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(Investigative Ophthalmology and Visual Science. 2003;44:4796-4804.)
© 2003 by The Association for Research in Vision and Ophthalmology, Inc.
DOI:  10.1167/iovs.02-1083
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Effect of Heparin II Domain of Fibronectin on Aqueous Outflow in Cultured Anterior Segments of Human Eyes

Amy J. Santas,1,2 Cindy Bahler,2,3 Jennifer A. Peterson,1 Mark S. Filla,1,4 Paul L. Kaufman,4 Ernst R. Tamm,5 Douglas H. Johnson,3 and Donna M. Pesciotta Peters1,4

1From the Departments of Pathology and Laboratory Medicine and 4Ophthalmology and Visual Sciences, University of Wisconsin, Madison, Wisconsin; the 3Department of Ophthalmology, Mayo Clinic, Rochester, Minnesota; and the 5Department of Anatomy, University Erlangen-Nürnberg, Germany.


   Abstract
Top
 Abstract
Materials and Methods
 Results
 Discussion
 References
 
PURPOSE. To determine whether an integrin/syndecan-binding domain of fibronectin, called the heparin II (Hep II) domain, affects outflow facility in the human eye.

METHODS. Anterior segments of human eyes were placed in perfusion organ culture. One eye of each pair received the Hep II domain, and the fellow eye received DMEM or a heat-denatured Hep II domain. The Hep II domain was produced as a recombinant glutathione S-transferase (GST)-fusion protein. Microscopic changes were assessed.

RESULTS. Outflow facility in anterior segments treated with Hep II domain increased by 93% compared with that in anterior segments treated with DMEM. In contrast, facility in anterior segments treated with the heat-denatured Hep II domain showed very little change. Outflow facility remained high during Hep II domain perfusion and returned to baseline after removal of the protein. Electron microscopy revealed disruptions in the endothelial lining of Schlemm’s canal in anterior segments fixed during maximum effect and in anterior segments after facility had returned to baseline. Scattered disruptions of canal cells were noted in control anterior segments. Trabecular cells in other regions looked normal. Major changes in the extracellular matrix of the juxtacanalicular tissue were not observed. Repeated doses of the Hep II domain administered after facility returned to baseline increased facility in two of three anterior segments.

CONCLUSIONS. The Hep II domain of fibronectin increases outflow facility in the human anterior segment. This suggests that fibronectin-mediated interactions may have a role in modulating aqueous hydrodynamics. Such interactions may represent avenues of novel therapeutic interventions for glaucoma.

Recent studies indicate that the actomyosin cytoskeleton in the trabecular meshwork (TM) may be involved in regulating aqueous humor hydrodynamics.1 2 3 In enucleated calf and human eyes and in living eyes of monkeys and humans, chemical agents that disrupt the cytoskeleton or the signaling pathways that maintain the network cause an increase in outflow facility. Among the agents used to disrupt the actomyosin cytoskeleton in the TM and alter outflow facility are cytochalasins B and D,4 5 H-7,6 7 Y-27632,3 latrunculin-A8 (Peterson JA, et al. IOVS 1996;37:ARVO Abstract S825), and latrunculin-B.9 10 These chemicals are believed to exert their effect on outflow by disrupting the actomyosin network and hence weakening cell-cell and cell-extracellular matrix adhesions used to maintain tissue architecture.

Several signaling pathways control the assembly and contractility of the actomyosin network. Among the pathways that regulate the actomyosin cytoskeleton and help maintain cell contacts are signaling events regulated by various members of the integrin and/or syndecan receptor families.11 12 13 14 Integrins are transmembrane receptors composed of an {alpha}- and ß-subunit covalently associated into a heterodimer.15 There are more than 20 different integrin receptors, each having their own ligand specificity. In general, however, they bind various extracellular matrix proteins, usually through a specific Arg-Gly-Asp (RGD) sequence. Human TM tissues express a spectrum of integrin subunits including {alpha}1, {alpha}3, {alpha}4, {alpha}5, {alpha}6, {alpha}v, ß1, ß3, ß4, and ß5.16 Syndecans are transmembrane heparan sulfate proteoglycans that bind several extracellular matrix proteins, cytokines, and growth factors, usually through interactions with their heparan sulfate side chains.14 17 The anterior segment of the human eye has been found to contain high levels of syndecan-3 and -4 as well as low levels of syndecan-1 and -2 (Filla MS, et al. IOVS 2002;43:ARVO E-Abstract 1025). Thus, cellular interactions between integrins and syndecans could play a regulatory role in modulating outflow facility.

One of the major extracellular matrix components closely associated with trabecular cells and capable of interacting with integrins and syndecans is fibronectin. Fibronectin has been found in the juxtacanalicular region and along the inner wall of Schlemm’s canal.18 19 Fibronectin is also present as a soluble protein in aqueous humor.20 21 The expression of fibronectin in the TM is regulated by factors believed to play a role in regulating outflow facility, including glucose,22 glucocorticoids,23 24 25 transforming growth factor-ß,26 and ascorbic acid.25 27 It is unclear, however, whether changes in the level of expression of fibronectin occur during aging and glaucoma.18 19 28

Fibronectin is a matrix glycoprotein that consists of two disulfide-bonded polypeptide chains.29 Each chain contains a series of homologous repeating units arranged into distinct biological domains. Cell adhesion to fibronectin and the subsequent assembly and contractility of the actin cytoskeleton are mediated by two specific domains in fibronectin: the central cell binding domain and a heparin-binding domain, Hep II.30 31 Both of these domains contain a binding site for members of the integrin family. The central cell-binding domain contains an RGDS sequence,32 33 and the Hep II domain contains a homologous sequence, IDAPS.34 In addition, the Hep II domain contains binding sites for various members of the syndecan family, including syndecan-1, -2, and -4.35 36 37

In this study, we examined whether integrin/syndecan-binding domains of fibronectin could have a role in regulating outflow facility in the human eye. To determine this, anterior segments of human eyes in culture were perfused with a recombinant Hep II domain of fibronectin. In some experiments, a heat-denatured Hep II domain was used. Our results show that the nondenatured Hep II domain significantly increased outflow facility.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
Preparation of Recombinant Proteins
An expression construct containing the Hep II domain of fibronectin was made by cloning the cDNA for the III12-14 repeats of fibronectin (Fig. 1) into a glutathione S-transferase (GST) expression system, as previously described.38 The recombinant GST-Hep II fusion protein was purified by using glutathione Sepharose 4B beads, according to the manufacturer’s instructions (Amersham Biosciences Corp., Piscataway, NJ). The Hep II domain was separated from the GST tag with a thrombin (1 NIH U/50 mL of original culture; Hematologic Technologies, Inc., Essex, VT) digest at 4°C, while the GST fusion protein was still on the column. Thrombin was removed from the supernatant containing the Hep II domain by using p-aminobenzamide Sepharose 6B beads as instructed by the manufacturer (Sigma-Aldrich, St. Louis, MO). Heat-denatured Hep II domains were generated by heating the protein to 80°C for 5 minutes. An immunofluorescence matrix assembly microscopy assay was used to verify the inactivation of the Hep II domain.38 39 On a 10% SDS-PAGE gel stained with Coomassie blue R,40 both the nondenatured and denatured Hep II domains migrated as a single but diffuse band of ~30 kDa (data not shown). The Hep II domain was dialyzed against 0.2 M ammonium bicarbonate and then lyophilized. For all the perfusion studies, the Hep II domain was reconstituted in DMEM. For the remainder of this article the recombinant Hep II domain will be referred to as rHep II.



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  		FIGURE 1. Model of fibronectin. Diagram shows the location of the rHep II domain (shaded ovals) in a fibronectin monomer relative to the arginine-glycine-aspartic acid (RGD) integrin-binding sequence in the 10th type III repeat. The type III repeats are numbered. The IIICS domain is the alternatively spliced variant region of fibronectin. The amino (N) and carboxyl (C) termini of fibronectin are noted.

 
Culture Technique
Twelve pairs of fresh normal human eye bank eyes were studied (obtained within 17 hours after death). The average age of the donor eyes was 72 ± 16 years (range, 51–90 years). No eyes had glaucoma or uveitis, nor were the donors using topical medications. The culture technique was similar to that described previously.41 42 Eyes were bisected at the equator, and the iris, lens, and vitreous were removed. The anterior segment was clamped in a modified Petri dish and the eye perfused with Dulbecco’s modified Eagle’s medium with added antibiotics (penicillin, 10,000 U; streptomycin, 10 mg; amphotericin B, 25 mg; and gentamicin, 17 mg in 100 mL medium; Sigma-Aldrich) at the normal human flow rate (2.5 µL/min). The anterior segments were cultured at 37°C in a 5% CO2 atmosphere. Intraocular pressures were continuously monitored with a pressure transducer connected to the second access cannula built into the dish and recorded with an automated computerized system.

Protein Infusion
After an initial adaptation period in culture usually lasting 2 to 4 days, one anterior segment from a pair of human eyes was given rHep II through an anterior chamber (AC) exchange, whereas that from the fellow control eye underwent an AC exchange with either DMEM or heat-denatured rHep II. The osmolarity of the rHep II solution was 349 mmol/kg H2O at pH 7.8. The AC exchanges were performed with a gravity-driven, constant-pressure method over a 10-minute period. The infusion pumps were turned off for 3 to 4 hours to allow time for the protein to act competitively at its sites without being washed away by the continual flow of culture medium. The concentration of the rHep II used for the AC exchange was 1.25 mg/mL (46 µM).

The experimental anterior segments were perfused continuously with 30 µM rHep II (0.833 mg/mL) dissolved in DMEM. Control anterior segments were perfused with DMEM or 30 µM heat-denatured rHep II (0.833 mg/mL) dissolved in DMEM. In two anterior segments, the rHep II remained in the medium until the eyes were fixed, approximately 24 hours after the initial infusion of the protein. Intraocular pressure remained low during this time, which allowed histologic observation of eyes while they were still affected by the protein. In other experiments, the rHep II was removed approximately 18 hours after its initial infusion, by changing the culture medium to standard medium. This allowed a gradual washing out of the protein from the AC (estimated t1/2 = 5 hours). Pressures were monitored for another 24 hours to see whether the effect of the rHep II would wear off before fixation.

Repeated-dose studies were performed on three anterior segments after the rHep II had been washed out of the initial infusion. After the removal of the rHep II, a baseline was established, and an AC exchange with a fresh solution of rHep II followed by a perfusion with rHep II was repeated. In all experiments, outflow facility (C = F/P) was calculated every hour, beginning 3 hours before drug infusion and continuing for the duration of the culture. Results from each pair were combined into a group mean.

A double-switch experiment with nondenatured and heat-denatured rHep II was performed on two anterior segments. In these experiments, one anterior segment was perfused with the rHep II domain and the other with the heat-denatured rHep II, as just explained. The domains were washed out, and then a second dose of nondenatured and heat-denatured rHep II was perfused into the opposite anterior segment previously used.

All anterior segments were fixed with 4% paraformaldehyde in 0.1 M phosphate buffer through either AC perfusion or immersion into fixative. Perfusion fixation was performed at the pressure level equal to that within the eye when the experiment was completed. Immersion fixation was used in some eyes to prevent artifactual loss of cells potentially weakened in their attachment to the underlying extracellular matrix by the rHep II.

Histologic Examination
For the light and transmission electron microscopy studies, at least four wedges of tissue were dissected from each quadrant. The wedges were of 1 to 1.5 mm circumferential width containing anterior ciliary muscle, scleral spur, and TM. Tissue was dehydrated in ascending concentrations of alcohol and embedded in epoxy resin according to standard protocols. Semithin sections were stained with toluidine blue. Thin sections were stained with lead citrate and uranyl acetate.

The cultures were evaluated by light microscopy of 1-µm semithin sections and by transmission electron microscopy to assess the appearance of the trabecular cells, evidence of toxicity, and changes in the endothelial cells of the canal. Examination was performed on at least two sections from each quadrant in a masked fashion, by using predetermined criteria including cell shape, nuclear shape, preservation of cellular and nuclear membranes, cell-cell attachments, cytoplasmic covering of trabecular lamellae, preservation of endothelial lining of Schlemm’s canal, and integrity of trabecular lamellae.42 43 Meshworks were considered normal in appearance if trabecular cells remained in their usual position on the lamellae, a normal number of cells was present (subjective assessment), and little disruption of the juxtacanalicular tissue and trabecular lamellae was seen.

Immunoperoxidase Staining of Cultured Eyes
Paraffin-embedded sections from four pairs of anterior segments were stained for rHep II with the mAb IST-244 45 at 8 µg/mL. As a negative control, sections were labeled with the mAb GAL-13 against ß-galactosidase (Sigma-Aldrich). Antibodies were localized with an immunoperoxidase protein assay kit (ABC Elite; Vector Laboratories, Burlingame, CA), using a red substrate (NovaRed; Vector Laboratories).

Statistical Analysis
Drug effects were expressed as the outflow facility after drug infusion (Cd) divided by the baseline facility (Co) for each anterior segment.46 The percentage of change in outflow facility was expressed as the ratio of the experimental to the control anterior segment of each pair. A percent increase or decrease in facility compared to baseline was calculated as (Cd/Co - 1) x 100. The no-effect value is assumed to be 1. Results from each pair were combined into a group mean. Values are expressed as the mean ± SEM. Statistical analyses were performed with a Wilcoxon-signed rank test, which is the nonparametric analogue to the paired sample t-test. Confidence limits were determined based on the differences in the paired population. Outliers were not discarded.


   Results
Top
 Abstract
 Materials and Methods
 Results
Discussion
 References
 
Facility of Outflow
Perfusion of anterior segments with rHep II (Fig. 1) resulted in a decrease in intraocular pressure (IOP; Fig. 2 ). The onset of this effect appeared immediately after the infusion pump was restarted (4 hours after it had been turned off). The decrease in pressure was at its peak at this time point and lasted as long as the rHep II was in the perfused culture medium. After removal of the recombinant domain from the culture medium, pressure returned to near baseline values in seven of the eight eyes allowed to return to baseline. The time required to return to baseline ranged from 2 to 12 hours. Within 9 hours of perfusion of the rHep II through 10 pairs of anterior segments, outflow facility in the rHep II-treated anterior segment was found to be statistically greater (P < 0.01) than in control anterior segments. In anterior segments treated with rHep II, facility increased by 120% (Cd/Co = 2.20 ± 0.43; Table 1 ) compared with baseline. In contrast, fellow control anterior segments increased by only 14% (Cvehicle/Co = 1.14 ± 0.07; Table 1 ) compared with baseline. The overall change in facility when compared with control anterior segments was 93%.



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  		FIGURE 2. Effect of rHep II and DMEM on outflow facility. An AC was exchanged with 1.25 mg/mL rHep II (•) or DMEM ({circ}). After the exchange, the pumps were turned off for 4 hours to allow the domains time to interact with possible receptors in the TM. The pumps were then turned on, and the ACs were perfused with 30 µM rHep II or buffer. Arrows: times when the pumps were turned off and then on again. After 27 hours (arrowhead), the rHep II was washed out and pressure returned to baseline.

 

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  		TABLE 1. Effect of rHep II on Outflow Facility in Cultured Anterior Segments

 
The magnitude of the facility increase caused by rHep II varied, ranging from 20% to 400% in 9 of the 10 anterior segments tested. Five anterior segments had an increase ranging from 20% to 55%, whereas four anterior segments ranged from 101% to 400% (Table 1) . One anterior segment had a decrease in facility after treatment with the rHep II. These differences did not correlate with gender, time after death, or time in culture.

To demonstrate that the biological activity of the rHep II was responsible for the increase in outflow, fellow anterior segments were perfused with heat-denatured rHep II. By SDS-PAGE analysis, the heat-denatured domain remained intact and was not degraded by the treatment (data not shown). Heat denaturation, however, affected the biological activity38 39 of the rHep II, because the domain no longer inhibited fibronectin fibrillogenesis (data not shown). As shown in Table 2 (donors 11 and 12), the heat-denatured rHep II had little effect on outflow facility compared with fellow anterior segments perfused with nondenatured rHep II. To demonstrate that the effect of the heat denatured rHep II on outflow facility was not due to an unresponsiveness of those anterior segments, double-switch experiments were performed in which the anterior segment previously perfused with heat-denatured rHep II was now perfused with the nondenatured rHep II and vice versa. In one experiment (donor 10), the control anterior segment was perfused with DMEM and then with the heat-denatured rHep II. As expected, anterior segments that were previously unresponsive to heat-denatured rHep II were now responsive to the nondenatured rHep II. In contrast, the anterior segment that was previously responsive to rHep II showed very little response to heat-denatured rHep II (Table 2) . The anterior segment first treated with DMEM (donor 10) was also unresponsive to heat-denatured rHep II. The facility for anterior segments perfused with the heat-denatured rHep II (n = 4; donors 11 and 12) increased by 5% (CdrHep/Co = 1.05 ± 0.06) compared to baseline. In contrast, facility increased by 162% (CnrHep II/Co = 2.62 ± 0.36) compared with baseline in fellow anterior segments treated with the nondenatured rHep II. This suggests that the decrease in pressure observed with the rHep II was due to its specific biological activity.


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  		TABLE 2. Effect of Heat Denaturation of rHep II Reduces Effect on Outflow Facility

 
Table 3 shows that two of three anterior segments remained sensitive to repeated treatments with the rHep II, even after the initial treatment was washed out. After the first treatment, an increase of 52% in facility was observed in anterior segments treated with the rHep II compared with control anterior segments. During the second treatment, two of the three anterior segments receiving repeated doses of rHep II showed an increase in facility, whereas all the fellow anterior segments treated with repeated doses of vehicle showed minimal change or even a small decrease in facility. The overall increase in facility for all three anterior segments receiving the second treatment of rHep II was 90% compared with the control anterior segments.


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  		TABLE 3. Effect of Repeated Doses of rHep II on Outflow Facility

 
Histologic Examination
Immunoperoxidase labeling studies were performed with a monoclonal antibody to rHep II to localize the site where the domain was binding. These anterior segments were immersion-fixed before washout of the rHep II. In the control anterior segment, endogenous fibronectin (containing the Hep II domain) was observed throughout much of the anterior segment (Figs. 3A 3D) . Staining was most prominent around and within the trabecular beams. In anterior segments perfused with rHep II, the level of rHep II increased above the level of endogenous fibronectin and was observed uniformly distributed throughout the TM and adjacent tissues (Figs. 3B 3E) . Large precipitates of the protein were not observed anywhere within the anterior segments. Serial sections labeled with a negative control antibody against ß-galactosidase demonstrated little or no significant staining in either the control anterior segment (not shown) or the anterior segment perfused with rHep II (Figs. 3C 3F) .



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  		FIGURE 3. Immunohistochemistry of the anterior segment with IST-2. Anterior segment perfused with buffer (A, D) or rHep II (B, C, E, F). Sections labeled with the mAb IST-2 (A, B, D, E) or with the negative control mAb GAL-13 (C, F). The TM and Schlemm’s canal (SC) are indicated. White arrows: collector channels. Black arrows: location of SC. The spaces present within the corneoscleral junction are artifacts of the sectioning. Regions in (A), (B), and (C) marked by asterisks are shown at higher magnification in (B), (E), and (F), respectively. The compression in the TM in (B) and (C) was not seen in other anterior segments treated with the Hep II and thus is not the result of the treatment with the Hep II. Scale bars: (A) 100 µm; (D) 50 µm.

 
Light microscopy studies indicated that meshworks from eyes treated with the rHep II and fixed at maximal pressure showed patchy areas of breaks or disruptions in the endothelial cells lining the inner wall of Schlemm’s canal (Fig. 4B) . These changes were clearly seen in all eight sections, from the circumference of each specimen that was analyzed. In these areas, endothelial cells were frequently missing over larger distances, and the extracellular matrix of the juxtacanalicular tissue (JCT) was in direct contact with the lumen of Schlemm’s canal (Fig. 4B) . Such areas were not seen in control anterior segments (Fig. 4A) , in which the endothelial lining of Schlemm’s canal was usually complete. In some control anterior segments, however, some scattered, rare breaks in the lining of the canal were observed (not shown). If breaks were present in control eyes, they were only in one or two sections of the circumference. In addition, they were much smaller than those observed in experimental eyes and corresponded to the length of one or two endothelial cells. Cells in the juxtacanalicular region remained intact without any ultrastructural evidence of toxicity. Trabecular cells on the lamellae in the corneoscleral and uveal regions appeared normal, and remained in place with intact cytoplasm and cytoplasmic membranes (Fig. 4C) . In these regions some scattered cell loss was apparent in both anterior segments treated with rHep II and control anterior segments, consistent with previous findings in organ cultured anterior segments.46



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  		FIGURE 4. Light microscopy of Schlemm’s canal after perfusion. (A) Anterior segment perfused with buffer. Schlemm’s canal (SC) endothelium (arrows) and juxtacanalicular meshwork appeared to be intact. (B) Eye perfused with the rHep II. Schlemm’s canal endothelial cells were missing in some areas along the inner and outer wall of the canal (arrows). Semithin sections were stained with toluidine blue. (C) Anterior segment perfused with the rHep II (same anterior segment as B, but different section). By electron microscopy, TM cells in the corneoscleral region appeared to be structurally intact. Anterior segments were fixed and prepared for microscopy after the rHep II was washed out and baseline facility returned to normal. Scale bar: (A, B) 10 µm; (C) 1.1 µm.

 
Meshworks from anterior segments fixed after the rHep II had been removed and facility returned to baseline revealed similar disruptions and breaks in the lining of Schlemm’s canal as in eyes fixed at the maximum change in pressure. In some regions, larger sheaths of inner wall endothelial cells were separated from the underlying extracellular matrix, whereas the cells and their cell junctions were intact (Fig. 5B) . In other areas, disconnected endothelial cells were ruptured (Fig. 5D) . Such changes were not observed in control eyes (Figs. 5A 5C) . In experimental eyes, detached endothelial cells were seen in the lumen of Schlemm’s canal. The nuclei of these cells often showed condensed chromatin, indicating apoptotic changes (Fig. 5F) . Overall, it appeared that the canal cells detached from their underlying extracellular matrix and lifted off, presumably because of the continual perfusion of fluid into the canal. The cells then ruptured and were lost. In addition, there were focal areas in which the endothelial lining was incomplete, but vesicle-like cellular structures were still seen between the lumen of Schlemm’s canal and the extracellular matrix of the JCT (Fig. 6B) . The vesicle-like structures were 80 to 100 nm in diameter and were surrounded by a cell membrane, indicating a cellular origin (Fig. 7A) . They lined the extracellular matrix of the JCT, but were also present in the lumen of Schlemm’s canal. Adjacent sections often showed partially detached inner wall cells that formed a cytoplasmic protrusion similar in diameter to the vesicle-like structures (Fig. 7B) , indicating that at least some of the vesicle-like structures might be part of such protrusions.



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  		FIGURE 5. Electron microscopy of Schlemm’s canal (SC) endothelium and juxtacanalicular meshwork (JCT). Anterior segment was perfused with rHep II (B, D–F), whereas the contralateral eye was perfused with DMEM (A, C). The anterior segment was fixed and prepared for electron microscopy after the rHep II domain was washed out and baseline facility returned to normal. (A) In the control anterior segment, the inner wall of Schlemm’s canal was regularly in close contact with the extracellular matrix of the JCT (arrows). (B) In anterior segments treated with rHep II, Schlemm’s canal endothelium was separated from the extracellular matrix of the JCT in parts of the circumference (arrows). (C) Higher magnification of the association between extracellular matrix and Schlemm’s canal endothelial cells in the control eye. Fine fibrillar material was in close contact with the cell membrane of the endothelial cells. In areas of contact, the cell membrane was more electron dense than on the luminal side (solid arrows). Open arrow: cell junction between two adjacent endothelial cells. (D) In treated eyes, Schlemm’s canal endothelial cells were detached from the extracellular matrix of the JCT (solid arrows). Open star: fragments of a detached cell in the lumen of Schlemm’s canal; open arrow: junction between adjacent endothelial cells. (E, F) Areas of detached Schlemm’s canal endothelium along inner (E) and outer (F) wall of Schlemm’s canal in the left eye. The so-called connecting fibrils of the cribriform network were disconnected from Schlemm’s canal endothelium (solid arrows). Open arrows: endothelial cells that are still covering the lumen of Schlemm’s canal; open star: nucleus of a detached endothelial cell. The nucleus shows heavily condensed chromatin, indicating apoptosis. Scale bar: (A, B) 1.83 µm; (C) 0.6 µm; (D) 1 µm; (E, F) 2.8 µm.

 


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  		FIGURE 6. Electron microscopy of Schlemm’s canal (SC) endothelium and juxtacanalicular meshwork (JCT). The anterior segment was perfused with rHep II (B), whereas the contralateral eye was perfused with DMEM (A). Eyes were fixed and prepared for electron microscopy after the rHep II domain was washed out and baseline facility returned to normal. (A) In the control anterior segment, the endothelial lining of the inner wall of Schlemm’s canal (SC) was complete and in close contact with the extracellular matrix of the JCT (arrows). (B) In the anterior segment treated with rHep II, an accumulation of electron-dense, fine granular material was seen between Schlemm’s canal endothelium and cells of the JCT (solid star). In some areas, openings in Schlemm’s canal endothelium were present and endothelial cells were detached from the extracellular matrix of the JCT (solid arrows). In addition, there were focal areas in which the endothelial lining was incomplete, and vesicle-like cellular structures were seen between the lumen of Schlemm’s canal and the extracellular matrix of the JCT (open arrows). Cell junctions are within the circles. Scale bars, 1.1 µm.

 


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  		FIGURE 7. Electron microscopy of Schlemm’s canal (SC) endothelium and juxtacanalicular meshwork (JCT). (A) Higher magnification of the area in Figure 6B showing the vesicle-like structures (open arrows). These structures were 80 to 100 nm in diameter and were surrounded by a cell membrane. They lined the extracellular matrix of the JCT, but were also in the lumen of Schlemm’s canal. (B) An adjacent section shows a cell of the inner wall of Schlemm’s canal (SC) that was detached from its underlying extracellular matrix (solid arrows) and formed a cytoplasmic protrusion similar in diameter to the vesicle-like structures shown in Figure 8A . Scale bars, 0.4 µm.

 
These findings were present in both immersion-fixed and perfusion-fixed eyes (data not shown). In some areas, the connecting fibrils of the tendon network in the juxtacanalicular region were completely disconnected from the canal endothelial cells (Fig. 5E) . The extracellular matrix in the meshwork generally appeared unaffected by the treatment with rHep II. In areas along the canal wall where endothelial cells were lost, the underlying basement membrane and subendothelial matrix material usually remained intact, maintaining a sharp linear border adjacent to the canal (Fig. 5D) . In some areas, the subendothelial matrix formed a basal lamina-like band of fine filamentous material (Fig. 8A) . In other regions, the subendothelial matrix was composed of more irregularly dispersed fine fibrillar material (Fig. 8B) . Some regions showed some amorphous, fine granular material within the juxtacanalicular tissue that was not observed in control eyes (Figs. 6A 6B) . The nature of this material is unclear, but might reflect changes in the ultrastructure caused by the rHep II.



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  		FIGURE 8. Electron microscopy of Schlemm’s canal (SC) endothelium and juxtacanalicular meshwork (JCT). Anterior segments perfused with rHep II were fixed and prepared for electron microscopy after the rHep II domain was washed out and baseline facility returned to normal. (A) Area in which endothelial cells were lost and vesicle-like structures were seen (solid arrow). The subendothelial matrix formed a basal lamina-like band of fine filamentous material (open arrows). (B) In a different region of the same eye, the subendothelial matrix was composed of more irregularly dispersed fine fibrillar material (open arrows). Scale bars, 0.4 µm.

 

   Discussion
Top
 Abstract
 Materials and Methods
 Results
 Discussion
References
 
A recombinant Hep II from fibronectin, containing both an integrin- and syndecan-binding site, caused a 93% increase in outflow facility in human anterior segments. This effect was dependent on the biological activity of the rHep II, because heat-denatured rHep II did not change facility. This suggests that the rHep II interfered with specific interactions between cells and extracellular matrix proteins in the meshwork that regulate outflow facility. Such interactions may be influential in the regulation of aqueous outflow.

The effect of the rHep II on the TM appeared to be similar to that previously reported in organ culture eyes treated with cytochalasin D and H-7.46 47 48 Both treatments resulted in breaks in the inner and outer wall cells of Schlemm’s canal, whereas cells in the uveal and corneoscleral meshwork remained intact. In cultured anterior segments treated with cytochalasin D, the 40% increase in outflow facility was accompanied by a 5% loss of canal cells. Similarly, treatment with H-7, which increased outflow facility by 43%, was also accompanied by a loss of canal cells in cultured anterior segments. In contrast, cytochalasin B and H-7 did not cause a loss of canal cells in monkeys, even though both drugs increased outflow facility in the monkey.4 7 48 Treatment with cytochalasin B, however, affected the TM in monkeys in which separation, degeneration, and disappearance of meshwork cells was observed in the uveal and corneoscleral regions.

These breaks did not appear to be artifacts, because the remnants of the canal wall cells still on the basement membrane had rounded cell boundaries. The cell fragments were encircled by cytoplasmic membrane in the regions of disruptions, suggesting that the cell membranes reconfigured themselves after the initial disruption. In contrast, artifactual breaks would be linear and jagged, with bare cytoplasm in the broken region. More important, the changes were present in the experimental eyes but not in the control eyes. If a systematic processing artifact were present that caused loss of cells, it should affect both eyes. Thus, it appears that the breaks occurred in response to the rHep II.

Giant vacuoles were not routinely seen in either the control or rHep II-treated anterior segments. This was probably because the formation of vacuoles is pressure dependent and is most commonly seen in perfusion fixed tissue. Thus, many of our fixed anterior segments that were fixed by immersion would not be expected to have giant vacuoles. In addition, treatment with the rHep II caused a loss of Schlemm’s canal cells, which most likely eliminated the resistance of this layer to aqueous outflow, and so the media could simply bypass the intact cells through the holes created by the rHep II.

The increase in facility caused by the rHep II could not have been due to the disruption of canal cells along the inner and outer wall of Schlemm’s canal. When Bill and Svedbergh49 analyzed the contribution of the pores in the canal cells on outflow facility, they calculated that the canal cells alone account for less than 10% of outflow resistance. This suggests that the removal of Schlemm’s canal cells was not the sole cause of the increase in outflow facility by the rHep II. In addition, outflow facility returned to baseline, even though breaks in the cellular lining of the canal persisted and repeated doses of the rHep II were effective in increasing outflow facility. Thus, a part of the regulatory mechanism that controls outflow facility was still intact and responsive to the rHep II, despite the loss of Schlemm’s canal cells. Although, the effect of repeated doses of rHep II could be from the loss of additional canal wall cells, as not all cells may have been lost after a single dose, this would not explain why facility returns to baseline in the presence of the breaks along Schlemm’s canal.

This is not to say, however, that canal cells do not have a role in modulating outflow facility. The canal cells could modify the resistance of the underlying extracellular matrix and have a greater influence on facility. In this scenario, the canal cells and underlying basement membrane function as a unit, with the giant vacuoles and pores of the cells causing aqueous humor to "funnel" or be concentrated in regions of the juxtacanalicular tissue.50 This effectively increases the resistance of the adjacent extracellular matrix. Loss of canal cells by the rHep II could therefore contribute to the increase in outflow facility by destroying the funneling effect.

Alternatively, the effect of the rHep II on outflow facility could be through effects on cellular contractility of juxtacanalicular cells. By binding to cellular receptors, the rHep II could cause changes in signaling pathways within cells and perhaps decrease the contractility of the juxtacanalicular cells, allowing this region to expand as may occur in monkeys after treatment with H-7.7 Although no obvious changes in the configuration of the juxtacanalicular region were evident on microscopic examination, subtle changes could have occurred after treatment with the rHep II.

Cellular contractility of meshwork cells has been demonstrated in vitro and involves modulation of Rho GTPase and a PKC-dependent signaling pathway.2 Contractility of the meshwork could be a target of the rHep II, consistent with its signaling role in other cell types. In fibroblasts, rHep II is responsible for the organization of the actin cytoskeleton.30 31 Several agents, including the serine-threonine kinase inhibitor H-77 51 and the Rho-kinase inhibitor Y27632,3 affect the same pathways that are used by rHep II to modulate formation of stress fibers and focal adhesions in fibroblasts.31 52 If rHep II was modulating the contractility of the meshwork, facility could return to baseline when the drug perfusion stopped, and repeated doses of rHep II could further increase facility in the presence of breaks in the canal cell lining.

The biological activity of rHep II responsible for the enhanced outflow is unknown. rHep II can interact with several transmembrane receptors, including integrins ({alpha}4ß1/ß7) and syndecan-1, -2, and -4, and both {alpha}4ß1 integrins and syndecan-4 are in the TM of the human eye (Zhou L, et al. IOVS 1999;40;ARVO Abstract 1279; Filla MS, et al. IOVS 2002;43 ARVO E-Abstract 1025). Interactions with syndecan-4 and {alpha}4ß1 integrins have been found to modulate a number of actomyosin-based processes.31 52 53 54 In some instances control of these actomyosin-based processes involves cooperative interactions between cell surface proteoglycans and integrins.55 Thus, the biological activity of the rHep II could involve either integrin-mediated signaling events or cooperative signaling events between integrins and heparan sulfate proteoglycans.


   Footnotes
 
2 Contributed equally to the work and therefore should be considered equivalent senior authors. Back

Supported in part by National Eye Institute Grants EY12515 (DPP), EY02698 (PLK), and EY07065 (DJ), and by a Research to Prevent Blindness Physician Scientist Award (PLK), SFB 539 (ERT), and AHAF (DPP) grants.

Submitted for publication October 23, 2002; revised April 23, 2003; accepted June 23, 2003.

Disclosure: A.J. Santas, None; C. Bahler, None; J.A. Peterson, None; M.S. Filla, None; P.L. Kaufman (P); E.R. Tamm, None; D.H. Johnson, None; D.M.P. Peters (P)

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Corresponding author: Donna M. Peters, Department of Pathology and Laboratory Medicine, University of Wisconsin-Medical School, 1300 University Ave., Madison, WI 53706; dmpeter2{at}facstaff.wisc.edu.


   References
Top
 Abstract
 Materials and Methods
 Results
 Discussion
 References
 

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