). Indentation stiffness of the cancellous bone in the distal human tibia. Clin. Orthop. 201,264 -270.[Medline]
Akizuki, S., Mow, V. C., Muller, F., Pita, J. C., Howell, D. S. and Manicourt, D. H. (1986). Tensile properties of human knee joint cartilage. I. Influence of ionic conditions, weight bearing, and fibrillation on the tensile modulus. J. Orthop. Res. 4, 379-392.[Medline]
Cole, F. J. (1905). A monograph on the general morphology of the myxinoid fishes, based on a study of Myxine. 1. The anatomy of the skeleton. Trans. R. Soc. Edinburgh 41,749 -791.
Comper, W. D. and Lyons, K. C. (1993). Non-electrostatic factors govern the hydrodynamic properties of articular cartilage proteoglycan. Biochem. J. 289,543 -547.[Medline]
Comper, W. D. and Zamparo, O. (1990). Hydrodynamic properties of connective-tissue polysaccharides. Biochem. J. 269,561 -564.[Medline]
Courtland, H.-W. (2001). Equilibrium mechanical properties of two noncollagenous cartilages in the sea lamprey Petromyzon marinus. M.Sc. Thesis. University of Prince Edward Island, Charlottetown, PE, Canada.
Craatz, S., Weiss, J. and Schmidt, W. (1999). Histologic-histochemical and immunocytochemical investigations of cartilage canals in human rib cartilage. Anat. Anz. 181,359 -363.[Medline]
Dimery, N. J., Alexander, R. McN. and Deyst, K. A. (1985). Mechanics of the ligamentum nuchae of some artiodactyls. J. Zool. 206A,341 -351.
Freeman, P. M., Natarjan, R. N., Kimura, J. H. and Andriacchi, T. P. (1994). Chondrocyte cells respond mechanically to compressive loads. J. Orthop. Res. 12,311 -320.[Medline]
Guilak, F. (2000). The deformation behavior and viscoelastic properties of chondrocytes in articular cartilage. Biorheology 37,27 -44.[Medline]
Hamaide, A., Arnoczky, S. P., Ciarelli, M. J. and Gardner, K. (1998). Effects of age and location on the biomechanical and biochemical properties of canine tracheal ring cartilage in dogs. Am. J. Vet. Res. 59,18 -22.[Medline]
Hubbs, C. L. and Potter, I. C. (1971). Distribution, phylogeny and taxonomy. In The Biology of Lampreys, vol. 1 (ed. M. W. Hardisty and I. C. Potter), pp. 1-5. London: Academic Press.
Jurvelin, J. S., Buschmann, M. D. and Hunziker, E. B. (1997). Optical and mechanical determination of Poisson's ratio of adult bovine humeral articular cartilage. J. Biomech. 30,235 -241.[CrossRef][Medline]
Langille, R. M. and Hall, B. K. (1993). Calcification of cartilage from the lamprey Petromyzon marinus (L.) in vitro. Acta. Zool. 74, 31-41.
Mansour, J. M. and Mow, V. C. (1976). The permeability of articular cartilage under compressive strain and at high pressures. J. Bone Joint Surg. 58,509 -516.[Abstract]
Maroudas, A. (1979). Physicochemical properties of articular cartilage. In Adult Articular Cartilage, 2nd edition (ed. M. A. R. Freeman), pp. 215-290. Turnbridge Wells, UK: Pitman Medical.
Mow, V. C., Proctor, C. S. and Kelly, M. A. (1989). Biomechanics of articular cartilage. In Cartilage: Biomechanics of the Musculoskeletal System, 2nd edition (ed. M. Nordin and V. H. Frankel), pp.31 -58. Philadelphia: Lea and Febiger.
Mow, V. C., Kuei, S. C., Lai, W. M. and Armstrong, C. G. (1980). Biphasic creep and stress relaxation of articular cartilage in compression? Theory and experiments. J. Biomech. Eng. 102,73 -84.[Medline]
Oloyede, A. and Broom, N. (1996). The biomechanics of cartilage load-carriage. Conn. Tiss. Res. 34,119 -143.
Presnell, J. K. and Schreibman, M. P. (1997). Staining pigments and minerals. In Humason's Animal Tissue Techniques, 5th edition (ed. J. K. Presnell and M. P. Schreibman), pp. 223-224. Baltimore: Johns Hopkins University Press.
Proctor, C. S., Schmidt, R. R., Whipple, R. R., Kelly, M. A. and Mow, V. C. (1989). Material properties of the normal medial bovine meniscus. J. Orthop. Res. 7, 771-782.[Medline]
Prophet, E. B., Mills, B., Arrington, J. B. and Sobin, L. H. (ed.) (1992). Armed Forces Institute of Pathology: Laboratory Methods in Histotechnology. Washington, DC: American Registry of Pathology.
Quinn, P., Barros, C. and Whittingham, D. G. (1982). Preservation of hamster oocytes to assay the fertilization capacity of human spermatozoa. J. Reprod. Fertil. 66,161 -168.[Abstract]
Rains, J. K., Bert, J. L., Roberts, C. R. and Pare, P. D.
(1992). Mechanical properties of human tracheal cartilage.
J. Appl. Physiol. 72,219
-225.
Roberts, C. R., Rains, J. K., Paré, P. D., Walker, D. C., Wiggs, B. and Bert, J. L. (1998). Ultrastructure and tensile properties of human tracheal cartilage. J. Biomech. 31, 81-86.[Medline]
Robson, P., Wright, G. M. and Keeley, F. W. (2000). Distinct non-collagen based cartilages comprising the endoskeleton of the Atlantic hagfish, Myxine glutinosa. Anat. Embryol. 202,281 -290.[CrossRef][Medline]
Robson, P., Wright, G. M., Youson, J. H. and Keeley, F. W. (1997). A family of non-collagen-based cartilages in the skeleton of the sea lamprey, Petromyzon marinus. Comp. Biochem. Physiol. B 118,71 -78.[CrossRef]
Robson, P., Wright, G. M., Sitarz, E., Maiti, A., Rawat, M.,
Youson, J. H. and Keeley, F. W. (1993). Characterization of
lamprin, an unusual matrix protein from lamprey cartilage. J. Biol.
Chem. 268,1440
-1447.
Rovainen, C. M. (1973). Projections of individual axons in lamprey spinal cord determined by tracings through serial sections. J. Comp. Neurol. 149,193 -201.[Medline]
Schmidt, M. B., Schoonbeck, J. M., Mow, V. C., Eyre, D. R. and Chun, L. E. (1987). The relationship between collagen crosslinking and the tensile properties of articular cartilage. Trans. Orthop. Res. Soc. 12, 134.
Schmidt, M. B., Mow, V. C., Chun, L. E. and Eyre, D. R. (1990). Effects of proteoglycan extraction on the tensile behavior of articular cartilage. J. Orthop. Res. 8, 353-363.[Medline]
Serafini-Fracassini, A. and Smith, J. W. (1974a). Glycosaminoglycans and proteoglycans. In The Structure and Biochemistry of Cartilage (ed. A. Serafini-Fracassini and J. W. Smith), pp. 64-112. London: Churchill Livingstone.
Serafini-Fracassini, A. and Smith, J. W. (1974b). Elastic cartilage. In The Structure and Biochemistry of Cartilage (ed. A. Serafini-Fracassini and J. W. Smith), pp. 220-228. London: Churchill Livingstone.
Shingleton, W. D., Mackie, E. J., Cawston, T. E. and Jeffcott, L. B. (1997). Cartilage canals in equine articular/epiphyseal growth cartilage and a possible association with dyschondroplasia. Equine Vet. J. 29,360 -364.[Medline]
Stockwell, R. A. (1979). Biology of Cartilage Cells. Cambridge: Cambridge University Press.
Vogel, S. (1988). A matter of materials. In Life's Devices (ed. S. Vogel), pp.177 -200. Princeton: Princeton University Press.
Wong, M., Ponticiello, M., Kovanen, V. and Jurvelin, J. S. (2000). Volumetric changes of articular cartilage during stress relaxation in unconfined compression. J. Biomech. 33,1049 -1054.[CrossRef][Medline]
Wright, G. M. and Youson, J. H. (1983). Ultrastructure of cartilage from young adult sea lamprey, Petromyzon marinus L: a new type of vertebrate cartilage. Am. J. Anat. 167,59 -70.[Medline]
Wright, G. M., Keeley, F. W. and Robson, P. (2001). The unusual cartilaginous tissues of jawless craniates, cephalochordates and invertebrates. Cell Tissue. Res. 304,165 -174.[CrossRef][Medline]
Wright, G. M., Keeley, F. W. and Youson, J. H. (1983). Lamprin: a new vertebrate protein comprising the major structural protein of adult lamprey cartilage. Experientia 39,495 -497.
Wright, G. M., Armstrong, L. A., Jacques, A. M. and Youson, J. H. (1988). Trabecular, nasal, branchial, and pericardial cartilages in the sea lamprey, Petromyzon marinus: fine structure and immunohistochemical detection of elastin. Am. J. Anat. 182, 1-15.[Medline]
Yao, J. Q. and Seedhom, B. B. (1993). Mechanical conditioning of articular cartilage to prevalent stresses. Brit. J. Rheum. 32,956 -965.