JBJS | Histological Changes in the Human Anterior Cruciate Ligament After Rupture*

The Journal of Bone & Joint Surgery, Volume 82, Issue 10

Articles | October 01, 2000

Histological Changes in the Human Anterior Cruciate Ligament After Rupture*

M. M. Murray, M.D.†; S. D. Martin, M.D.†; T. L. Martin, M.D.†; M. Spector, Ph.D.†

Investigation performed at the Department of Orthopaedic Surgery, Brigham and Women's Hospital, Boston, Massachusetts
*No benefits in any form have been received or will be received from a commercial party related directly or indirectly to the subject of this article. Funds were received in total or partial support of the research or clinical study presented in this article. The funding source was the Brigham Orthopedic Foundation.
†Department of Orthopaedic Surgery, Brigham and Women's Hospital, 75 Francis Street, Boston, Massachusetts 02115. E-mail address for M. M. Murray: mmmurray@partners.org.

The Journal of Bone & Joint Surgery. 2000; 82:1387-1387

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Abstract

Background: Four phases in the response to injury of the ruptured human anterior cruciate ligament are observed histologically; these include an inflammatory phase, an epiligamentous repair phase, a proliferative phase, and a remodeling phase. One objective of this study was to describe the histological changes that occur in the ruptured human anterior cruciate ligament during these phases. Myofibroblast-like cells that contain a-smooth muscle actin are present in the midsubstance of the intact human anterior cruciate ligament. A second objective of this study was to determine whether an increased number of myofibroblast-like cells is found in the midsubstance of the ruptured human anterior cruciate ligament because it was thought that those cells might be responsible in part for the retraction of the ruptured anterior cruciate ligament. In the early phase of this study, it was found that the number of myofibroblast-like cells in the midsubstance of the ruptured anterior cruciate ligament was actually decreased, and this hypothesis was abandoned. During the epiligamentous repair phase, synovial tissue was formed that covered the ends of the ruptured anterior cruciate ligament. Most of the synovial lining cells were myofibroblast-like cells that contained a-smooth muscle actin. The primary objective of this study was to determine the location and the characteristics of the a-smooth muscle actin-containing myofibroblast-like cells that appear in the human anterior cruciate ligament following rupture.

Methods: Twenty-three ruptured and ten intact human anterior cruciate ligaments were evaluated for cellularity, nuclear morphology, blood vessel density, and percentage of cells containing a contractile actin isoform, a-smooth muscle actin. The histological features of the synovial and epiligamentous tissues were also described.

Results: At no time after rupture was there evidence of tissue-bridging between the femoral and tibial remnants of the anterior cruciate ligament. The ruptured ligaments demonstrated a time-dependent histological response, which consisted of inflammatory cell infiltration up to three weeks, gradual epiligamentous repair and resynovialization between three and eight weeks, and neovascularization and an increase in cell number density between eight and twenty weeks. Compared with the intact ligaments, there was a decrease in the percentage of myofibroblast-like cells containing a-smooth muscle actin within the remnant of the ligament. However, many of the epiligamentous and synovial cells encapsulating the remnants contained a-smooth muscle actin.

Conclusions: After rupture, the human anterior cruciate ligament undergoes four histological phases, consisting of inflammation, epiligamentous regeneration, proliferation, and remodeling. The response to injury is similar to that reported in other dense connective tissues, with three exceptions: formation of an a-smooth muscle actin-expressing synovial cell layer on the surface of the ruptured ends, the lack of any tissue bridging the rupture site, and the presence of an epiligamentous reparative phase that lasts eight to twelve weeks. Other characteristics reported in healing dense connective tissue, such as fibroblast proliferation, expression of a-smooth muscle actin, and revascularization, also occur in the ruptured human anterior cruciate ligament.

Clinical Relevance: Unlike extra-articular ligaments that heal after injury, the human intra-articular anterior cruciate ligament forms a layer of synovial tissue over the ruptured surface, which may impede repair of the ligament. Moreover, a large number of cells in this synovial layer and in the epiligamentous tissue express the gene for a contractile actin isoform, a-smooth muscle actin, thus differentiating into myofibroblasts. These events may play a role in the retraction and lack of healing of the ruptured anterior cruciate ligament.

Figures in this Article

Introduction

Introduction | Materials and Methods | Results | Discussion | References

Ligaments that heal, such as the medial collateral ligament, do so by progressing through a series of inflammatory, proliferative, and remodeling phases^3,5, which result in the formation of a functional scar. In contrast, the anterior cruciate ligament does not appear to form a bridging scar after rupture^1,17,28,29, even when a primary repair has been performed^28,29. After rupture, the human anterior cruciate ligament appears to retract initially, with no evidence of healing^20,34. Surgical repair initially seemed to yield promising results, with one two-year study demonstrating a good or excellent outcome in twenty-five of thirty patients⁹. Five-year follow-up, however, demonstrated that 94 percent (thirty) of thirty-two patients had instability and 72 percent (twenty-three) had pain¹⁰. A similar study, of seventy athletes, demonstrated good results at an average of two and a half years after repair, with 93 percent (sixty-five) of the patients returning to their sport and none with unpredictable giving-way²¹. However, the longer-term follow-up of fifty-two patients revealed an overall failure rate of 17 percent (nine patients), with 42 percent (twenty-two) of the patients demonstrating laxity on clinical examination¹⁸. These results have since been corroborated by the long-term findings of other investigators^6,8,27,32. The poor wound-healing response of the anterior cruciate ligament has thus been noted both histologically in animal models and clinically in humans.

Why the anterior cruciate ligament does not heal after rupture is an important question that remains unanswered. In this study of histological changes in the ruptured anterior cruciate ligament, we sought to determine what cellular events occur that might impede satisfactory healing. We recently showed that myofibroblast-like cells that contain a-smooth muscle actin are present in the midsubstance of the intact human anterior cruciate ligament²⁵, and our initial objective was to determine whether an increased number of myofibroblast-like cells is formed in the midsubstance of the ligament after rupture and might be responsible for retraction of the ligament. However, in the initial phase of this study, we found that the number of myofibroblast-like cells in the midsubstance of the ligament was actually decreased, and our initial hypothesis was abandoned. Unexpected observations were then made that led to the formulation of a totally different hypothesis. The human anterior cruciate ligament (which is normally covered by synovial tissue) undergoes four phases in its response to rupture; these include inflammation, epiligamentous repair, proliferation, and remodeling. During the epiligamentous repair phase, synovial tissue was reformed and covered the ends of the ruptured ligament. Most of the synovial lining cells were myofibroblast-like cells that contained a-smooth muscle actin. These observations suggested that the early reformation of synovial tissue over the ends of the ligament combined with the formation of myofibroblast-like cells with contractile properties in this synovial tissue might be responsible in part for the retraction of the ligament and its poor healing.

The main objective of this study was to define the location and characteristics of the a-smooth muscle actin-containing myofibroblast-like cells that appear in the human anterior cruciate ligament following rupture.

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Fig. 1:: Schematic of the gross and histological appearance of the four phases of the healing response in the human anterior cruciate ligament.
A: The inflammatory phase, showing mop-ends of the remnants (a), disruption of the epiligament and synovial covering of the ligament (b), intimal hyperplasia of the vessels (c), and loss of the regular crimp structure near the site of injury (d).
B: The epiligamentous regeneration phase, involving a gradual recovering of the ligament remnant by vascularized, epiligamentous tissue and synovial tissue (e).
C: The proliferative phase, with revascularization of the remnant with groups of capillaries (f).
D: The remodeling and maturation phase, characterized by a decrease in cell number density and blood vessel density (g) and by retraction of the ligament remnant (h).

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Fig. 2:: Case 12. Photomicrograph of the epiligamentous tissue three weeks after rupture of the anterior cruciate ligament, demonstrating a high density of thin-walled blood vessels. Immunohistochemical analysis revealed a-smooth muscle actin where red designates a positive stain. No a-smooth muscle actin was demonstrated in control sections incubated with nonimmune mouse serum instead of the mouse monoclonal antibody to a-smooth muscle actin.

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Fig. 3:: Case 24. Photomicrograph of the epiligamentous tissue sixteen weeks after rupture of the anterior cruciate ligament, demonstrating vascular tissue with an overlying well defined synovial layer containing cells staining positive for a-smooth muscle actin (arrows). Immunohistochemical analysis revealed a-smooth muscle actin where red designates a positive stain. No a-smooth muscle actin was demonstrated in control sections incubated with nonimmune mouse serum instead of the mouse monoclonal antibody to a-smooth muscle actin.

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Fig. 4:: Case 16. Photomicrograph of the epiligamentous tissue six weeks after rupture of the anterior cruciate ligament, demonstrating vascular tissue with multiple cells staining positive for a-smooth muscle actin, which may be of vascular origin.

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Fig. 5:: Case 17. Area in the midsubstance of the remnant, showing several a-smooth muscle actin-containing fibroblasts displaying the red chromogen in the cytoplasm. The elongated cells are aligned along the long axis of the collagen bundles. No a-smooth muscle actin was demonstrated in control sections incubated with nonimmune mouse serum instead of the mouse monoclonal antibody to a-smooth muscle actin. BV = blood vessel.

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Fig. 6:: Histogram demonstrating the changes in cell number density near the site of injury as a function of time after complete rupture of the anterior cruciate ligament and comparison with the cell number density in the proximal part of the intact anterior cruciate ligament. Error bars represent the standard error of the mean.

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Fig. 7:: Histogram demonstrating the changes in blood vessel density near the site of injury as a function of time after complete rupture of the anterior cruciate ligament and comparison with the blood vessel density in the proximal part of the intact anterior cruciate ligament. Error bars represent the standard error of the mean.

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Table I: Patient Demographics for Intact and Ruptured Anterior Cruciate Ligaments

*The time from rupture was designated to the nearest week, or to the nearest four-week period for the later specimens.

Case	Age (yrs.)	Gender	Time from Rupture* (wks.)
Intact ligaments
1	61	M
2	65	F
3	65	F
4	83	F
5	73	F
6	75	F
7	62	F
8	65	M
9	65	F
10	71	M
Ruptured ligaments
11	34	M	1
12	25	M	3
13	28	F	3
14	45	F	4
15	24	M	6
16	24	F	6
17	14	F	8
18	20	F	8
19	24	M	8
20	29	M	8
21	45	M	12
22	42	M	16
23	41	M	16
24	24	M	16
25	31	M	16
26	46	M	20
27	34	M	20
28	30	M	52
29	22	M	64
30	21	M	104
31	20	M	104
32	44	F	104
33	36	M	156

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Table I: Patient Demographics for Intact and Ruptured Anterior Cruciate Ligaments

*The time from rupture was designated to the nearest week, or to the nearest four-week period for the later specimens.

Case	Age (yrs.)	Gender	Time from Rupture* (wks.)
Intact ligaments
1	61	M
2	65	F
3	65	F
4	83	F
5	73	F
6	75	F
7	62	F
8	65	M
9	65	F
10	71	M
Ruptured ligaments
11	34	M	1
12	25	M	3
13	28	F	3
14	45	F	4
15	24	M	6
16	24	F	6
17	14	F	8
18	20	F	8
19	24	M	8
20	29	M	8
21	45	M	12
22	42	M	16
23	41	M	16
24	24	M	16
25	31	M	16
26	46	M	20
27	34	M	20
28	30	M	52
29	22	M	64
30	21	M	104
31	20	M	104
32	44	F	104
33	36	M	156

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Table II: Histomorphometric Measurements of the Intact and Ruptured Human Anterior Cruciate Ligaments*

*The values are given as the mean and the standard error of the mean. a-SMA = a-smooth muscle actin.

Intact, or No. of Weeks After Rupture	Proximal Edge	1 mm from Edge	2 mm from Edge	4 mm from Edge	6 mm from Edge
Intact ligaments
Cell density (no./mm²)	701 ± 120	525 ± 108	539 ± 91	294 ± 39	265 ± 37
Nuclear aspect ratio	6.1 ± 0.9	4.5 ± 0.8	4.3 ± 0.6	3.6 ± 0.6	2.4 ± 0.5
Blood vessel density (no./mm)	1.5 ± 0.16	1.2 ± 0.2	1.0 ± 0.2	0.60 ± 0.12	0.24 ± 0.03
Cells positive for a-SMA (percent)	4.7 ± 1.0	7.3 ± 1.7	10.7 ± 3.0	15 ± 3.9	17 ± 4.3
No. of specimens	10	10	10	10	10
1 to 6 weeks
Cell density (no./mm²)	614 ± 249	476 ± 267	420 ± 210	254 ± 48	231 ± 30
Nuclear aspect ratio	4.5 ± 1.0	3.9 ± 0.8	3.7 ± 0.9	4.2 ± 0.7	4.3 ± 1.2
Blood vessel density (no./mm)	4 ± 3.3	2.9 ± 2.6	5.0 ± 2.9	2.0 ± 1.2	0.8 ± 0.2
Cells positive for a-SMA (percent)	2.3 ± 1.4	1.9 ± 1.1	1.0 ± 0.3	0.8 ± 0.31	0.4 ± 0.12
No. of specimens	6	6	6	6	6
8 to 12 weeks
Cell density (no./mm²)	1541 ± 451	1272 ± 363	965 ± 249	701 ± 162	497 ± 151
Nuclear aspect ratio	6.2 ± 1.0	4.3 ± 1.0	3.8 ± 1.0	2.9 ± 1.0	4.1 ± 1.3
Blood vessel density (no./mm)	5.1 ± 3.1	4.0 ± 2.6	3.0 ± 2.1	2.2 ± 1.0	2.1 ± 1.0
Cells positive for a-SMA (percent)	1.3 ± 0.8	1.3 ± 0.3	1.1 ± 0.3	0.5 ± 0.3	0.3 ± 0.2
No. of specimens	5	5	5	5	5
16 to 20 weeks
Cell density (no./mm²)	2244 ± 526	1522 ± 285	1037 ± 280	833 ± 312	1009 ± 437
Nuclear aspect ratio	5.4 ± 1.0	4.8 ± 0.2	4.6 ± 0.5	5.3 ± 1.2	3.8 ± 1.3
Blood vessel density (no./mm)	13.3 ± 4.9	4.0 ± 1.3	5.2 ± 2.0	2.9 ± 1.6	3.3 ± 2.0
Cells positive for a-SMA (percent)	0.6 ± 0.3	0.4 ± 0.2	0.3 ± 0.2	0.3 ± 0.3	1.2 ± 0.7
No. of specimens	6	6	6	6	6
52 to 104 weeks
Cell density (no./mm²)	559 ± 115	601 ± 204	718 ± 241	590 ± 46	546 ± 45
Nuclear aspect ratio	3.7 ± 0.6	4.0 ± 0.9	4.2 ± 0.5	3.3 ± 1.1	3.7 ± 0.5
Blood vessel density (no./mm)	2.1 ± 2.0	1.5 ± 1.3	1.2 ± 0.7	1.6 ± 0.8	1.3 ± 0.6
Cells positive for a-SMA (percent)	0.5 ± 0.3	0.2 ± 0.2	0.2 ± 0.1	0.5 ± 0.3	1.1 ± 0.9
No. of specimens	6	6	6	6	6

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Table II: Histomorphometric Measurements of the Intact and Ruptured Human Anterior Cruciate Ligaments*

*The values are given as the mean and the standard error of the mean. a-SMA = a-smooth muscle actin.

Intact, or No. of Weeks After Rupture	Proximal Edge	1 mm from Edge	2 mm from Edge	4 mm from Edge	6 mm from Edge
Intact ligaments
Cell density (no./mm²)	701 ± 120	525 ± 108	539 ± 91	294 ± 39	265 ± 37
Nuclear aspect ratio	6.1 ± 0.9	4.5 ± 0.8	4.3 ± 0.6	3.6 ± 0.6	2.4 ± 0.5
Blood vessel density (no./mm)	1.5 ± 0.16	1.2 ± 0.2	1.0 ± 0.2	0.60 ± 0.12	0.24 ± 0.03
Cells positive for a-SMA (percent)	4.7 ± 1.0	7.3 ± 1.7	10.7 ± 3.0	15 ± 3.9	17 ± 4.3
No. of specimens	10	10	10	10	10
1 to 6 weeks
Cell density (no./mm²)	614 ± 249	476 ± 267	420 ± 210	254 ± 48	231 ± 30
Nuclear aspect ratio	4.5 ± 1.0	3.9 ± 0.8	3.7 ± 0.9	4.2 ± 0.7	4.3 ± 1.2
Blood vessel density (no./mm)	4 ± 3.3	2.9 ± 2.6	5.0 ± 2.9	2.0 ± 1.2	0.8 ± 0.2
Cells positive for a-SMA (percent)	2.3 ± 1.4	1.9 ± 1.1	1.0 ± 0.3	0.8 ± 0.31	0.4 ± 0.12
No. of specimens	6	6	6	6	6
8 to 12 weeks
Cell density (no./mm²)	1541 ± 451	1272 ± 363	965 ± 249	701 ± 162	497 ± 151
Nuclear aspect ratio	6.2 ± 1.0	4.3 ± 1.0	3.8 ± 1.0	2.9 ± 1.0	4.1 ± 1.3
Blood vessel density (no./mm)	5.1 ± 3.1	4.0 ± 2.6	3.0 ± 2.1	2.2 ± 1.0	2.1 ± 1.0
Cells positive for a-SMA (percent)	1.3 ± 0.8	1.3 ± 0.3	1.1 ± 0.3	0.5 ± 0.3	0.3 ± 0.2
No. of specimens	5	5	5	5	5
16 to 20 weeks
Cell density (no./mm²)	2244 ± 526	1522 ± 285	1037 ± 280	833 ± 312	1009 ± 437
Nuclear aspect ratio	5.4 ± 1.0	4.8 ± 0.2	4.6 ± 0.5	5.3 ± 1.2	3.8 ± 1.3
Blood vessel density (no./mm)	13.3 ± 4.9	4.0 ± 1.3	5.2 ± 2.0	2.9 ± 1.6	3.3 ± 2.0
Cells positive for a-SMA (percent)	0.6 ± 0.3	0.4 ± 0.2	0.3 ± 0.2	0.3 ± 0.3	1.2 ± 0.7
No. of specimens	6	6	6	6	6
52 to 104 weeks
Cell density (no./mm²)	559 ± 115	601 ± 204	718 ± 241	590 ± 46	546 ± 45
Nuclear aspect ratio	3.7 ± 0.6	4.0 ± 0.9	4.2 ± 0.5	3.3 ± 1.1	3.7 ± 0.5
Blood vessel density (no./mm)	2.1 ± 2.0	1.5 ± 1.3	1.2 ± 0.7	1.6 ± 0.8	1.3 ± 0.6
Cells positive for a-SMA (percent)	0.5 ± 0.3	0.2 ± 0.2	0.2 ± 0.1	0.5 ± 0.3	1.1 ± 0.9
No. of specimens	6	6	6	6	6

Materials and Methods

Introduction | Materials and Methods | Results | Discussion | References

Twenty-three ruptured anterior cruciate ligament remnants were obtained from seventeen male patients and six female patients (age range, fourteen to forty-six years; average age, thirty-one years) undergoing reconstruction of the anterior cruciate ligament (Table I). The ruptured ligaments were obtained from ten days to three years after rupture. Ten intact ligaments (controls) were obtained from three men and seven women (age range, sixty-one to eighty-three years; average age, sixty-nine years) undergoing total knee arthroplasty because of degenerative joint disease (Table I). The intact ligaments were resected from their insertion sites with a scalpel by the surgeon. The majority of the ruptured ligaments were gently lifted from the posterior cruciate ligament, transected at their tibial attachment, and removed arthroscopically by the surgeon. Ruptured ligaments retrieved at ten days to three weeks were removed at the time of open reconstruction for the treatment of multiple ligament injury. Our Institutional Review Board approved the use of the human ligament tissue.

Histological and Immunohistochemical Analysis

The ligaments were marked with a suture at the site of tibial transection and fixed in neutral buffered formalin for one week. After fixation, specimens were embedded longitudinally in paraffin, and seven-micrometer-thick longitudinal sections were cut and fixed on glass slides. Representative sections from each ligament were stained with hematoxylin and eosin, and immunohistochemical studies were carried out with use of a mouse monoclonal antibody to a-smooth muscle actin (Sigma Chemical, St. Louis, Missouri). In the immunohistochemical procedure, deparaffinized, hydrated slides were digested with 0.1 percent trypsin (Sigma Chemical) for twenty minutes. Endogenous peroxidase was quenched with 3 percent hydrogen peroxide for five minutes. Nonspecific sites were blocked with use of 20 percent goat serum for thirty minutes. The sections were then incubated with the mouse monoclonal antibody to a-smooth muscle actin for one hour at room temperature. A negative control section on each microscope slide was incubated with nonimmune mouse serum diluted to the same protein content, instead of with the a-smooth muscle actin antibody, to monitor for nonspecific staining. The sections were then incubated with a biotinylated goat anti-mouse IgG secondary antibody for thirty minutes followed by thirty minutes of incubation with affinity purified avidin. The labeling was developed with use of the AEC chromogen kit (Sigma Chemical) for ten minutes. Counterstaining with Mayer hematoxylin for twenty minutes was followed by a twenty-minute tap-water rinse, and a coverslip was applied with glycerol gelatin.

Method of Evaluation

Histological slides were examined with use of a Vanox-T AH-2 microscope (Olympus, Tokyo, Japan) with normal and polarized light. For the histomorphometric measurements, the intact ligaments were evaluated at the site of transection from the femoral attachment and at one, two, four, and six millimeters distal to the transection site. These analyses did not include the insertion of the ligament into bone. The ruptured ligaments were evaluated at the ruptured end and at one, two, four, and six millimeters distal to the site of the rupture (toward the tibial insertion). At each location, three 0.1-square-millimeter areas were evaluated by determining the total cell number density, the nuclear morphology, the blood vessel density, and the percentage of cells that were positive for the a-smooth muscle actin isoform. The nuclear morphology was classified on the basis of the nuclear shape, which was fusiform, ovoid, or spheroid. Fibroblasts with nuclei with a ratio of length divided by width of greater than ten were classified as fusiform, those with a ratio of between five and ten were classified as ovoid, and those with a ratio of less than five were classified as spheroid. Between twenty and 230 cells were counted at each of the three areas. At each location, the total number of cells was counted and divided by the area of analysis in order to yield the cell number density, or cellularity. The total number of blood vessels crossing the section at each location was divided by the width of the section at each location to determine the blood vessel density for each location.

Smooth muscle cells surrounding vessels were used as internal positive controls for determination of a-smooth muscle actin-positive cells. Positive cells were those that demonstrated chromogen intensity similar to that seen in the smooth muscle cells on the same microscope slide and that had markedly more intense stain than the perivascular cells on the negative control section. Any cell with a questionable intensity of stain (for example, a light pink tint) was not counted as positive. The a-smooth muscle actin-positive cell density was reported as the number of stained cells divided by the area of analysis, and the percentage of a-smooth muscle actin-positive cells was determined by dividing the number of stained cells by the total number of cells in a particular histological zone.

Results

Introduction | Materials and Methods | Results | Discussion | References

Gross and Histological Observations

After the complete rupture of the human anterior cruciate ligament, four progressive phases of healing response were seen.

Phase I: Inflammation

Within the first few weeks after the rupture, the synovial fluid that was encountered on entering the joint was rust-colored and was easily suctioned from the knee. No blood clots were found within the knee joint. The remnants of the anterior cruciate ligaments were swollen and edematous, and the synovial and epiligamentous tissue was grossly disrupted. Blood clot was seen covering part of the ligament remnants, but no connection between the femoral and tibial ends was visible grossly. Near the site of the rupture, the ligament ends were composed of friable, stringy tissue previously described as "mop-ends" (Fig. 1, A).

Histologically, the ligament remnants retrieved in this time-period were populated by fibroblasts, polymorphonuclear neutrophils, lymphocytes, and macrophages. The inflammatory cells were found in greatest concentration around blood vessels near the site of the injury. Macrophages appeared to be actively phagocytosing cell and tissue debris.

Arterioles near the site of the injury were dilated, with intimal hyperplasia associated with dramatic proliferation of the smooth muscle cells and thickening of the walls of the arterioles. Venules were dilated, with less hyperplasia of the smooth muscle cells. Capillaries were congested, with rouleaux and thrombus formation in their lumens.

Phase II: Epiligamentous Regeneration

Between three and eight weeks after rupture, gradual growth of epiligamentous and synovial tissue was noted over the ruptured end of the ligament, giving it a smoother, mushroom-like appearance (Fig. 1, B), different from the mop-ends seen in the earlier specimens. No tissue bridged the gap between the proximal and distal segments, although several of the distal remnants were adherent to the sheath of the intact posterior cruciate ligament.

Histologically, the epiligamentous repair phase was characterized by an unchanged cell number density and blood vessel density in the midsubstance of the ligament. After the initial influx of inflammatory cells and removal of cell and tissue debris seen in the inflammatory phase, the number of inflammatory cells decreased and fibroblasts became the dominant cell type. The cell number density of fibroblasts was similar to that seen in the intact ligament, and the remaining blood vessels displayed essentially normal morphology, with little intimal hyperplasia. No neovascularization was noted within the ligament fascicles.

Most of the changes occurred in the epiligament, which displayed an increase in cell number density and blood vessel density. The vascular epiligamentous tissue extended over the end of the ruptured ligament, encapsulating the mop-ends (Fig. 2). Thickening of the epiligament and fibroblast proliferation were seen to occur during this time-period. A synovial cell layer was found extending over the surface of the ruptured ligament.

Phase III: Proliferation

By eight weeks, the distal remnant of the anterior cruciate ligament was completely encapsulated by a synovial sheath, and few remaining mop-ends were seen grossly (Fig. 1, C). No tissue was visible between the proximal and distal ligament remnants. Several of the distal remnants were noted to be adherent to the periligamentous tissue of the posterior cruciate ligament.

Histologically, the period between eight and twenty weeks after rupture was characterized by increasing cell number density and blood vessel density in and among the collagen fascicles of the ligament remnant. Fibroblasts were the predominant cell type. The entire remnant became increasingly cellular, with a peak cell number density at sixteen to twenty weeks. The orientation of the fibroblasts remained disorganized, with few cell nuclei with longitudinal axes parallel to that of the ligament. Vascular endothelial capillary buds were seen during this phase, and loops from anastomoses of proximal sprouts were noted to form a diffuse network of immature capillaries (Fig. 1, C).

Between six and twenty weeks, a continuous layer of synovial tissue had formed over the ends of the ruptured ligament (Fig. 3). The most dramatic - and unexpected - observation was the abundance of a-smooth muscle actin-containing cells making up the synovial layer in selected regions (Fig. 3). Many a-smooth muscle actin-expressing cells in the synovial layer were clearly separate from vascular smooth muscle cells and pericytes in the underlying epiligamentous tissue (Fig. 3). No staining for a-smooth muscle actin was demonstrated in control sections incubated with nonimmune mouse serum instead of mouse monoclonal antibody to a-smooth muscle actin. In other cases, however, such a distinction was not possible as the synovial layer merged with a highly vascular epiligament, forming a syncytium of a-smooth muscle actin-containing cells (Fig. 4). In specimens obtained in this time-period, immunohistochemical analysis also revealed a-smooth muscle actin-containing fibroblasts distributed throughout the midsubstance of the remnant (Fig. 5), albeit in relatively low percentages (as presented below and in Table II).

Phase IV: Remodeling and Maturation

Between one and two years after ligament rupture, remodeling and maturation of the ligament remnant were seen. The ligament ends were dense and white, with little fatty synovial tissue overlying them (Fig. 1, D). No tissue was noted to connect the two ends of the ligament.

Histologically, the fibroblast nuclei were fusiform with the long axis of the nucleus aligned with the longitudinal axis of the ligament. There was decreased blood vessel density within the ligament remnant. The epiligamentous tissue continued to decrease in thickness; however, the synovial sheath persisted. A more axial alignment of the collagen fascicles was seen. The cell number density decreased to a level similar to that seen in the intact human anterior cruciate ligament.

Histomorphometric Results

The histomorphometric results for the ligaments at each of the time-points are provided in Table II. The evaluation of the percentage of a-smooth muscle actin-positive cells did not include the synovial or epiligamentous tissue when the distinction of vascular and nonvascular cells could not be made confidently.

In the control group of intact anterior cruciate ligaments, there was a decrease in cell number density and vascularity proceeding from proximal to distal and an increase in the sphericity of the cell nuclei and in the percentage of a-smooth muscle actin-positive cells.

Two-factor analysis of variance demonstrated that the cell number density in the ruptured anterior cruciate ligaments was significantly affected by the location in the ligament remnant and by the time after the rupture. The cell number density was highest near the site of injury at all time-points up to one year after the injury. This cellularity increased significantly, to a maximum at sixteen to twenty weeks (Bonferroni-Dunn post hoc testing, p < 0.005), and then decreased between twenty and fifty-two weeks after the injury (Bonferroni-Dunn post hoc testing, p < 0.005) (Fig. 6). With the number of ligaments available, age and gender were not found to significantly affect cell number density (two-factor analysis of variance, p > 0.80 and p < 0.40, respectively).

Two-factor analysis of variance demonstrated that the morphology of the cell nuclei was significantly affected by the location in the ligament remnant (p < 0.003) but not by the time after the injury (p < 0.40) or by age (p < 0.70), with the numbers available. The effect of gender on this parameter was close (p < 0.06) to meeting our criterion for significance (p < 0.05). The proximal part of the ligament remnant was found to have cells with a higher nuclear aspect ratio when compared with cells in the more distal part of the remnant (Bonferroni-Dunn post hoc testing, p < 0.0005). This pattern was also observed in the intact ligaments.

Two-factor analysis of variance showed the blood vessel density to be significantly affected by the time after the injury. The blood vessel density reached its highest value at sixteen to twenty weeks (Bonferroni-Dunn post hoc testing, p < 0.003) and decreased after that time-point (Fig. 7). The blood vessel density appeared to decrease with the distance from the ruptured end. However, the effect of location on blood vessel density (p < 0.09) did not reach significance. With the numbers available, two-way analysis of variance showed no significant effect of age (p < 0.30) or gender (p > 0.25) on blood vessel density.

Cells that stained positively for the a-smooth muscle actin isoform were present throughout the intact and ruptured anterior cruciate ligaments. The time after the injury was not found to have a significant effect on staining for a-smooth muscle actin (p < 0.30). However, compared with the intact ligaments, the ruptured ligaments had a smaller percentage of cells that stained positive for a-smooth muscle actin in their midsubstance. Neither the location in the ligament (p < 0.90) nor the age of the patient (p < 0.61) was found to have a significant effect on the percentage of cells staining positive for a-smooth muscle actin. Gender was found to have a significant effect, with women having a greater percentage of cells staining positive for the isoform than men (p < 0.002).

Discussion

Introduction | Materials and Methods | Results | Discussion | References

The human anterior cruciate ligament undergoes four histological phases after rupture: an inflammatory phase, an epiligamentous reparative phase, a proliferative phase, and a remodeling phase. The response to injury is similar to that reported in other dense connective tissues, with three important exceptions that are likely interrelated: (1) the formation of an a-smooth muscle actin-expressing synovial cell layer on the surface of the ruptured ends, (2) the lack of any tissue bridging the rupture site, and (3) the presence of an epiligamentous reparative phase that lasts eight to twelve weeks. Other reported characteristics of the healing of dense connective tissue, such as fibroblast proliferation, expression of a-smooth muscle actin, and angiogenesis, all occur in the human anterior cruciate ligament.

The proliferative fibroblast response, while similar to that seen in tissues that heal, occurs at a later time-point after ruptures of the anterior cruciate ligament. For example, in the medial collateral ligaments of rabbits, this proliferative response begins in the second week after injury, when the fibroblasts are noted to be synthesizing extracellular matrix actively¹¹.

Synovial Cell Expression of a-Smooth Muscle Actin and Contractile Behavior

The percentage of nonvascular a-smooth muscle actin-containing cells distributed throughout the midsubstance of the ruptured remnants was unremarkable. However, a large number of a-smooth muscle actin-containing cells appeared in the synovial tissue (Fig. 3), which may be responsible in part for the retraction of the ends of the anterior cruciate ligament. Several prior studies have implicated synovial cells in contraction.

In studies of the healing of the knee meniscus¹³ and the temporomandibular joint disc³¹, it was proposed that synovial tissue was the source of the myofibroblasts that contributed to the reparative process through their contraction. A recent investigation²² showed that a small but substantial percentage (14 ± 3 percent) of synovial fibroblasts isolated from patients with rheumatoid arthritis or osteoarthritis expressed a-smooth muscle actin, as determined by immunolocalization. Of note, however, was that there was no remarkable staining for a-smooth muscle actin in the synovial cells in these samples in situ. The investigators posited that the lack of staining in some samples may have been due to the low numbers of fibroblasts in the samples and the effects of certain cytokines in down-regulating a-smooth muscle actin expression.

The results of this study suggested that a contiguous layer of a-smooth muscle actin-containing cells around a tissue, rather than individual myofibroblasts distributed through it, may be responsible for contraction (that is, retraction of the remnants of the anterior cruciate ligament). This "picture frame" model of tissue contraction¹⁴ was supported by the in vitro finding of contraction of porous collagen-glycosaminoglycan analogs of extracellular matrix by a multiple layer of bovine tendon-derived myofibroblasts on the surface of the specimen³³. Evidence of circumferential contraction by the myofibroblast layer included the finding that the peripheral pores were more constricted than the internal pores.

Absence of Tissue Bridging the Rupture Gap

A notable finding in this study was the lack of any tissue in the gap between the ligament remnants. In extra-articular tissues that heal, a fibrin clot forms that is invaded by fibroblasts and is gradually replaced by collagen fibers. This has been demonstrated to be instrumental in the healing process in both tendon^7,30 and the medial collateral ligament¹¹. However, it has been demonstrated that fibrin clot does not form in the intraarticular milieu, probably because of the presence of fibrinolytic enzymes in synovial fluid^2,15. An experimental study of hemarthroses (induced by drilling across the joint) in canine knees demonstrated no clot formation from five minutes to four hours after the injury; only bloody fluid was found within the joint¹⁵. In the present study, only one of the ruptured human anterior cruciate ligaments demonstrated any fibrin clot bridging adjacent fascicles of the tibial remnant, and none of the ruptured ligaments had any clot or tissue bridging the proximal and distal remnants or bridging a gap of greater than 700 micrometers within a remnant. As the early specimens were obtained with an open technique, it is possible that the blood clot seen on the remnants had formed at the time of surgery, after the synovial fluid had been removed from the joint. In the knees that were operated on in the first ten to twenty-one days after the injury, the hemarthrosis had already been lysed to a viscous liquid incapable of holding the ruptured ligament remnants together. One reason, then, for the failure of the human anterior cruciate ligament to heal is the lack of a bridging scaffold between the ruptured ligament ends that persists until the proliferative cell response occurs.

Epiligamentous Regeneration

The presence of an epiligamentous reparative phase distinguishes the ruptured human anterior cruciate ligament from connective tissues that heal. In the medial collateral ligaments of rabbits, the inflammatory phase is followed immediately by the start of fibroblast proliferation during the second week after injury¹¹. However, an epitendinous reparative phase was seen in lapine tendon-healing when the synovial sheath had been violated; thickening and fibroblast proliferation of the epitenon were noted to occur before any proliferative response of the endotenon cells¹². If the synovial sheath had not been violated, an epitenon and endotenon demonstrated increases in cell proliferation simultaneously²³.

The presence of the epiligamentous reparative phase also reconciles the findings in this study with previous reports of failure of anterior cruciate ligament cells to respond to rupture. In one of the two previous histological studies of the human anterior cruciate ligament after injury, biopsies performed up to forty-five days after rupture demonstrated cell death and minimal proliferation²⁶. This finding is consistent with the results seen in our six to eight-week specimens - that is, the fibroblast remained the predominant cell type but the cell number density remained low. A substantial cell response within the ligament remnant was not seen in our study until more than eighty-four days after the rupture.

The presence of the epiligamentous reparative phase is also important with regard to analysis of the results of primary repair or augmentation techniques. These procedures may have different results depending on the timing of the repair after the injury. Repair done in the first few weeks after an injury may result in filling of the gap with the proliferative epiligamentous vascular tissue that is active at that time. Repair performed months after an injury, when the endoligamentous tissue is proliferating, may result in a different mode of repair. Additional work is needed to determine if there is any difference in the repair strength or histological findings between primary repair performed late and that performed early.

Neovascularization

The neovascularization of the anterior cruciate ligament remnant, along with the fibroblastic proliferative response, was similar to that reported in studies of other healing soft tissues, including the medial collateral ligament³, tendon^7,30, and the periphery of the meniscus¹⁹. Proliferation of perivascular smooth muscle cells was most pronounced in the ten-day and two-week anterior cruciate ligament remnants, and intimal hyperplasia was seen up to three weeks after rupture.

Lack of neovascularization after injury has been mentioned as one of the factors responsible for the failure of other avascular or poorly vascularized tissues (for example, articular cartilage) to heal. Techniques involving perforation of the subchondral bone in an attempt to stimulate vascular invasion of the lesion, initially performed in rabbits²⁴, are currently in widespread clinical use. The lack of neovascularization is also thought to be one of the primary reasons why the inner zone of the meniscus does not heal after injury. Tears in the peripheral, vascularized zone of the meniscus are capable of healing with repair, while those in the inner, avascular zone are incapable of mounting a reparative response¹⁹. Other studies have shown experimentally that, after sharp incisional injury in the avascular meniscus, no gross or histological evidence of repair is seen^4,13,16. Thus, the response that we observed in the human anterior cruciate ligament is markedly dissimilar from that seen in other intra-articular tissues that fail to heal in that the human anterior cruciate ligament is capable of neovascularization of the remnants after complete rupture.

The fact that fibroblast proliferation and angiogenesis occur within the human anterior cruciate ligament remnant is also critical to the development of future methods of facilitating healing of that ligament. The pronounced neovascularization and cell proliferation suggest that the harnessing of these responses, and their extension into the gap between the ruptured ligament ends, may provide a cell biology-based method of repair of the anterior cruciate ligament, with the ligament remnants providing the cellular constituents of the repair tissue.

In summary, the biology of the response of the human anterior cruciate ligament to complete rupture is a complex process. It differs from that of tissues that successfully heal in that there is an epiligamentous reparative phase and a lack of any bridging scar formation. Moreover, the formation of a synovial layer, comprising cells with a contractile actin isoform, over the epiligamentous tissue may explain in part the retraction of the remnants that disfavors a reparative bridging tissue. The anterior cruciate ligament is different from other intra-articular tissues that fail to heal (for example, articular cartilage) in that it exhibits proliferative fibroblastic and angiogenic responses to rupture. A deeper understanding of the biology of the response of the anterior cruciate ligament to injury is likely to lead to new approaches to facilitate regeneration of this important musculoskeletal tissue.

Note: The authors acknowledge the support of the Brigham Orthopedic Foundation. They also thank Sandra Zapatka-Taylor for her assistance with the preparation of the histological specimens as well as Dr. Arthur Boland, Dr. Charles Brown, Dr. Mark Steiner, and Dr. Bertram Zarins for their help in obtaining the tissue for study.

References

Introduction | Materials and Methods | Results | Discussion | References

Amiel, D.; Kuiper, S.; and Akeson, W. H.: Cruciate ligaments. Response to injury. In Knee Ligaments. Structure, Function, Injury, and Repair, pp. 365-377. Edited by D. M. Daniel, W. H. Akeson, and J. J. O'Connor. New York, Raven Press, 1990.

Andersen, R. B., and Gormsen, J.: Fibrin dissolution in synovial fluid. Acta Rheumat. Scandinavica, 16: 319-333, 1970.

Andriacchi, T.; Sabiston, P.; DeHaven, K.; Dahners, L.; Woo, S. L-Y.; Frank, C. B.; Oakes, B.; Brand, R.; and Lewis, J.: Ligament: injury and repair. In Injury and Repair of the Musculoskeletal Soft Tissues: Workshop, Savannah, Georgia, June 1987, pp. 103-128. Edited by S. L-Y. Woo and J. A. Buckwalter. Park Ridge, Illinois, American Academy of Orthopaedic Surgeons, 1988.

Arnoczky, S. P.; Rubin, R. M.; and Marshall, J. L.: Microvasculature of the cruciate ligaments and its response to injury. An experimental study in dogs. J. Bone and Joint Surg., 61-A: 1221-1229, Dec. 1979.

Arnoczky, S. P.: Physiologic principles of ligament injuries and healing. In Ligament and Extensor Mechanism Injuries of the Knee: Diagnosis and Treatment, pp. 67-81. St. Louis, C. V. Mosby, 1991.

Balkfors, B.: The course of knee ligament injuries. Acta Orthop. Scandinavica, Supplementum 198, 1982.

Buck, R. C.: Regeneration of tendon. J. Pathol. and Bacteriol., 66: 1-18, 1953.

Engebretsen, L.; Benum, P.; and Sundalsvoll, S.: Primary suture of the anterior cruciate ligament. A 6-year follow-up of 74 cases. Acta Orthop. Scandinavica, 60: 561-564, 1989.

Feagin, J. A.; Abbott, H. G.; and Roukous, J. A.: The isolated tear of the anterior cruciate ligament. In Proceedings of the American Academy of Orthopaedic Surgeons. J. Bone and Joint Surg., 54-A: 1340, Sept. 1972.

Feagin, J. A., Jr., and Curl, W. W.: Isolated tear of the anterior cruciate ligament: 5-year follow-up study. Am. J. Sports Med., 4: 95-100, 1976.

Frank, C.; Schachar, N.; and Dittrich, D.: Natural history of healing in the repaired medial collateral ligament. J. Orthop. Res., 1: 179-188, 1983.

Gelberman, R.; Goldberg, V.; An, K.-N.; and Banes, A.: Tendon. In Injury and Repair of the Musculoskeletal Soft Tissues: Workshop, Savannah, Georgia, June 1987, pp. 1-40. Edited by S. L-Y. Woo and J. A. Buckwalter. Park Ridge, Illinois, American Academy of Orthopaedic Surgeons, 1988.

Ghadially, F. N.; Wedge, J. H.; and Lalonde, J. M.: Experimental methods of repairing injured menisci. J. Bone and Joint Surg., 68-B(1): 106-110, 1986.

Grillo, H. C.; Watts, G. T.; and Gross, J.: Studies in wound healing: I. Contraction and the wound contents. Ann. Surg., 148: 145-152, 1958.

Harrold, A. J.: The defect of blood coagulation in joints. J. Clin. Pathol., 14: 305-308, 1961.

Heatley, F. W.: The meniscus - can it be repaired? An experimental investigation in rabbits. J. Bone and Joint Surg., 62-B(3): 397-402, 1980.

Hefti, F. L.; Kress, A.; Fasel, J.; and Morscher, E. W.: Healing of the transected anterior cruciate ligament in the rabbit. J. Bone and Joint Surg., 73-A: 373-383, March 1991.

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Kohn, D.: Arthroscopy in acute injuries of anterior cruciate-deficient knees: fresh and old intraarticular lesions. Arthroscopy, 2: 98-102, 1986.

Marshall, J. L.; Warren, R. F.; and Wickiewicz, T. L.: Primary surgical treatment of anterior cruciate ligament lesions. Am. J. Sports Med., 10: 103-107, 1982.

Mattey, D. L.; Dawes, P. T.; Nixon, N. B.; and Slater, H.: Transforming growth factor beta 1 and interleukin 4 induced alpha smooth muscle actin expression and myofibroblast-like differentiation in human synovial fibroblasts in vitro: modulation by basic fibroblast growth factor. Ann. Rheumat. Dis., 56: 426-431, 1997.

Matthews, P., and Richards, H.: Factors in the adherence of flexor tendon after repair: an experimental study in the rabbit. J. Bone and Joint Surg., 58-B(2): 230-236, 1976.

Mitchell, N., and Shepard, N.: The resurfacing of adult rabbit articular cartilage by multiple perforations through the subchondral bone. J. Bone and Joint Surg., 58-A: 230-233, March 1976.

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Neurath, M. F.; Printz, H.; and Stofft, E.: Cellular ultrastructure of the ruptured anterior cruciate ligament. A transmission electron microscopic and immunohistochemical study in 55 cases. Acta Orthop. Scandinavica, 65: 71-76, 1994.

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Table I: Patient Demographics for Intact and Ruptured Anterior Cruciate Ligaments

*The time from rupture was designated to the nearest week, or to the nearest four-week period for the later specimens.

Case	Age (yrs.)	Gender	Time from Rupture* (wks.)
Intact ligaments
1	61	M
2	65	F
3	65	F
4	83	F
5	73	F
6	75	F
7	62	F
8	65	M
9	65	F
10	71	M
Ruptured ligaments
11	34	M	1
12	25	M	3
13	28	F	3
14	45	F	4
15	24	M	6
16	24	F	6
17	14	F	8
18	20	F	8
19	24	M	8
20	29	M	8
21	45	M	12
22	42	M	16
23	41	M	16
24	24	M	16
25	31	M	16
26	46	M	20
27	34	M	20
28	30	M	52
29	22	M	64
30	21	M	104
31	20	M	104
32	44	F	104
33	36	M	156

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Table I: Patient Demographics for Intact and Ruptured Anterior Cruciate Ligaments

*The time from rupture was designated to the nearest week, or to the nearest four-week period for the later specimens.

Case	Age (yrs.)	Gender	Time from Rupture* (wks.)
Intact ligaments
1	61	M
2	65	F
3	65	F
4	83	F
5	73	F
6	75	F
7	62	F
8	65	M
9	65	F
10	71	M
Ruptured ligaments
11	34	M	1
12	25	M	3
13	28	F	3
14	45	F	4
15	24	M	6
16	24	F	6
17	14	F	8
18	20	F	8
19	24	M	8
20	29	M	8
21	45	M	12
22	42	M	16
23	41	M	16
24	24	M	16
25	31	M	16
26	46	M	20
27	34	M	20
28	30	M	52
29	22	M	64
30	21	M	104
31	20	M	104
32	44	F	104
33	36	M	156

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Table II: Histomorphometric Measurements of the Intact and Ruptured Human Anterior Cruciate Ligaments*

*The values are given as the mean and the standard error of the mean. a-SMA = a-smooth muscle actin.

Intact, or No. of Weeks After Rupture	Proximal Edge	1 mm from Edge	2 mm from Edge	4 mm from Edge	6 mm from Edge
Intact ligaments
Cell density (no./mm²)	701 ± 120	525 ± 108	539 ± 91	294 ± 39	265 ± 37
Nuclear aspect ratio	6.1 ± 0.9	4.5 ± 0.8	4.3 ± 0.6	3.6 ± 0.6	2.4 ± 0.5
Blood vessel density (no./mm)	1.5 ± 0.16	1.2 ± 0.2	1.0 ± 0.2	0.60 ± 0.12	0.24 ± 0.03
Cells positive for a-SMA (percent)	4.7 ± 1.0	7.3 ± 1.7	10.7 ± 3.0	15 ± 3.9	17 ± 4.3
No. of specimens	10	10	10	10	10
1 to 6 weeks
Cell density (no./mm²)	614 ± 249	476 ± 267	420 ± 210	254 ± 48	231 ± 30
Nuclear aspect ratio	4.5 ± 1.0	3.9 ± 0.8	3.7 ± 0.9	4.2 ± 0.7	4.3 ± 1.2
Blood vessel density (no./mm)	4 ± 3.3	2.9 ± 2.6	5.0 ± 2.9	2.0 ± 1.2	0.8 ± 0.2
Cells positive for a-SMA (percent)	2.3 ± 1.4	1.9 ± 1.1	1.0 ± 0.3	0.8 ± 0.31	0.4 ± 0.12
No. of specimens	6	6	6	6	6
8 to 12 weeks
Cell density (no./mm²)	1541 ± 451	1272 ± 363	965 ± 249	701 ± 162	497 ± 151
Nuclear aspect ratio	6.2 ± 1.0	4.3 ± 1.0	3.8 ± 1.0	2.9 ± 1.0	4.1 ± 1.3
Blood vessel density (no./mm)	5.1 ± 3.1	4.0 ± 2.6	3.0 ± 2.1	2.2 ± 1.0	2.1 ± 1.0
Cells positive for a-SMA (percent)	1.3 ± 0.8	1.3 ± 0.3	1.1 ± 0.3	0.5 ± 0.3	0.3 ± 0.2
No. of specimens	5	5	5	5	5
16 to 20 weeks
Cell density (no./mm²)	2244 ± 526	1522 ± 285	1037 ± 280	833 ± 312	1009 ± 437
Nuclear aspect ratio	5.4 ± 1.0	4.8 ± 0.2	4.6 ± 0.5	5.3 ± 1.2	3.8 ± 1.3
Blood vessel density (no./mm)	13.3 ± 4.9	4.0 ± 1.3	5.2 ± 2.0	2.9 ± 1.6	3.3 ± 2.0
Cells positive for a-SMA (percent)	0.6 ± 0.3	0.4 ± 0.2	0.3 ± 0.2	0.3 ± 0.3	1.2 ± 0.7
No. of specimens	6	6	6	6	6
52 to 104 weeks
Cell density (no./mm²)	559 ± 115	601 ± 204	718 ± 241	590 ± 46	546 ± 45
Nuclear aspect ratio	3.7 ± 0.6	4.0 ± 0.9	4.2 ± 0.5	3.3 ± 1.1	3.7 ± 0.5
Blood vessel density (no./mm)	2.1 ± 2.0	1.5 ± 1.3	1.2 ± 0.7	1.6 ± 0.8	1.3 ± 0.6
Cells positive for a-SMA (percent)	0.5 ± 0.3	0.2 ± 0.2	0.2 ± 0.1	0.5 ± 0.3	1.1 ± 0.9
No. of specimens	6	6	6	6	6

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Table II: Histomorphometric Measurements of the Intact and Ruptured Human Anterior Cruciate Ligaments*

*The values are given as the mean and the standard error of the mean. a-SMA = a-smooth muscle actin.

Intact, or No. of Weeks After Rupture	Proximal Edge	1 mm from Edge	2 mm from Edge	4 mm from Edge	6 mm from Edge
Intact ligaments
Cell density (no./mm²)	701 ± 120	525 ± 108	539 ± 91	294 ± 39	265 ± 37
Nuclear aspect ratio	6.1 ± 0.9	4.5 ± 0.8	4.3 ± 0.6	3.6 ± 0.6	2.4 ± 0.5
Blood vessel density (no./mm)	1.5 ± 0.16	1.2 ± 0.2	1.0 ± 0.2	0.60 ± 0.12	0.24 ± 0.03
Cells positive for a-SMA (percent)	4.7 ± 1.0	7.3 ± 1.7	10.7 ± 3.0	15 ± 3.9	17 ± 4.3
No. of specimens	10	10	10	10	10
1 to 6 weeks
Cell density (no./mm²)	614 ± 249	476 ± 267	420 ± 210	254 ± 48	231 ± 30
Nuclear aspect ratio	4.5 ± 1.0	3.9 ± 0.8	3.7 ± 0.9	4.2 ± 0.7	4.3 ± 1.2
Blood vessel density (no./mm)	4 ± 3.3	2.9 ± 2.6	5.0 ± 2.9	2.0 ± 1.2	0.8 ± 0.2
Cells positive for a-SMA (percent)	2.3 ± 1.4	1.9 ± 1.1	1.0 ± 0.3	0.8 ± 0.31	0.4 ± 0.12
No. of specimens	6	6	6	6	6
8 to 12 weeks
Cell density (no./mm²)	1541 ± 451	1272 ± 363	965 ± 249	701 ± 162	497 ± 151
Nuclear aspect ratio	6.2 ± 1.0	4.3 ± 1.0	3.8 ± 1.0	2.9 ± 1.0	4.1 ± 1.3
Blood vessel density (no./mm)	5.1 ± 3.1	4.0 ± 2.6	3.0 ± 2.1	2.2 ± 1.0	2.1 ± 1.0
Cells positive for a-SMA (percent)	1.3 ± 0.8	1.3 ± 0.3	1.1 ± 0.3	0.5 ± 0.3	0.3 ± 0.2
No. of specimens	5	5	5	5	5
16 to 20 weeks
Cell density (no./mm²)	2244 ± 526	1522 ± 285	1037 ± 280	833 ± 312	1009 ± 437
Nuclear aspect ratio	5.4 ± 1.0	4.8 ± 0.2	4.6 ± 0.5	5.3 ± 1.2	3.8 ± 1.3
Blood vessel density (no./mm)	13.3 ± 4.9	4.0 ± 1.3	5.2 ± 2.0	2.9 ± 1.6	3.3 ± 2.0
Cells positive for a-SMA (percent)	0.6 ± 0.3	0.4 ± 0.2	0.3 ± 0.2	0.3 ± 0.3	1.2 ± 0.7
No. of specimens	6	6	6	6	6
52 to 104 weeks
Cell density (no./mm²)	559 ± 115	601 ± 204	718 ± 241	590 ± 46	546 ± 45
Nuclear aspect ratio	3.7 ± 0.6	4.0 ± 0.9	4.2 ± 0.5	3.3 ± 1.1	3.7 ± 0.5
Blood vessel density (no./mm)	2.1 ± 2.0	1.5 ± 1.3	1.2 ± 0.7	1.6 ± 0.8	1.3 ± 0.6
Cells positive for a-SMA (percent)	0.5 ± 0.3	0.2 ± 0.2	0.2 ± 0.1	0.5 ± 0.3	1.1 ± 0.9
No. of specimens	6	6	6	6	6