Knowledge of cartilage hydration (water uptake) is essential to understanding and predicting its load bearing and lubrication properties since it is primarily the osmotic properties of cartilage that cause it to resist applied compressive loads. Cartilage swelling behavior is also exquisitely sensitive to changes occurring in development, disease, degeneration, and aging. Moreover, controlled hydration or swelling of cartilage also provides a means of determining functional properties of cartilage and of other extracellular matrices. One concept that we introduced previously was the controlled hydration of cartilage could also be used to measure important physical/chemical properties of the collagen network and proteoglycans (PG) in cartilage independently within the extracellular matrix. This approach entailed modeling the cartilage tissue matrix as a composite material consisting of two distinct phases: a collagen network and a proteoglycan solution trapped within it; applying various known levels of equilibrium osmotic stress; and using physical-chemical principles and additional experiments to determine a """"""""pressure-volume"""""""" relationship for both the PG and collagen phases independently. In pilot studies, we used this approach to determine pressure-volume curves for the collagen network and the PG phases in native and in trypsin treated normal human cartilage specimen, as well as in cartilage specimen from osteoarthritic (OA) joints. In both normal and trypsin-treated specimen, collagen network stiffness appeared unchanged, whereas in the OA specimen, collagen network stiffness decreased. Our findings highlighted the role of the collagen network in limiting normal cartilage hydration, and in ensuring a high PG concentration in the matrix, both of which are essential for effective load bearing in cartilage and lubrication, but are lost in OA. These data also suggest that the loss of collagen network stiffness, and not the loss or modification of PGs may be the incipient event leading to the subsequent disintegration of cartilage observed in OA. One shortcoming of this approach, however, was that it required many person-days to study a single cartilage specimen, so the approach was not suitable to routine pathological analysis or use in tissue engineering applications. Secondly, a significant amount of the tissue specimen was required to obtain the osmotic titration curves. This contributed to long equilibration times in obtaining data as well as large amounts of sample, which are not routinely available in most clinical and biological applications. Therefore, we have recently developed a new micro-osmometer to perform these experiments in a practical and rapid manner. This instrument can measure minute amounts of water absorbed by small tissue samples (<1 microgram) as a function of the equilibrium activity (pressure) of the surrounding water vapor. A quartz crystal detects the water uptake of a specimen attached to its surface. The high sensitivity of its resonance frequency to small changes in the amount of adsorbed water allows us to measure the mass uptake of the tissue specimen precisely. Varying the equilibrium vapor pressure surrounding the specimen induces controlled changes in the osmotic pressure of the tissue layer. To illustrate the applicability of the new apparatus, we recently measured the swelling pressure of tissue-engineered cartilage specimen. The micro-osmometer will eventually permit us to obtain a profile of the osmotic compressibility or stiffness of multiple cartilage specimens simultaneously as a function of depth from the articular surface to the bone interface. It will also allow us to quantify the contributions of individual components of ECM (such as aggrecan and hyaluronic acid) to the total osmotic pressure. Moreover, it should allow us to assess the osmotic compatibility and mechanical integrity of developing tissues and of tissue-engineered cartilage (or ECM) for implantation.
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