Critical Size Defect in Adult Articular Cartilage: A Preclinical Study

Abstract

Objective

Lesions of adult articular cartilage occur due to trauma or disease, such as osteoarthritis. If they do not penetrate the subchondral bone, they are called partial-thickness defects (PTDs), which are believed not to heal. However, some reports indicate that minor PTDs can be repaired. We hypothesize that a critical-size PTD exists below which spontaneous healing occurs.

Design/Methods

In an adult pig model, we created PTDs of minimal width (a scalpel cut) and systematically increased their width up to 0.5 mm. Defect analyses were conducted at 1 and 3 months post-surgery using light microscopy and histomorphometry.

Results

None of the defects healed by repair cartilage; therefore, all PTDs are of a critical size. Surprisingly, a critical defect-size range was identified where significant mesenchymal tissue (MT) formation occurs, specifically in defects measuring 50-100 μm in width. The presence of this MT was limited to a 1-month time window. Furthermore, physiological joint loading during the postsurgical phase was associated with substantial structural tissue deformation, often leading to an overlapping of the side walls of the smallest defects. This results in a pseudo-covering of the defect void, which may thus be invisible when observed from above.

Conclusions

The main novel finding of this study is that there is no critical width below which PTDs undergo repair.

Keywords

articular cartilage repair critical-size defect partial-thickness defect spontaneous repair

Introduction

Sports injuries and other types of traumas, or surgical interventions in synovial joints (arthroscopy, meniscectomy, etc) may lead to or be associated with the creation of structural lesions of the articular cartilage. They may also occur during the early stages of osteoarthritis.¹ As long as such lesions remain confined to the articular cartilage layer itself, they are called partial-thickness defects (PTDs).² PTDs generally do not heal spontaneously in the adult organism.^3,4 PTDs are to be distinguished from full-thickness articular cartilage defects (FTDs), which, by definition, span the entire articular cartilage layer and penetrate the underlying subchondral bone. Concurrent damage to osseous blood vessels leads to bleeding and a spontaneous repair response.⁴

The lateral width of PTDs generally falls within the millimeter-to-centimeter range. However, literature suggests that PTDs with widths below the millimeter mark, such as those created by a simple scalpel-blade cut, can heal spontaneously.^5,6 However, whether there is a critical size below which PTDs heal on their own, similar to FTDs or bone defects,^7,8 has not been systematically explored.

We suggest that, even for PTDs, a critical defect size exists—below which they heal on their own and above which they do not. The current study aims to identify this parameter.

The relevance of this parameter to orthopedic surgeons, rheumatologists, arthroscopists, and traumatologists pertains to many practical issues that need to be considered, such as, for example:

The treatment of joint fractures implicating the articular-cartilage layer and the subchondral bone tissue: Is the precision of the bony and articular cartilage reduction crucial for the repair of the cartilage layer?

Are additional measures, such as compressive reduction, necessary to improve the healing of the cartilage layer (which becomes a PTDS if a perfect bony reduction is achieved underneath the cartilage layer)?

Is an additional bioactive therapy necessary to restore the structure and, by implication, the functionality of the articular cartilage layer, particularly above a critical-size cartilage defect?

Do small (narrow) PTDs, generated inadvertently during arthroscopy, heal spontaneously?

What are the requirements for a tissue-engineering approach aimed at restoring the structure and function of a PTD caused by trauma or during early osteoarthritis? For example, is lateral annealing necessary between an implanted tissue-engineered construct and the parent tissue wall? And if so, above what critical defect width?

Determining the critical size of PTDs will help answer these questions and indicate whether supplementary postoperative measures, such as targeted physiotherapy, are likely to be beneficial or harmful.

To determine whether PTDs generated within adult articular cartilage are characterized by a critical size below which they heal spontaneously, we conducted in vivo experiments with skeletally mature Goettingen miniature pigs. PTDs, starting with the smallest possible defect width of a scalpel cut (theoretically, a few nanometers to 2 µm wide) and gradually increasing defect widths up to 0.5 mm, were surgically created. The repair response was analyzed histologically, histochemically, and histomorphometrically at 1 and 3 months after defect creation.

Materials and Methods

Study Design

PTDs were surgically created in adult Göttingen miniature pigs (2–4 years old) in eight experimental groups with varying defect widths (Table 1). At 1 and 3 months post-surgery, the defects were examined using light microscopy, histochemistry, and histomorphometry to evaluate healing responses, including the types of tissue involved, the pattern of repair tissue growth, and the quantitative levels of these responses. Chemical preservation of cartilage was performed using a cationic compound, cetyl pyridinium chloride (CPC), to prevent proteoglycan loss from the cartilage matrix and preserve cell shape during histological processing.

Table 1.

Experimental Groups/Defect Widths (in µm).

≈0-2

100

200

300

500

Preliminary In Vitro Experiments

Using fresh knee joints from adult female miniature pigs (which had been used for cardiovascular studies by another research group), we tested a specially designed sawing tool (see below) to evaluate its ability to produce consistent sawing depths and defect widths. We also assessed whether it reliably creates vertical defect edges and channel-like excavations of the intended width. To do this, tissue blocks were cut from the native material with an Exact Diamond saw band (Exakt Technologies, Inc., Oklahoma City, OK) and frozen in liquid nitrogen; cryosections were stained with Safranin O. Several tissue blocks were chemically fixed, dehydrated, embedded (see details below), and stained to determine if these histological processing steps can preserve the original defect shape and geometry. Ensuring that no such artifacts occur was crucial to accurately evaluating potential shape changes during natural postsurgical joint loading. Additionally, these preliminary studies helped quantify the amount of tissue debris (hyaline cartilage) produced during the sawing process.

Experimental Animals, Anesthesia, Surgery

We used 24 adult female Göttingen miniature pigs (aged 2-4 years). General anesthesia was induced by intravenous injections of Vetalar® (2-bromo-2-chloro-1,1,1-trifluoroethane) and maintained with nitrous oxide (via intubation) and ketamine.³ Each animal underwent surgery on one of its knees. PTDs were created using a dental saw with blades of specified thicknesses (Table 1).

The stainless-steel sawing blades were manufactured in-house at our mechanical workshop on a custom basis. Blade thicknesses (25 μm-500 μm) were measured using a digital caliper and a stainless-steel micrometer to confirm accuracy. Depth control of the PTDs was achieved by placing round metal disks (approximately 0.6 mm thick) laterally on each side of the saw blade (Fig. 1). The disk’s outer cutting surface circumference was designed so that it ended about 100 μm below the saw blade`s cutting circumference. During cutting, it was ensured that the disks only touched the articular cartilage surface, thus achieving the desired sawing depth of approximately 100 μm with the rotating saw blade. The bilaterally placed support disks also helped mechanically stabilize the thin sawing blades. The reproducibility of this procedure was verified using adult Göttingen miniature pig knee joints (see above: preliminary experiments). The freshly created defects were then analyzed for correct thickness, depth, and shape preservation using cryosections (see above).

Figure 1.

(A) Photograph of the sawing instrument showing the fine central sawing blade (blue arrow) and the metal support discs on each side of the blade, which mechanically stabilize this delicate tool (green arrow). (B) Macroscopic photograph of a Göttingen miniature pig`s distal femoral end, showing the patellar groove and the condyles.

Practically zero-width defects (i.e., the smallest possible) were created by vertical scalpel-blade incisions using an ophthalmic diamond scalpel (Rhein Medical Inc., St. Petersburg, FL). The edge thickness of this diamond blade is reported by the manufacturer’s data sheet and in the scientific literature to be in the order of tens of nanometers.⁹ Defect lengths were limited to 10 mm (Fig. 2).

Figure 2.

(A) Drawing of a pig’s knee joint. The topographical locations of the experimental defects are marked on the patellar groove and the medial femoral condyle. (B) Graph showing the mapping of defects and their dimensions (lateral distances between them, lengths).

Defect locations are shown in Figure 2 . Eight defect-group widths were defined, and 9 defects were created within each group. For each joint, one defect from the same group was made, with defect locations systematically varied.

Each animal had one joint undergo surgery. This setup resulted in 56 defects at each time point. Therefore, a total of 120 defects (including 8 reserves) were created for the entire study.

Animals were housed at a nearby farm in an on-site stable with access to an outdoor area for unrestricted movement. They received standard commercial pig feed. A DVM performed daily health assessments. Euthanasia was conducted under general anesthesia, followed by an intravenous infusion of potassium chloride to induce cardiac arrest.

The study received approval from the local animal ethics committee. It was conducted in accordance with its regulations, as well as Directive 2010/63/EU on the protection of animals used for scientific purposes, and the ARRIVE guidelines were strictly followed.

Tissue Sampling and Processing

Condyles were cut into bone blocks measuring 2 cm in length (Figs. 1 and 2); the patellar grooves were cut into two blocks, each containing the proximal and distal two defects. Blocks were sectioned perpendicular to the long axis of the defect into four slices of equal thickness, starting at a random point (systematic random sampling principle),¹⁰ using a Leco diamond saw (Leco Corporation, St. Joseph, MI). The first slice was prepared for cryofixation, the second and third slices for methylmethacrylate embedding (for 80 μm sections), and the fourth slice for Epon 812 embedding for semithin (1 μm) sections.¹¹

Light Microscopy and Histochemistry

Chemical fixation was performed using 4% glutaraldehyde containing 2.5% CPC to induce proteoglycan precipitation.¹² CPC is a colorless substance used here to enable surface staining of thick sections (which otherwise would not remain transparent for light microscopy). Dehydration was initiated with 70% ethanol, the minimum concentration necessary to prevent cartilage tissue swelling^11,13, and continued in a graded series to 100% ethanol. Tissue embedding was carried out in methyl methacrylate, and sections (approximately 200 μm thick) were produced (sampling strategy: see above) using a Leco diamond saw (Leco Corporation, St. Joseph, MI). The sections were then glued to plexiglass holders and milled down to about 80 μm in thickness. Section surface staining was performed with McNeil’s Tetrachrome and Toluidine Blue O (as previously described^3,12).

Cryotechnical Processing

Cryotechnical processing prevents chemical artifacts, preserves the original structure, and enables detection of potential shape changes in defects that may occur during the animal’s postsurgical, unrestricted movement, as well as additional structural changes that may arise during histological processing. Sections stained with Safranin O help identify potential proteoglycan loss during the postsurgical period¹². Cryosection thickness: 30 μm.

Morphological Analysis

LM analyses (thick and semithin sections) were performed with a focus on the shape of the defects and the numerical areal density of chondrocytes near the cut defect surfaces, comparing them to those in the deeper regions.

Mesenchymal tissue (MT) is an avascular tissue composed solely of undifferentiated stellate or spindle-like cells and a fine network of intercellular fibrils in which these cells are embedded¹⁴. This tissue type is mainly encountered during embryonic and fetal development.

Histomorphometry

To quantify the neoformation of repair and/or other tissue, each section was photographed, and positive prints were made. Test systems on a transparent foil were randomly placed over the prints. Using the standard point counting procedures (cf. Gundersen et al, APMIS,96(5)379-394), the volume fractions (percentages) of the following structures were estimated: total defect space, hyaline cartilage tissue debris (resulting from the sawing process), newly formed fibrocartilage tissue, mesenchymal tissue, mesenchymal cells, and empty defect space.

Statistics

All statistical analyses were performed using EZR software, version 1.64.¹⁵ Data comparisons among multiple groups were conducted with one-way ANOVA, followed by Tukey’s multiple comparison test.

Results

The preliminary experiments showed that the sawing tools produce the desired vertically channeled defects with the intended shape and width (Figs. 3 and 4). Figure 3 depicts the histology of normal adult knee cartilage (Goettingen miniature pig). A freshly created defect, chemically fixed, dehydrated, and embedded in plastic, demonstrates that the original shape and geometry of the defect are preserved, indicating that tissue processing (dehydration and embedding) is homogeneous and symmetrical at and around defect areas and does not alter the defect’s original structure.

Figure 3.

(A) Light micrograph showing the structure of the normal adult articular cartilage of the Göttingen miniature pig at low magnification. Stained with McNeil`s Tetrachrome and Fuchsin Red. Abbreviations used: S = superficial zone; T = transitional zone; UR = upper radial zone; LR = lower radial zone; C = calcified cartilage zone. Magnification bar = 50 µm. (B) Light micrograph of a surface-stained thick section (80 μm thick) of a vertical partial-thickness defect measuring 250 μm in width (red double arrow). The shape and vertical orientation of this defect are preserved, demonstrating that dehydration and embedding methods do not cause artifactual structural distortions in tissue. Stained with McNeil`s Tetrachrome and Fuchsin Red. Magnification: see the double arrow indicating 250 μm in length.

Figure 4.

Light micrographs of cryosections from freshly made defects, stained with Safranin O. Defect widths are in (A): 250 µm, (B): 150 µm, (C): 200 µm. The different defects show the variability in size, shape, and location of sawing debris or shearings of hyaline articular cartilage. Magnifications match the defect widths.

The cryosections (Fig. 4) of freshly made defects also showed (and confirmed) the desired vertical edges of the defect across different sizes. They also revealed that the sawing process produces shearings (debris) from articular cartilage within the defect void, occupying roughly 5–40% of the defect space (Fig.7). This confirms that the sawing tools can create cartilage defects with the desired geometry. These defects remain structurally intact throughout histological processing, which is essential for identifying postsurgical shape changes resulting from joint use.

Visual inspection of the joint surfaces after euthanasia revealed that not all defects remained detectable. Specifically, simple scalpel cuts and minor defects (up to 100 μm) were no longer visible in approximately 30% of cases, suggesting, based on a simple visual inspection, that they had healed. However, this is a false and misleading impression for the surgeon (see the following paragraph for explanation). The precise topographical mapping of the defects (Fig. 2B) was thus essential for reidentifying the defect sites for analysis.

Histomorphological examination showed that none of the created defects had healed with fibrocartilage repair (FCR) tissue (Figs. 5 and 6). Therefore, all partial-thickness defects (PTDs), ranging from 0-2 μm in width (scalpel cut) up to 500 μm, are of critical size in adult articular cartilage. The systematic review of the different defect-size groups revealed that most of them remained empty.

Figure 5.

Light micrographs of three scalpel cuts, taken at different magnifications, reveal the wide variability in their structure and defect volume 1 month after creation. (A) The image shows a scalpel cut that remained nearly unchanged during joint use but has a slightly widened defect volume. (B) The image depicts a cut with further widening of the space, likely caused by joint loading and internal stretching or strain forces at that location. (C) The image shows an irregularly shaped cut (highlighted by a black dashed line for clarity) after 1 month in situ. The variable internal strain and stress forces within the tissue probably contributed to this unusual shape. Staining with Toluidine Blue O (A, B) and Fuchsin Red (C). Magnification bars: 50 μm.

Figure 6.

Light micrographs showing defects observed 1 month after surgery. Defect widths are: (A): 25 μm, (B): 50 μm, (C): 75 μm, (D): 100 μm, (E): 100 μm, (F): 100 μm. Figures G and H display defects 3 months post-surgery, with widths of 25 μm in Figure G and 100 μm in Figure H. These images illustrate the significant changes that occur during the postsurgical period under normal knee joint loading. On one hand, the defect widths vary widely, and the original defect sizes often become unrecognizable. The shapes of the defects also change markedly—from highly irregular (A, B, C, G) to V-shaped (E), and to minor variations (D, F, H)—depending on the topographic location within the joint and the internal forces of strain and stress acting on the cartilage during loading. Conversely, some defects exhibit minimal shape change from their original form (D, F, H). The varying degrees of defect filling with repair connective tissue are clearly shown, with little to no filling in A and C, compared to nearly complete filling in D, F, and H. In addition, the overlapping of defect side walls creates the appearance of defect closure and repair when viewed from above the native joint surface. These phenomena are clearly visible in A, B, and G. The staining methods used include McNeil’s Tetrachrome and basic Fuchsin. Magnification bars are 100 μm.

In particular, the PTDs created by the mere incision by the corneal scalpel blade remained “empty” (viz., devoid of neo-formed tissue) throughout the entire course of the follow-up period (i.e., up to 3 months). No repair cartilage tissue nor mesenchymal tissue (MT) was visible at either 1 month or 3 months after surgery (Fig. 5); specifically, no individual repair cells, such as mesenchymal cells, were observed adhering to the defect surfaces.

PTDs of 25 μm, 200 μm, 300 μm, and 500 μm showed partial filling with mesenchymal tissue (MT), which was less than 20% of the defect volume in each of these groups (Fig. 8) at each sampling time point. In contrast, those with widths of 50 μm, 75 μm, and 100 μm showed a significantly higher degree of defect filling with MT, ranging from 45% to 55% of the defect volume (Fig 8). However, there was no evidence of remodeling of the MT into fibrocartilage in any of the PTDs, regardless of width or sampling time. These findings suggest that there is no lower width-limit (i.e., critical size) below which FCR is formed.

A notable and novel finding was that the original shape of the defects—a long channel with vertical walls (Figs. 3 and 4)—was not preserved; instead, they deformed into irregular shapes. This was especially true for minor defects, between 25 μm and 100 μm in width, and particularly for scalpel cuts. The opposing cut surfaces often did not stay in contact but became secondarily separated, creating a void between them or causing them to overlap (Figs. 5 and 6). These shape changes were less apparent in larger defects (~200 μm and wider) (Fig. 6).

For these reasons, upon simple visual inspection, the scalpel cuts and the minor (25-100 μm) defects were often no longer visible from above the articular surface to the naked eye.

The final width of the scalpel cut defects at the time of sacrifice varied depending on the anatomical location in the joint. In areas with a convex joint surface shape (femoral condyle), the defect widths tended to become wider (up to a few µm). In contrast, in concave-shaped or flat areas (mainly the patellar groove), they tended to remain of zero µm width. This may relate to the cartilage’s site-specific internal structure and the predominant tissue internal forces, which are of a tensile and/or compressive nature.

As expected, the defects created by the sawing method contained hyaline articular cartilage fragments from the sawing process itself, meaning shearing debris generated during defect excavation (Fig. 4). All defects (25-500 μm wide) contained a similar amount of this “sawdust” that filled about 5%-35% of the defect volume (Fig. 4); the amount of sawdust retained varied significantly among different defect-size groups (0%-35%; Fig. 7). The retention of these tissue fragments remained consistent over time for up to 3 months (Fig. 7). These unavoidable fragments are a sawing technique induced artifact of the experiment. They are considered part of the empty defect space.

Figure 7.

(A, B) Graphical representation of histomorphometry estimates for articular cartilage sawing debris (fragments) encountered in defects at 1 and 3 months postsurgically (expressed as percentages of total defect space, compared to the remaining empty defect space). The percentages of cartilage fragments varied considerably with high coefficients of error (Table 2); only the 25-μm and the 300-μm defects at 1 month showed significantly lower fragment percentages in their defect spaces than the other defects. The space these fragments occupied was assigned, in the final analysis, to the defect’s empty space (since this is not a repair activity).

Table 2.

Summary of the Coefficients of Error (CE) expressed as a percentage of the mesenchymal tissue response (MTR) to defect creation in each experimental group (excluding scalpel-blade cuts), and a comparison of the MTR percentages at 1 month post-surgery between groups that showed significant differences in MTR activity, namely the 50 μm, 75 μm, and 100 μm defect groups, compared to larger defect groups and the 25-μm defect group (which has lower P-values).

A: 1 Month			B: 3 Months			C: Significances
Width	n	MT	Width	n	MT	Comparisons	MT
(μm)		CE (%)	(μm)		CE (%)	(width in μm)	(at 1 Month)
25	7	35	25	11	31	200-25	P = 0.183
50	8	15	50	11	50	300-25	P = 0.330
75	4	28	75	10	36	500-25	P = 0.403
100	6	28	100	12	30	200-50	P < 0.01
200	7	100	200	10	51	300-50	P < 0.01
300	9	76	300	6	53	500-50	P < 0.01
500	10	42	500	14	42	200-75	P < 0.05
						300-75	P < 0.05
						500-75	P = 0.054
						200-100	P < 0.01
						300-100	P < 0.01
						500-100	P < 0.01

Interestingly, the high levels of MT filling in defects with widths of 50 μm, 75 μm, and 100 μm did not increase over time, suggesting that these MT activities are not progressing in an anabolic direction; instead, at the 3-month mark, the MT filling in these three defect sizes was reduced to the level of the other defects (Fig. 8); this indicates that degradative remodeling activities are occurring.

Figure 8.

Graphical representation of histomorphometry estimates for mesenchymal tissue response (MTR) percentages in defect areas compared to empty areas. One month after surgery (A), minor defects measuring 50-100 μm exhibit the highest MTR activity. The 25-μm defect group exhibits significantly less activity but still more than defects wider than 200 μm. This highlights a critical defect-size window of 50-100 μm. Scalpel cuts did not demonstrate any MTR activity (not illustrated). At 3 months (B) post-surgery, these high MTR activities are no longer observed, indicating that a critical size and time window for MTR also exist. This suggests that any therapeutic intervention aimed at enhancing spontaneous repair, such as biological activation, must be applied within a critical early time window after trauma or surgery.

The finding that no fibrocartilage repair (FCR) tissue was present indicates a lack of active repair processes. The observation that the empty space (Fig. 8) is similar in size across all groups at 3 months postoperatively and remains stable over time suggests that ongoing tissue neoformation activity has ceased. Additionally, the high variability in the volume of the empty space among different defect-size groups at both 1 month and 3 months postoperatively further suggests that there is no effective repair tissue response.

Chondrocyte cluster formations appeared irregularly near the defect surfaces. Surprisingly, the histochemical matrix staining intensity of the cartilage matrix in the parent tissue at the immediate sub-incision surface remained unchanged throughout the 3-month experimental period, indicating that metachromasia (and thus proteoglycan content) did not decline.

The histomorphometry measurements (Figs. 7 and 8) show that the defect spaces contain minimal tissue filling and mostly remain empty (55%-85%, with notable variations; coefficients of error range from 15% to 100% (Table 2)). The sawing debris portions ranged from 5% to 45% (Fig. 7). Without these debris, the total remaining empty defect space would range from 75% to 95% across all groups, except for the 50-100 μm defect groups at 1 month (Figs. 8).

The presence of MT in defect areas typically ranged from 10% to 25%, except for those with a width of 50-100 μm, which showed significantly higher mesenchymal tissue neoformation activity, ranging from 45% to 55%. However, this increased MT neoformation activity was temporary and had ceased by 3 months after surgery. FCR does not appear at any time in any group.

Some defects were cut too deeply, reaching the subchondral bone tissue space; once identified, they were excluded from the analysis. As a result, the number of defects analyzed per group ranged from 4 (lowest) to 14 (highest); see Table 2 for the n-number details.

While it is generally understood that PTDs do not heal, this mainly applies to large PTDs.^3,4 Some studies suggest that small PTDs, like simple cuts, can heal.^5,6 The existence of a critical size for PTDs has not been systematically studied before. The new findings from this study show that all PTDs are of critical size and that there is a small defect-size range in which temporarily significant mesenchymal tissue responses occur, although they are ultimately abortive. Additionally, another new finding is that small PTDs are structurally significantly deformed under normal joint loads, which can make them potentially invisible when inspected from above, even if they did not heal. This could lead to a mistaken assessment of their outcome.

Discussion

Mesenchymal tissue (MT) was observed in all sizes of PTDs except for scalpel cuts, which showed none. This aligns with a previous study,¹⁶ indicating that an abortive repair process, such as MT formation in PTDs in adult articular cartilage, may occur and originate in the synovium. The cells responsible for repair are most likely transported to the defect site by the flow of synovial fluid.

A novel finding here is the presence of significant MT formation in small-width defects measuring 50-100 μm (Fig. 8). This suggests a critical defect-size window in which this initial, but abortive, repair-type process by the MT plays an important role. Another discovery is the time window for this MT-formation process, which ends within 3 months, likely due to degradative tissue remodeling activities, and the inability of this MT to spontaneously differentiate further into functionally competent FCR tissue, probably due to insufficient intrinsic chondrogenic differentiation activity. A previous study on large PTDs also showed a similar MT activity time window,¹⁷ in which, however, mainly mesenchymal cell adhesion to the defect surfaces was observed.

We interpret the existence of a critical-size window for PTD as resulting from specific synovial rheology conditions within these defect-size groups, leading to a significant, but abortive, MT-formation activity. The existence of defect-size-specific rheology, specifically its fluid streaming patterns, and the transport of synovium-derived mesenchymal cells for repair within its fluid may depend on local geometrical factors. The geometry of these defect groups is characterized by exceptionally high surface-to-volume ratios (SVRs). These SVRs are indeed highest in the 50-100 μm groups compared to other defect widths. Very high SVRs may indeed facilitate the temporary adhesion of mesenchymal cells and tissues, which is necessary for the induction of FCR. Therefore, defects with high SVRs may specifically facilitate or even promote the spontaneous influx of these cells from the synovium, thereby facilitating their adhesion and tissue formation. Thus, these narrower defects with high SVRs tend to support the local biological environment (niche biology)¹⁸ for spontaneous, host-supported MT formation.

In the early postsurgical and post-traumatic phases of PTDs, proteoglycan metabolism may be impaired, possibly due to minor proteoglycan loss from lesion surfaces and from nearby intercellular spaces. Such minor losses were not detected in this study by metachromasia staining. However, it suggests that active cell and matrix metabolism is still ongoing, and that an abortive chondrocyte repair response is present, as indicated by the formation of cell clusters.^16,19

Proteoglycans are known for their anti-cell-adhesive activity, which prevents cell adhesion during the early postsurgical period and thus hampers cell-based repair processes.^20,21 However, these effects are reversible¹⁷ and are no longer observed three months after surgery. This may explain the limited time window during which MT is present.

Surprisingly, the scalpel cuts did not exhibit any repair-cell adhesion to their cut surfaces. The 25-μm defects showed repair-cell adhesion, though at lower levels than the 50- to 100-μm defects (Fig. 8). The 25-μm width aligns with the size of average cells in the body,²² especially synovial lining cells.²³ This suggests it might be a geometric issue, such as steric hindrance, that prevents more active cell migration and physiological rheology from transporting cells into this tiny defect space.

Lesions within the critical defect window size (50-100 μm) appear to have sufficient mesenchymal tissue temporarily present that could be stimulated to fill the defect space and complete repair by differentiation into FCR tissue, without needing additional defect space-filling carriers³ as long as the surgeon achieves proper reduction of the defect walls to this critical width and provides sufficient bioactive local stimulation (such as with growth factors).^3,24

The discovery that scalpel cuts and especially tiny defects become highly distorted, often with overlapping defect walls, has important implications for practical joint surgery: it can create a false impression for the surgeon during visual inspection that these defects have healed. Therefore, simple visual inspection alone is insufficient to accurately evaluate the repair response to minor defects, whether they are isolated defects or narrow gaps between tissue-engineered implants and native tissue, where tissue integration is intended to be induced. In particular, when therapeutically tissue-engineered constructs are deposited in cartilage defects, the side walls between the built and native tissue need to be firmly joined and mechanically stabilized using appropriate biological glues to ensure tight adhesion from the beginning of therapy.

The basis for this conclusion is our findings, which indicate that daily physiological load forces are linked to widespread but reversible structural distortions at both the tissue and cellular levels (compression/expansion effects), as well as to internal tissue strain forces, in areas with structural defects in articular cartilage. Because the force-absorbing collagen arcade structure becomes disrupted,^25,26 these deformations become abnormal in extent near defects. As a result, local overloading and underloading forces develop,²⁷ which are associated with the long-term progression of osteoarthritis.

In addition, if bioactive therapeutic measures are put in place (tissue engineering) to induce FCR tissue formation and tissue integration/annealing measures are taken, it appears essential to avoid even normal joint loading for a specific period of time to prevent excessive tissue deformations and fragment separations, which can happen even around minor defects. Therefore, it helps to temporarily prevent tissue deformation, thereby preserving local stability and supporting healing processes and remodeling. A suitable postoperative physiotherapy protocol might thus include, for example, intermittent passive joint motion,²⁸ which provides nutritional benefits to the cartilage while preventing harmful tissue deformation during cartilage healing.

Footnotes

Acknowledgements

The authors thank Eva Kapfinger and Britt Hoffmann for their technical assistance, as well as Dr. Kay Juergensen for his help with the surgery and logistics. The English language and grammar were corrected using the Grammarly software (San Francisco, CA, USA). This article is dedicated to the memory of Dr. Lawrence C. Rosenberg (1928-2012), a highly respected scholar and cherished friend.

ORCID iD

Ernst B. Hunziker

Ethical Considerations

The study received approval from the local animal ethics committee. It was carried out in accordance with its regulations, as well as Directive 2010/63/EU on the protection of animals used for scientific purposes, and the ARRIVE guidelines were strictly followed.

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This study was supported by the University of Bern, Bern, Switzerland, and the Spine-Pelvis, AG, Zurich, Switzerland.

Declaration of Conflicting Interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

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