Abstract
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-A
8 (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 α- and β-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 α1, α3, α4, α5, α6, αv, β1, β3, β4, and β5.
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-β,
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.
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.
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 Svedbergh
49 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-7
7 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 (α4β1/β7) and syndecan-1, -2, and -4, and both α4β1 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 α4β1 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.
Contributed equally to the work and therefore should be considered equivalent senior authors.
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;
[email protected].
Table 1. Effect of rHep II on Outflow Facility in Cultured Anterior Segments
Table 1. Effect of rHep II on Outflow Facility in Cultured Anterior Segments
Donor | Treatment | Anterior Segment | IOP Pre-rHep II | IOP 9 h Post rHep II | C o | C d9 | C d/C o | % Change | Time of Fixation (h) | Method of Fixation |
1 | rHep II | Exp. | 22 | 15 | 0.11 | 0.17 | 1.47 | 55 | 45 | P |
| DMEM | Control | 21 | 22 | 0.12 | 0.11 | 0.95 | — | | |
2 | rHep II | Exp. | 21 | 5 | 0.12 | 0.50 | 4.20 | 320 | 20 | P |
| DMEM | Control | 15 | 15 | 0.17 | 0.17 | 1.00 | — | | |
3 | rHep II | Exp. | 24 | 45 | 0.10 | 0.06 | 0.53 | −66 | 60 | P |
| DMEM | Control | 19 | 12 | 0.13 | 0.21 | 1.58 | — | | |
4 | rHep II | Exp. | 24 | 17 | 0.10 | 0.15 | 1.41 | 31 | 42 | IM |
| DMEM | Control | 26 | 24 | 0.10 | 0.10 | 1.08 | — | | |
5 | rHep II | Exp. | 21 | 15 | 0.12 | 0.17 | 1.40 | 22 | 24 | IM |
| DMEM | Control | 15 | 13 | 0.17 | 0.19 | 1.15 | — | | |
6 | rHep II | Exp. | 15 | 3 | 0.17 | 0.83 | 5.00 | 400 | 34 | IM |
| DMEM | Control | 16 | 16 | 0.16 | 0.16 | 1.00 | — | | |
7* | rHep II | Exp. | 15 | 7 | 0.17 | 0.36 | 2.14 | 53 | — | NA |
| DMEM | Control | 14 | 10 | 0.18 | 0.25 | 1.40 | — | | |
8* | rHep II | Exp. | 17 | 9 | 0.15 | 0.28 | 1.89 | 101 | — | NA |
| DMEM | Control | 15 | 16 | 0.17 | 0.16 | 0.94 | — | | |
9* | rHep II | Exp. | 28 | 17 | 0.09 | 0.15 | 1.65 | 20 | — | NA |
| DMEM | Control | 11 | 8 | 0.23 | 0.31 | 1.38 | — | | |
10, † | rHep II | Exp. | 14 | 6 | 0.18 | 0.42 | 2.33 | 165 | — | NA |
| DMEM | Control | 15 | 17 | 0.17 | 0.15 | 0.88 | — | | |
Mean ± SEM | | Exp. | | | | | 2.20 ± 0.43 | 93 | — | |
| | Control | | | | | 1.14 ± 0.07 | | | |
Table 2. Effect of Heat Denaturation of rHep II Reduces Effect on Outflow Facility
Table 2. Effect of Heat Denaturation of rHep II Reduces Effect on Outflow Facility
Donor | Anterior Segment | First Treatment | C o | C d9 | C d/C o | % Change | Second Treatment | C o | C d9 | C d/C o | % Change |
11 | Right | dn-rHep II | 0.12 | 0.13 | 1.05 | 158 | rHep II | 0.17 | 0.28 | 1.67 | 67 |
| Left | rHep II | 0.13 | 0.36 | 2.71 | — | dn-rHep II | 0.17 | 0.17 | 1.00 | — |
12 | Right | dn-rHep II | 0.15 | 0.18 | 1.21 | 123 | rHep II | 0.15 | 0.50 | 3.40 | 257 |
| Left | rHep II | 0.09 | 0.25 | 2.70 | — | dn-rHep II | 0.14 | 0.13 | 0.95 | — |
10* | Right | DMEM | 0.17 | 0.15 | 0.88 | 165 | dn-rHepII | 0.17 | 0.18 | 1.06 | 0 |
| Left | rHep II | 0.18 | 0.42 | 2.33 | — | DMEM | 0.17 | 0.18 | 1.06 | — |
Table 3. Effect of Repeated Doses of rHep II on Outflow Facility
Table 3. Effect of Repeated Doses of rHep II on Outflow Facility
Donor | Anterior Segment | C o | C d9 | 1st Dose C d/C o | % Change | C o | C d9 | 2nd Dose C d/C o | % Change |
7* | Experiment | 0.17 | 0.36 | 2.14 | 69 | 0.36 | 0.83 | 2.33 | 191 |
| Control | 0.18 | 0.25 | 1.40 | — | 0.31 | 0.25 | 0.80 | — |
8* | Experiment | 0.15 | 0.28 | 1.89 | 89 | 0.23 | 0.23 | 1.00 | 6 |
| Control | 0.17 | 0.16 | 0.94 | — | 0.17 | 0.16 | 0.94 | — |
9* | Experiment | 0.09 | 0.15 | 1.65 | 59 | 0.15 | 0.23 | 1.55 | 87 |
| Control | 0.23 | 0.31 | 1.38 | — | 0.25 | 0.21 | 0.83 | — |
Mean | Experiment | | | 1.89 ± 0.14 | 52 | | | 1.63 ± 0.39 | 90 |
| Control | | | 1.24 ± 0.15 | — | | | 0.86 ± 0.04 | — |
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