Polarized Light Microscopy Study - CiteSeerX

Connective Tissue Research, 48:76–84, 2007 c Informa Healthcare Copyright ISSN: 0300-8207 print / 1521-0456 online DOI: 10.1080/03008200601130950

Morphological Changes in Articular Cartilage Due to Static Compression: Polarized Light Microscopy Study Hisham A. Alhadlaq Department of Physics and Center for Biomedical Research, Oakland University, Rochester, Michigan, USA Department of Physics and Astronomy, King Saud University, Riyadh, Saudi Arabia

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Yang Xia Department of Physics and Center for Biomedical Research, Oakland University, Rochester, Michigan, USA

Fay M. Hansen Department of Biological Sciences, Oakland University, Rochester, Michigan, USA

Clifford M. Les Bone and Joint Center, Henry Ford Hospital, Detroit, Michigan, USA

George Lust James A. Baker Institute for Animal Health, Cornell University, Ithaca, New York, USA

INTRODUCTION Articular cartilage is a soft connective tissue that protects the underlying bone in synovial joints and is responsible for distributing mechanical stress. The tissue consists of a network of collagen fibrils, embedded in a hydrated proteoglycan matrix. In healthy (and unloaded) cartilage, the alignment of collagen fibrils in the network differentiates the tissue histologically into three zones: superficial (or tangential) zone (SZ), where the collagen fibrils run parallel to the articular surface; transitional (or intermediate) zone (TZ), where the collagen fibrils run obliquely and randomly; and radial (or deep) zone (RZ), where the collagen fibrils run mostly perpendicular to the articular surface [1–4]. This highly ordered ultrastructure helps to maintain the shape of cartilage and to provide biomechanical stiffness and integrity to the soft tissue. A mechanical load will inevitably result in deformation of the collagen matrix in articular cartilage. If a loaded specimen can be studied in high-resolution imaging, the extent of the local fibril deformation could provide important information regarding the fibril structure and molecular interactions at different tissue depths. Several scanning electron microscopy (SEM) studies examined the effect of mechanical loading on the organization of the collagen fibrils in cartilage matrix, in either excised cartilage specimens [5, 6] or intact joints [7, 8]. The deformation of chondrocytes also was investigated as a means of quantifying cartilage deformation as a function of joint loading [9–11]. To preserve the deformation, specimens in those studies were either chemically fixed [10, 12] or were excised from intact joints after cryofixation [7, 13].

We studied the deformation of the extracellular matrices in articular cartilage using a new compression-preservation method in histology. A Hoffman clamp was used to compress the tissue, which remained throughout the paraffin procedure and was removed from the embedded tissue block just before microtoming. Then 14 cartilage-bone blocks from 2 canine humeri were compressed for various strain levels from 5% to 65%. The histological sections were studied using a polarized light microscope, which generated a pair of two-dimensional maps of the fibril orientation (angle) and fibril organization (retardance) for each section. Results were 3-fold. One there was little change in the angle and retardance profiles of the tissue for strain levels 0–15% and a significant change in these profiles for strain levels 15% and above. Two for higher compression, more fibrils became aligned parallel to the articular surface; and three at ∼30% strain, a second “transitional zone” was formed in the deep part of the tissue. We concluded that this novel compression procedure can be used effectively to study the altered architecture of the collagen matrix in compressed cartilage. YX and GL would like to dedicate this paper to a co-author of this study and our long-time collaborator in cartilage research, Nancy Burton-Wurster (1941–2006). Keywords

Cartilage, Collagen, Compression, Paraffin Embedding, Polarized Light Microscopy

Received 24 August 2006; accepted 20 November 2006. Address Correspondence to Yang Xia, PhD, Department of Physics, Oakland University, Rochester, Michigan 48309, USA. E-mail: xia@ oakland.edu

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ALTERED MATRICES OF COMPRESSED CARTILAGE

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FIG. 1. (a) Beagle humeral head where the specimens were harvested from the central weight-bearing area (the black lines show the location of specimens). Gross inspection revealed no signs of injury or disease. (b) Schematic diagram shows the compression of a cartilage-bone specimen in the clamp (not to scale). The three labels SZ, TZ, and RZ (and the short lines and crosses) illustrate the collagen orientations in three histological zones of a cartilage-bone block.

In our study, the aim was to develop a straightforward static compression procedure to compress the cartilage tissue and to maintain the deformation of the collagen matrix in cartilage throughout the paraffin embedding procedure. We hypothesized that this novel procedure would have the ability to preserve the morphological environment of the loaded cartilage. The deformed collagen structure in cartilage was imaged by a quantitative polarized light microscopy (PLM) technique, which has the ability to construct two-dimensional (2D) images of the fibril orientation (angle) and the fibril organization (retardance) [14, 15]. Although this imaging technique does not have the resolution to visualize individual fibrils in ultrastructure [16], this technique is capable of detecting a group of collagen fibrils in each pixel based on optical birefringence [14]. MATERIALS AND METHODS Specimen Preparation Fully 14 osteochondral specimens were excised from the central load-bearing region of 2 humeral heads obtained from 2 healthy mature beagles sacrificed for an unrelated experimental study (Figure 1a). The interface between the soft tissue and the bone was preserved, and the location and orientation of the individual specimens on the joint surface were closely monitored to identify any influence of topographical variation. First, 9 specimens were compressed for different strain levels (5–65%) and were prepared for histology while compressed. Second, 5 specimens from the adjacent sites were selected to serve as uncompressed tissue. Each specimen block had the dimensions of 1.5 × 2 × 7 mm, and the underlying bone of each specimen was polished with a grinding pad. The specimens were bathed in physiological saline with 1% protease inhibitor

cocktail (P2714 Sigma, St. Louis, MO, USA) before compression. Animal subjects were handled according to the protocols approved by the Institutional Animal Care and Use Committee. The unconfined compression was performed by placing the fresh cartilage-bone plugs in Hoffman clamps (Fisher Scientific, Pittsburg, PA, USA) and compressing by the means of screws (Figure 1b). The compressed specimens remained in the clamps during the entire histological process (fixation, decalcification, paraffin processing, and embedding) to ensure the preservation of the local environments in the compression state. The standard histology procedure was used to process the specimens, including fixing them over night in buffered formalin solution, decalcifying them in fresh EDTA solution every other day for a week, and processing/embedding them in a paraffin tissue processor (RMC 1530, Tuscan, AZ, USA). The clamp was removed in the paraffin container just before the specimen was sectioned into multiple 6-µm thick sections. The unstained sections were placed on microscope slides, mounted, and covered for examination under the microscope. Polarized Light Microscopy (PLM) Our PLM system consists of a Leica polarized light microscope equipped with a 12-bit CCD camera [14]. The image acquisition uses circularly polarized light together with a liquid crystal compensator consisting of two retarders such that the retardance of each could be varied separately under computer control. This configuration allows for the compensation of birefringence in a specimen of any orientation without requiring its rotation or mechanical movement of the optical components in the light path. For each histological section, multiple intensity images were acquired at different settings of the liquid crystal compensator. The method of Oldenbourg and Mei [17] allows


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the calculation of two quantitative images, representing the optical retardance and the angular orientation of birefrigent elements in the tissue. For articular cartilage, the angle maps represent the pixelaveraged orientations of the collagen fibrils in the tissue, where the fibrils are expected to have an ∼90◦ difference between the surface and the deep zones. The retardance maps are influenced by several factors in the measurements, including the randomness of the collagen fibrils, the fibril diameter, the packing density of the fibrils, and the thickness of the histological section. For a single section, a smaller retardance value generally indicates that the collagen fibrils are less ordered. At the 5× objective, a retardance sensitivity of 0.32 ± 0.17 nm was found in our system (the rms noise level ± the standard deviation of the noise level). The pixel size of the 2D images was 2.72 µm/pixel for a 5× objective and 0.34 µm/pixel for a 40× objective. Other experimental details for the PLM system and procedures have been documented extensively elsewhere [14]. Data Analysis Five to ten histological sections were processed for each specimen, and 1D profiles from the angle and the retardance image were extracted from several regions of interests (ROI) from each section. The final 1D profiles were the averaged profiles across the available sections at the identical ROI from each specimen. Of the 14 specimens, 5 compressed specimens were matched with 5 adjacent control specimens, and the compression was determined by measuring the change in cartilage total thickness from each compressed-control pair. For the other 4 compressed specimens, the total thickness of the cartilage was determined previously from the microscopic

magnetic resonance imaging (µMRI) results and was used to calculate the specimen compression. The depth in the 1D profiles was normalized; 0 corresponds to the articular surface and 1 corresponds to the cartilage-bone boundary. The normalized thickness of each histological zone was determined using the criteria previously validated and documented [14]. RESULTS Figure 2 shows the calculated angle and retardance images of 2 cartilage sections from a pair of adjacent specimens, compressed to 0% and 12% strain values. The arrows point to the change in total thickness as a result of compression. In the angle map of the normal and unloaded tissue, the collagen fibrils close to the surface have a nominal orientational difference of 90◦ from the fibrils deep in the tissue [14]. In the retardance map, the uncompressed specimen reaches a minimum at a normalized depth of ∼0.07 from the articular surface, marking the center of the transitional zone that has the most random orientation [18]. Under compression, the features of cartilage tissue in both angle and retardance maps change (Figure 3). Figure 3 compares the 1D angle and retardance profiles at selected strain values from several specimens. Several features are apparent. In Figure 3a at the 15% strain value, the transition in the angle profile from 0◦ to 90◦ moved deeper into the tissue from ∼0.07 normalized depth to 0.2 normalized depth (the first solid arrow in Figure 3a), suggesting that more fibrils near the surface of the loaded tissue become parallel with the articular surface. At this strain, the retardance profile shows a reduction in its values but maintains the essential shape of its profile. In Figure 3b at the 30% strain value, the changes in the images and profiles became more prominent. For the angle profile, the

FIG. 2. Angle maps (top row) and retardance maps (bottom row) from the sections of two adjacent specimens: one was the control (no compression) and the other was compressed at ∼ 12% strain. The resolution of the images was 2.72 µm/pixel. The scales for the angle and the retardance maps are degrees and nanometers, respectively.



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FIG. 3. 1D profiles extracted from several selected regions of interest of (a) angle maps and (b) retardance maps at different values of strain (solid dots = 0%, open squares = 15%, crosses = 30%). Two solid arrows in (a) point to the expansion of SZ in compressed tissue. The hollow arrow indicates the formation of the second transitional zone when the tissue was compressed by 30%. For the uncompressed tissue (0% strain), three labels (SZ, TZ, and RZ) mark the approximate location of the histological zones along the tissue depth.

“surface zone” became further widened (the second solid arrow in Figure 3a); In addition to the original transitional zone, a second “transitional zone” was formed in the deep part of the tissue (∼0.4 normalized depth, marked by the hollow arrow). For the retardance profile, the minimum in the retardance, which used to be a narrow dip, became a wide region occupying ∼40% of the total tissue thickness. To study the changes in collagen fibril organization within cartilage matrix as a function of strain, the thicknesses of the three histological zones were determined from 1D angle profiles using the previously validated criteria [14], shown in Figure 4. (The thickness of the “newly formed zones” was not considered.) It is clear that there were no significant changes in the normalized thickness of three zones from 0% to 15% strain values and that the most drastic changes in the normalized thickness occurred at the 20%–40% of strain values. Within this 20%–40% strain range, the normalized thicknesses of both the ‘superficial zone’ and ‘transitional zone’ increased dramatically, whereas at the same time the normalized thickness of the ‘radial zone’ decreased.

Figure 5a shows the angle map of one specimen that underwent uneven compression, creating a gradient of strain from one side to the other. The right side was the less compressed part of the tissue and the left side was the most compressed part of the tissue. The area that showed the most complicated changes (compressed in the strain gradient of 15%–30%) was imaged at a higher magnification (40× objective); and the angle image is shown in Figure 5b. The change in the collagen fibril orientation across the tissue as a result of various values of strain is striking. (Because a higher magnification translates to a more restricted field of view, the tissue at 40× was imaged in several independent imaging acquisitions across the depth/width, and the calculated retardance and angle images were pieced together digitally. A black line was purposely left between any two images to indicate the boundary.) The higher resolution image at 0.34 µm/pixel also enabled us to examine some specific details in the tissue, for example the shape of the chondrocytes. Ultrastructural studies show that a cell in cartilage can be surrounded by an apparently empty zone, a moat of amorphous material, and finally by a territorial


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FIG. 4. Relative thickness of the three histological zones (SZ, TZ, RZ) as a function of strain. Most data points correspond to the average of two or three tissue sections at the same local strains; the data at13% and 20% came from one specimen.

matrix consisting of some collagen fibrils organized as ordered circular “cocoons” around the clusters of chondrocytes [19, 20]. Figure 6 shows the angle maps of two chondrocytes, one from a location where the tissue was compressed by ∼ 15% strain and the other from a 30% strain location. Both locations are about the same depth from the articular surface. The deformation of the cell due to external strain is clear. In Figure 6a, inside the region that used to have the cell, the random angles indicate that this region has no ordered fibrils. Outside the cell region, the collagen fibril orientation in the circular cocoons can be measured. Figure 7 illustrates the angle measurement of the “cocoon fibrils” from several cells under different strains. Around each cell, 8 locations were measured, marked by the white arrows and labeled as 1 to 8 in Figure 6a. At 0% strain (no compression), the angle map shows the periodic variation of collagen fibril orientation surrounding the chondrocyte, as demonstrated in one of our previous studies [21]. (The angle calculation in our PLM technique falls into the range of 0–180◦ because the collagen fibrils have neither head nor tail.) At ∼ 20% strain, the periodic variation, even weaker, can still be seen. At any higher strain (eg., 29% and 35%), there were no periodic differences in fibril orientation around the chondrocytes, indicating the highly flattened cell shapes at high strains. DISCUSSION In this article, we report on using a mechanical clamp to compress the cartilage-bone block and a paraffin tissue processing protocol to maintain the compressed fibril architecture in articular cartilage. Compared with several reported protocols in the literature (eg, chemical fixation [10, 12], and cryofixation [7, 13]), our unconfined compression method is relatively straightforward and the paraffin embedding procedure has been the standard method in histology. To the best of our knowledge, this is a novel combination because it preserves

the compressed environment of cartilage while the tissue is being processed for histology. Because of the simplicity of our compression device, some specimens in our study were compressed unevenly. Although this might seem undesirable, our results show that uneven compression can be used to study the morphological changes of the collagen network as a result of different strain levels in the same piece of specimen. The deformation of the collagen matrix in cartilage was quantified using PLM that generated two 2D images: the retardance map that illustrated the local fibril organization and the angle map that illustrated the local fibril orientation. We have shown in our previous work that this digital PLM method is an effective tool in the study of the collagen architecture in cartilage [14, 15, 21]. At higher magnifications, we showed that the orientation of the collagen fibrils that forms the cocoon-shaped territorial matrix surrounding the clusters of chondrocytes can be imaged by our PLM technique (Figure 6) [21]. The ability to map both the fibril organization and fibril orientation demonstrates that even though the PLM technique does not have the resolution to visualize individual fibrils, this technique is capable of detecting a group of collagen fibrils such as the circular cocoons around the chondrocytes. In this PLM study of compressed cartilage, the zonalspecific changes in compressed cartilage correlated well with the previous µMRI findings [22]. We showed that mechanical compression above physiological strain level can have profound effects on the orientation of macromolecules, predominantly collagen fibrils, in each histological zone. Our observation of zonal deformation is consistent with a scanning electron microscopy study of bovine femoral cartilage, which showed an expansion of the upper zone depth (the superficial and transitional zone) at the costs of the radial zone [6]. The crimped form of collagen fibrils under mechanical loading has been observed previously in several studies [8, 20, 23].

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FIG. 5. (a) Angle map of cartilage tissue under uneven compression at 5× objective (2.72 µm/pixel). The strain increases from the right side to the left side. (AS = articular surface, AC = articular cartilage, TM = tide-mark, CC = calcified cartilage, and B = bone). (b) Angle map of the region in (a), which is marked by the rectangular box, imaged using a 40× objective (0.34 µm/pixel). Note that the final image at the 40× objective was pieced together by multiple independent images, each from a small region of the tissue.

Note that the discussion of tissue zone’s “growth/change/ formation” in this article does not mean that a particular part of the cartilage now belongs to a different histological zone under compression. The concept means that the orientational changes of the collagen fibrils due to external loading make them appear to have a different birefringent property in PLM (hence

belonging to a different zone). It is well known that compression of articular cartilage results in inhomogeneous consolidation of all zones of uncalcified cartilage, with the superficial zone being more compliant than the mid/deep zones [24]. However, the superficial zone of cartilage clearly increases in thickness when the tissue is loaded, either by µMRI [25] or by PLM (this work).



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FIG. 6. Angle images of two chondrocytes from two locations of the 40× objective image (0.34 µm per pixel). Both locations were close to the articular surface: (a) one cell from the right side of the image in Figure 5b (∼15% strain), and (b) one cell from the left side of the image in Figure 5b (∼ 30% strain). The arrows and labels mark the locations where the angle values are measured around the cell.

To account for these seemingly paradoxical findings, we previously hypothesized that this altered characteristic in compression is based on structural reorganization or reordering of the fibrillar collagen architecture [25]. Loss of interstitial water from the compressed cartilage concentrates the solid phase of the tissue and enhances the molecular interactions, and these imaging techniques appear exquisitely sensitive to the reorganization of the collagen fibrils, particularly near the articular surface. In fact, we have carried out repeated imaging

experiments for samples before/during/after (moderate) loadings and found that the tissue’s orientation and properties are recoverable once the specimen is allowed to rehydrate in saline. In summary, since the collagen matrix maintains the integrity of articular cartilage and cartilage plays a critical role in distributing mechanical stress in joints, it is important to characterize the altered environment of the collagen matrix under different mechanical stresses. This PLM study demonstrated that it was possible to preserve the altered morphology



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FIG. 7. Fibril orientation around the single cells, as marked in Figure 6a, at 0%, 20%, 29%, and 35% strain levels. An undeformed cell has an elliptical/round shape, which results in the collagen fibrils in the territorial matrix (solid dots) circling around the cell (from locations 1 to 3, 3 to 5, 5 to 7, and 7 to 1). As the strain increases, the cell is flattened out, which results in a flat profile of fibril orientation.

of the compressed cartilage using a novel paraffin protocol and to visualize this altered morphology using PLM. The understanding of the morphological adaptability of cartilage in various compressed conditions will be useful in characterizing the events in mechanically induced joint diseases and injuries. ACKNOWLEDGMENTS Yang Xia acknowledges the following funding sources: Research Excellence Fund in Biotechnology from Oakland University, an instrument endorsement from R.B. and J.N. Bennett, and an R01 grant (AR 45172) from the NIH. Hisham Alhadlaq expresses his gratitude to Professor Yang Xia, Oakland University, for hosting him over the summer of 2006 and to KACST, the national funding agency in Riyadh, Saudi Arabia, for making his summer trip to the United States possible. Niloufar Fozouni and Ilco Aksovski (Department of Physics, Oakland University) helped in histological processing of the specimens. Clifford Les acknowledges the NIW funding (RO1 AR47434). REFERENCES 1. Aspden, R.M., and Hukins, D.W. (1981). Collagen organization in articular cartilage, determined by X-ray diffraction, and its relationship to tissue function. Proc. R. Soc. Lond. B. Biol. Sci., 212, 299–304. 2. Muir, H., Bullough, P., and Maroudas, A. (1970). The distribution of collagen in human articular cartilage with some of its physiological implications. J. Bone. Joint. Surg. Br., 52, 554–563. 3. Broom, N.D., and Marra, D.L. (1985). New structural concepts of articular cartilage demonstrated with a physical model. Connect. Tissue Res., 14, 1–8. 4. Broom, N.D., and Silyn-Roberts, H. (1989). The three-dimensional ‘knit’ of collagen fibrils in articular cartilage. Connect. Tissue Res., 23, 75–88. 5. Kobayashi, S., Yonekubo, S., and Kurogouchi, Y. (1996). Cryoscanning electron microscopy of loaded articular cartilage with special reference to the surface amorphous layer. J. Anat., 188(Pt 2), 311–322.

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