The Aqueous Outflow System as a Mechanical Pump: Evidence from Examination of Tissue and Aqueous Movement in Human and Non-Human Primates : Journal of Glaucoma

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Basic Sciences in Clinical Glaucoma

The Aqueous Outflow System as a Mechanical Pump

Evidence from Examination of Tissue and Aqueous Movement in Human and Non-Human Primates

Johnstone, Murray A MD

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Journal of Glaucoma 13(5):p 421-438, October 2004. | DOI: 10.1097/01.ijg.0000131757.63542.24
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The trabecular meshwork (TM) is a specialized vessel wall located between the aqueous-containing anterior chamber of the eye and Schlemm’s canal (SC), a venous sinus that communicates with episcleral veins on the surface of the eye. Aqueous moves by bulk flow1 through the trabecular meshwork into SC (Fig. 1) down a pressure gradient initially set up by the heart. Mechanisms controlling aqueous outflow and intraocular pressure (IOP) reside in this region,2,3 as does the abnormality in open angle glaucoma.4 This report characterizes aqueous outflow as an active phenomenon driven by means of a mechanical pump.

(A) Aqueous outflow system appearance when intraocular pressure (IOP) is below episcleral venous pressure (EVP). (B) Change in aqueous outflow system appearance when intraocular pressure rises to physiologic level (IOP>EVP). In the aqueous outflow pump model, aqueous flows from the anterior chamber through the trabecular meshwork and SC valves into Schlemm’s canal. Schlemm’s canal endothelium (SCE) refers to portion of endothelium facing juxtacanalicular space. From: Johnstone MA. Pressure-dependent changes in configuration of the endothelial tubules of Schlemm’s canal. Am J Ophthalmol 1974;78:630–8. Copyright © (1974) AJO. All rights reserved.

In brief, the aqueous outflow pump receives power from transient increases in IOP, such as occur in systole of the cardiac cycle, during blinking and during eye movement (Fig. 2). These transient pressure spikes cause microscopic deformation in the elastic structural elements of the TM, juxtacanalicular cells, and SC inner wall endothelium (SCE) (Figs. 3 and 4). During systole, SCE moves outward into SC forcing aqueous into collector channel ostia and aqueous veins. At the same time, the IOP increase of systole forces aqueous into one-way collecting vessels or valves that span SC. When the pressure spike decays, the elastic elements respond by returning to their original configuration causing a relative pressure reduction in SC, thus inducing aqueous to flow from the valves into the canal. Pump efficiency is pressure sensitive, providing a mechanism to maintain IOP homeostasis. Laboratory and clinical evidence for this aqueous outflow model will be presented and parallels drawn with similar tissue pumps in other body systems.

Effect of the ocular pulse, eye movement and blinking on IOP. The continually changing pressure gradients transmit forces to aqueous outflow structures. From: Coleman DJ, Trokel S. Direct-Recorded Intraocular Pressure Variations in a Human Subject. Arch Ophthalmol 1969;82:637–40. Copyright © (1969) American Medical Association. All rights reserved.
Evidence for trabecular meshwork (TM) compliance (upper figures, light microscopy; lower figures, illustrations). (A) Intraocular pressure (IOP) < episcleral venous pressure (EVP). Living monkey eye. IOP is 0 and EVP is 8–9 mm Hg. Schlemm’s canal (SC) fills with red blood cells. Compression of SCE against underlying juxtacanalicular tissue in turn compresses the juxtacanalicular space against the trabecular lamellae of the trabecular meshwork (TM) almost obliterating the juxtacanalicular space. A rounded nuclear profile is visible (N). (B) (IOP = EVP). Hypotonous enucleated eye (IOP and EVP both = 0) or living eyes (IOP and EVP both = 9 mm Hg). Schlemm’s canal endothelium, juxtacanalicular tissue and trabecular lamellae are slightly separated from each other. SCE lacks evidence of cellular distension. (C) (IOP>EVP). Fellow eye of living monkey in A (eyes fixed simultaneously). IOP is 25 mm Hg and EVP is ∼8–9 mm Hg. Inner wall endothelium of SC distends into SC approaching external or corneoscleral wall (CSW). Schlemm’s canal narrows to a potential space and contains small amounts of plasma and red blood cells (RBC). SC endothelium distends to form pseudovacuoles “giant vacuoles”. Juxtacanalicular space is large and spacing between trabecular lamellae is wide. From: Johnstone MA, Grant WM. Pressure-dependent changes in structure of the aqueous outflow system in human and monkey eyes. Am. J. Ophthalmol 1973;75:365–83. Copyright © (1973) AJO. All rights reserved.
(A) Mechanism of Schlemm’s canal (SC) endothelium (SCE) attachment to underlying trabecular lamellae. Cytoplasmic processes of SCE attach to juxtacanalicular cell (JC) processes. Juxtacanalicular cell processes (JCC) in turn attach to trabecular lamellae (TL). Intertrabecular cell processes rather than collagen attachments limit excursions of adjacent trabecular lamellae. IOP-induced tissue loading (hollow arrows). (B) Region of scanning electron micrograph in (C) (white square). Numerous cytoplasmic processes arise from juxtacanalicular cells creating extensive attachments to the undersurface of SCE. (D) Appearance and relationships of SCE to juxtacanalicular cells and trabecular meshwork when IOP is low. (E) Appearance and relationships at physiologic IOP. SCE attachments to the underlying TM modulate SCE distention into SC. When IOP progressively increases, structural elements responsible for resistance to pressure respond by configuration changes as illustrated by the configuration change from D to E. Configuration changes illustrated in transition from D to E in response to IOP-induced tissue loading, place resistance to aqueous outflow at SCE. The juxtacanalicular space enlarges, extracellular matrix material density reduces and cell processes (CP) restraining the inner wall endothelium change from a parallel to perpendicular configuration. Signs of pressure-induced cell stresses are present at cell process origins where cytoplasm and nuclei of both juxtacanalicular cells and SCE undergo deformation toward respective processes; cytoplasm and nuclei of SCE undergo elongation and attenuation. Evidence of SC endolethial cell tethering by extracellular matrix material is absent. Trabecular lamellae, which participate in modulating and restraining SCE distention into SC progressively separate from one another as seen in Fig. 3. (A) From: Johnstone MA. The morphology of the aqueous outflow channels. In: Drance SM, ed. Glaucoma: Applied Pharmacology in Medical Treatment. New York: Grune & Stratton, 1984. “Copyright © (1984) Grune & Stratton. All rights reserved.” (D and E) From: Johnstone MA. Pressure-dependent changes in nuclei and the process origins of the endothelial cells lining Schlemm’s canal. Invest. Ophthalmol. & Vis. Sci. 1979;18(1):44–51. Copyright © (1979) IOVS. All rights reserved.

Characterization of flow controlled by a mechanical pump differs from the traditional view that aqueous movement through the outflow system is a passive process.5,6 However, structural and positional characteristics of the outflow system fulfill requisites for a mechanical pump. The first structural requisite of a pump, the presence of one-way valves, is fulfilled by collecting vessels or valves that span SC7 (Fig. 1). The second structural requisite, distention and recoil of tissue in response to small changes in IOP, is fulfilled by the trabecular meshwork8 (Fig. 2).

The trabecular tissues of the outflow system experience uninterrupted exposure to transient IOP changes (Fig. 3). Pressure transients include continuous oscillating or cyclic pressure changes from the ocular pulse9–11 and frequent transient pressure changes resulting from blinking and eye movement.12 These pressure gradient changes are sufficient to cause trabecular tissue excursions8 and are thus capable of driving an aqueous outflow pump.

Consistent with principles of biomechanics,13–18 this article examines in turn, the architecture or geometry of the outflow tissues, the composition of the tissues, laboratory evidence of the effects of tissue loading (in this case by IOP) and finally, the in vivo effects in humans of physiologic tissue loading by IOP transients. Tissue geometry considerations place emphasis on two features, first the attachment mechanism of SC endothelium to underlying trabecular lamellae (Fig. 4) and second the anatomy of Schlemm’s canal collecting vessels or valves (SC valves) (Fig. 1).

The article explores laboratory evidence that IOP-induced tissue loading acts at SCE. It emphasizes that at normal IOP, SCE experiences a continuous IOP-induced load distributed throughout the load-bearing trabecular meshwork, thus providing an intrinsic stabilizing tension (Fig. 4). This stabilizing tension permits finely tuned global responses of the TM tissues to cyclic and intermittent variations in load presented by changing IOP. Next, the article explores evidence that SC valves contain a lumen communicating with both the anterior chamber and SC. Responses to IOP-induced loads provide a mechanism to drive aqueous through the valve lumen to SC.

The article explores in vivo evidence of pulsatile flow into SC, into collector channels, and into aqueous veins in response to in vivo tissue loading by physiologic IOP-induced cyclic pressure transients in humans. Finally, the report explores functional biomechanics intrinsic to the model, including pumping and IOP regulatory mechanisms.


Schlemm’s canal endothelium attaches to the TM; otherwise, normal IOP would cause SCE to separate completely from the TM.19 Schlemm’s canal endothelial cells extend cytoplasmic processes into the juxtacanalicular space, as do the cells lining the trabecular lamellae. Juxtacanalicular cell processes attach to both SCE and trabecular-lamellae processes. Thus, juxtacanalicular cells through their processes provide a cellular linking mechanism between SCE and the trabecular lamellae8,20–25 (Fig. 4). Well-characterized robust desmosomes capable of sustaining cellular stress are present between cell process attachments.20,26 Such desmosomes attach to intracellular intermediate and actin support filaments27,28 enabling them to distribute stress throughout the cytoskeleton of involved cells23 of this tensionally integrated system16,29,30 (Fig. 4).

Trabecular lamellae adjacent to the wall of a vessel (SC) are analogous to the adventia of blood vessels that provide resistance to distention of the vessel wall and recoil in response to cyclic hydrodynamic loading. Because pressure gradients across the SC endothelial surface are higher on the abluminal side, trabecular lamellae require special adaptations to provide resistance to hydrodynamic loading. Like the adventia of other vessel walls, trabecular lamellae contain type I and III collagen providing structural support in tension and elastin that provides a recoverable response over large excursions. Organization and distribution of elastin in trabecular lamellae is similar to that found in tendons31 providing a mechanism for reversible deformation in response to cyclic hydrodynamic loading.8

Trabecular lamellae organize in parallel sheets with a circumferential orientation relative to SC32; intertrabecular collagen beams are hard to find19 (Figs. 1 and 4). Intertrabecular cell processes connect trabecular lamellae to one another19,26,32–36 via desmosomes.20 Cell processes of the endothelial cells on the trabecular lamellae are anchored to the underlying extracellular matrix (ECM) of the lamellae through integrins.37–39 The larger lamellae near the AC anchor at their ends to scleral spur and Schwalbe’s line8,26,35 while smaller trabecular lamellae close to the juxtacanalicular space attach to the larger lamellae by means of these interlamellar processes. Schlemm’s canal endothelium thus anchors to the underlying trabecular lamellae through a 3-dimensional force-distributing organization of cell process attachments (Fig. 4).


Discrete structural elements in a load-bearing network are capable of change in orientation and relative spacing to one another.16 In response to IOP-induced tissue loading, the following evidence demonstrates progressive pressure-induced distention of SCE into SC with the load distributed to the entire TM. As IOP increases, cell process reorientation from parallel to perpendicular relative to SCE occurs accompanied by a straightening of cell processes throughout the meshwork (Fig. 4).8,23,26,40 The juxtacanalicular space enlarges markedly as pressure increases8,21,23,26,35,40–43; enlargement may be as much as two to threefold.26 The resultant enlargement of the juxtacanalicular space reduces the density of both cellular elements and ECM materials.23,26,42 Progressively larger spaces develop between trabecular beams as the trabecular lamellae suspending SCE undergo progressive separation from one another in concert with progressive distention of SCE.8,20,21,23,35,40


Cellular and subcellular IOP-induced deformation provide evidence of resistance characteristics. Of special interest is evidence that deformation described in SC endothelial cells parallels mechanosensory and regulatory mechanisms modulating pressure and flow throughout the vascular system.13,16,30,44 Ultrastructurally, the endothelial and juxtacanalicular cell cytoskeleton that controls IOP-induced deformation is composed of microtubules,45–47 microfilaments,48–51 and intermediate filaments,52–57 the predominant cytoskeletal element of human trabecular meshwork cells.57,58

The SCE cell membrane and cytoplasmic contents progressively change shape from a spherical configuration in hypotony without an IOP-induced tissue load to an elongated plate-like configuration when subjected to an IOP-induced tissue load8,21,23,24,26,40 (Fig. 4). The entire nucleus shape including the nuclear membrane also undergoes a progressive change from a spherical to an elongated plate-like configuration with loss of nuclear folds when subjected to a progressive IOP-induced tissue load. A progressive IOP increase causes juxtacanalicular cell shape to change from spherical to stellate (Fig. 4). The IOP-induced juxtacanalicular cell configuration change also involves the cell membrane, the cytoplasm, the nuclear envelope, and the nuclear contents. At cell process origins of both SCE and juxtacanalicular cells, the cell membrane, the cytoplasm, the nuclear membrane, and the nuclear contents undergo a change in shape in response to a progressive increase in an IOP-induced tissue load (Fig. 4).23,24 The cellular shape changes described are reversible long after death8 and may be explained by the elastic properties and limited energy requirements of intermediate filaments.


Tissue geometry, composition, and deformation in response to systematic tissue loading permit identification of both resistance characteristics of tissue elements and integrated tissue responses.13,14 IOP is the loading force to which the outflow system normally responds. Accordingly, resistance as determined by IOP-induced tissue loading takes priority in the hierarchy of considerations related to resistance sites. The many responses to IOP-induced tissue loading at both the tissue and cellular level provide a body of evidence enabling development of an evidence-based functional model.

Evidence of tissue and cellular deformation in response to an IOP-induced load places TM resistance to IOP at SCE. The loading force of IOP acts at SCE causing a number of manifestations of SCE deformation at the cellular level. Tissue loading at SCE causes it to deform and to distend into SC lumen, but dynamic tension distributed throughout the entire load bearing elements of the TM restrains the distending wall of SCE. A resulting reorganization of juxtacanalicular cell shape, juxtacanalicular space, and trabecular lamellae follows. The cytoplasmic and nuclear shape changes at cell process origins of SCE and juxtacanalicular cells are reflective of the tensional integration that extends from the tissue to the cytoskeletal level.

IOP-induced deformation of cellular and tissue elements are progressive as IOP increases, providing a mechanism for graded responses.8,26,35,40 Comparison of the appearance without a load (0 IOP) to the appearance with a load as seen at physiologic IOP (Fig. 4) demonstrates that at physiologic IOP, SCE faces a continuous IOP-induced load.8,26,35,40 The continuous IOP-induced load provides a stabilizing tension or prestress.

Evidence from these tissue-loading experiments demonstrates IOP-induced tissue deformation that is not compatible with a hydraulic resistance in the juxtacanalicular space. As described above, progressive deformation of SCE, juxtacanalicular cells, and trabecular lamellae with increasing IOP occurs in concert with progressive enlargement of the juxtacanalicular space. Juxtacanalicular space enlargement causes cellular elements and ECM material to become less compact, progressively reducing the ability of the juxtacanalicular space to participate as a resistance element. Yet as IOP increases, resistance to aqueous outflow also increases,5,59–62 making the juxtacanalicular region an unlikely source of hydraulic resistance.43


Operating Microscope

Stegmann’s technique of unroofing SC in the course of non-penetrating filtration surgery,63 reveals SC valves as diaphanous cylindrical structures that contain a lumen and span the canal (Fig. 5). Schlemm’s canal valves undergo remarkable distention but eventually rupture when stretched by a probe or when SC walls are widely separated when viewed intraoperatively. A burst of aqueous discharges from the newly ruptured ends of disrupted SC valves.

Dissecting microscope. (A) (B) and (C) Illustration of effect of viscoelastic injected into Schlemm’s canal (SC) dilating the canal and collapsing the trabecular meshwork (TM) thus stretching SC valve across SC. Dissecting microscope light passes the full length of SC in each radial segment allowing analysis of SC valve frequency. Examination of the full circumference of SC demonstrates an average of two SC valves per mm. From: Smit & Johnstone 2002 (D) Human Eye. Schlemm’s canal valve (arrow from C to D) arises as funnel-shaped structure, bridges the lumen of SC as a semitransparent diaphanous structure and then inserts into the external or corneoscleral wall of the canal. (E) Courtesy Robert Stegmann. Operating room microscope-human eye. Corneoscleral wall (CSW) of SC is visible on undersurface of scleral flap dissected for deep sclerectomy/viscocanalostomy. SC cut ends indicated by small white arrows. Small black arrows indicate SC inner wall endothelium at origin of SC valve. Large white arrows point to SC valve. Upper structure attached to SC inner wall endothelium with disruption of distal attachment. Lower structure remains attached to CSW with SC inner wall attachment disrupted. From: Smit BA, Johnstone MA. Effects of viscoclastic injection into Schlemm’s canal in primate and human eyes: potential relevance to viscocanalostomy. Ophthalmology 2002;109(4):786–92. Copyright © (2002) Ophthalmology. All rights reserved.

Dissecting Microscope

Schlemm’s canal valves are easily viewed following viscoelastic dilation of SC (Fig. 5).64,65 Schlemm’s canal valves arise directly from SCE, coursing across the canal to attach to the external wall in the manner of a vessel.

Light Microscopy

Intraocular pressure reduction causes blood from the episcleral veins to reflux into SC. Blood reflux forces the TM toward the anterior chamber, collapsing it, at the same time dilating SC (Figs. 1 and 6). In the dilated canal, SC valves, which attach both to the TM and corneoscleral wall, then span almost directly across SC. In the widely dilated canal only a few histologic sections are necessary to characterize the appearance of SC valves (Fig. 6).7,66 Experiments that force the crystalline lens backward also cause SC valves to span directly across the canal (Fig. 7). 43

Light microscopy. Macaca mulatta (Rhesus) monkey eye fixed in vivo at 0 mm Hg IOP (episcleral venous pressure ∼8–9 mm Hg. Blood refluxes into Schlemm’s canal (SC) widely dilating the canal at the same time collapsing the trabecular meshwork (TM), separating it from the corneoscleral wall (CSW). (A) Serial histologic sections labeled A, B, C, and D encompass the length of Schlemm’s canal a valve that has a circumferential orientation within SC. Isometric drawing derived from placement of serial sections within the isometric grid. (B) In single histologic section, SC valve (inside white circle) appears as nondescript island of tissue. Large black arrow illustrates location of segment of SC valve marked “C” within the series of serial histologic sections in B. Endothelium lining walls of SC valves (ET) spanning across SC, (lu) lumen, primate red blood cells (prbc) refluxed into lumen of SC valves from SC lumen. From: Johnstone MA. Pressure-dependent changes in configuration of the endothelial tubules of Schlemm’s canal. Am J Ophthalmol 1974;78:630–8. Copyright © (1974) AJO. All rights reserved.
Light microscopy. Monkey eye. A device presses the crystalline lens backward, causing tension on the ciliary muscle via zonular attachments. Zonular attachments cause tension on scleral spur. Inward movement of scleral spur, through its attachment to the trabecular lamellae, causes Schlemm’s canal (SC) to dilate. SC valve (arrow) appears as a cord of highly cellular tissue, and spans between trabecular meshwork and external wall of SC near entrance of collector channel ostia (cc). From: Van Buskirk EM. Anatomic correlates of changing aqueous outflow facility in excised human eyes. Invest Ophthalmol Vis Sci 1982;22(5):625–32. Copyright © (1974) AJO. All rights reserved.

Amorphous material in SC valves has staining characteristics identical to the ECM in the juxtacanalicular space of the TM. Densely staining ECM material in SC valves is consistent with a greater concentration of ECM material within these structures than in the juxtacanalicular space; the higher concentration may present a hydraulic resistance in the SC valve lumen. The ECM presumably washes into the lumen of SC valves from the juxtacanalicular space where it has been extensively studied.37,67–73 A concentration of pigment is at times apparent in SC valves and is generally greater than that in the TM. The accumulation of pigment within SC valves7,65 further suggests that these structures are pathways for outflow of fluid and particulate material.

Scanning Electron Microscopy

Schlemm’s canal valve topographic relationships identified by scanning electron microscopy (SEM)66 confirm those described by light microscopy. Openings are visible at the distal end of SC valves (Figs. 8 and 9).

Scanning electron microscopy. Monkey eye-macaca mulatta. (A) Canal dilated by maintaining tension on scleral spur of 500-micron radial limbal section during fixation. Surface of endothelial lining (ET) of Schlemm’s canal valve spanning between trabecular meshwork (TM) and corneoscleral wall (CSW) is seen to be continuous with both inner wall endothelium of SC and outer or corneoscleral wall endothelium. (B) Region of opening at distal end of Schlemm’s canal valve in A. (C) Monkey eye-macaca nemistrina. Viscoelastic dilation of Schlemm’s canal (SC). Two valve-like structures (white arrows) span between trabecular meshwork and corneoscleral wall of SC. (cc) collector channel ostia. From: Smit BA, Johnstone MA. Effects of viscoelastic injection into Schlemm’s canal in primate and human eyes: potential relevance to viscocanalostomy. Ophthalmology 2002;109(4):786–92. Copyright © (2002) Ophthalmology. All rights reserved.
Scanning electron microscopy (SEM) Human eye. The surface of the endothelial lining (ET) of a SC valve spans between the trabecular meshwork (TM) and the corneoscleral wall (CSW) of Schlemm’s canal. The wall of the SC valve is continuous with the lining of the inner and outer wall of SC. (cc) collector channel, (AC) anterior chamber. An opening is visible at the distal end of the SC valve where it attaches to the external wall.

Transmission Electron Microscopy

Schlemm’s canal valves arise from the inner wall endothelium of SC and course across the canal to attach to the external wall as seen with transmission electron microscopy (TEM)66 (Fig. 10).66 The walls of SC valves are continuous with the walls of SCE. The lumen of SC valves is continuous with the juxtacanalicular space of the TM. Electron-dense material present in the valve lumen is like ECM material seen in the juxtacanalicular space.

Monkey eye-macaca mulatta. (A) The endothelial lining (ET) of the SC valve originates from the trabecular meshwork (TM) as a funnel-shaped region continuous with SCE along the inner wall. The lumen (Lu) of the SC valve is continuous with the juxtacanalicular space of the TM. The SC valve spans SC attaching at the distal end to the corneoscleral wall (CSW). Electron-dense extracellular matrix material is present in the lumen of the SC valve. Stellate cells with cytoplasmic processes are present in the valve lumen with characteristics like subendothelial cells (SEC) of the juxtacanalicular space of the trabecular meshwork. Collagenous supporting structures (CSS) are visible at the SC valve distal attachment to the corneoscleral wall with an appositionally closed potential space (small white arrows) between the two collagenous supporting structures. Bicuspid valves of the larger lymphatics have a similar arrangement. Avian red blood cells (arbc), intentionally introduced into the anterior chamber are visible in the juxtacanalicular space at the origin of the valve. A few arbc’s have passed through the SC valve lumen to the distal end. This figure represents a montage of five of the lowest power electron micrographs illustrating the difficulty in assessing the structures with transmission electron microscopy. (B) Cross section through a region of SC valve examined by TEM. The endothelial cells comprising the endothelial lining (ET) of the SC valve lumen enclose electron-dense extracellular matrix material. Stellate subendothelial cells with many cytoplasmic processes are present in the lumen. (C) Scanning electron microscopy with line indicated by x-y depecting cross section through valve seen in A.


Broad collagen sheets, attaching between the corneoscleral walls of SC, divide the canal into compartments at the entrance of collector channel ostia. At the dissecting microscope, septa appear as white collagen-like scleral collagen. Histologically septa are composed of dense collagen bundles continuous with identically staining collagen of the sclera (Fig. 11). Septa contain no lumen.

(A) Scanning electron microscopy (SEM) Macaca mulatta. Septum (SEP) arising from the corneoscleral wall (CSW) at the entrance of a collector channel (cc) that communicates with Schlemm’s canal (SC). Collagen bundles of the septum are continuous with the collagen bundles of the sclera. The cross section cuts through the septum so that in the center of the micrograph, the cross section ends (white arrows). The remaining portion of the septum passes out of the plane of the cut section before attaching to the corneoscleral wall and creates a cross section that on histologic examination looks like the septum in Fig. B. At the dissecting microscope, septa have a dense white appearance like the collagen of the sclera. (B) Septum-Light microscopy. Macaca mulatta. Schlemm’s canal dilates and fills with blood while the trabecular meshwork collapses. A large septum is present arising from the posterior wall of SC. Dense bundles of collagen continuous with the collagenous bundles of the sclera are present in the septum. Staining characteristics are identical to the staining characteristic of scleral collagen. Serial sections reveal that septa are sheets of collagen demarcating the entrance of collector channel ostia as illustrated in Fig. 10A. In contrast to aqueous valves, which always arise from the inner wall endothelium of SC, the collagenous structures regularly arise in the posterior portion of the canal from the region of the scleral spur and the corneoscleral wall. From: Johnstone et al.


Herniations are undisrupted hemispherical outpouchings or protrusions of the internal wall of SC8,74 that do not attach to SC external wall. Herniations are unrelated to SC valves that have a lumen, develop a cylindrical configuration, and span across SC where they attach to SC external wall. SC valves are disrupted when separating the wall of SC to examine the herniations by SEM.74


Monkey Red Blood Cell Tracers at Normal IOP (TM Flow-Enabling Configuration Maintained)

Maintaining a physiologic pressure of 25 mm Hg in vivo during fixation establishes the flow-enabling configuration. After fixation, in the still living animal, reduction of IOP causes blood to flow from the episcleral veins into SC as a tracer (Fig. 12). The technique results in uniform apposition of blood to SC endothelium. Intense staining of plasma by toluidine blue as well as the presence of red cells is an excellent tracer system to demonstrate openings in SC endothelium.75 Despite extensive examination, no blood or plasma crosses SC endothelium.

(A) Hypothesis. (B) Illustrations 1–3. Depiction of findings involving in vivo fixation for 30 minutes in 3 eyes and 60 minutes in one eye while maintaining 25 mm Hg IOP. Fixation then followed by in vivo reduction of IOP to atmospheric pressure. Macaca mulatta, findings in 4 eyes, 31 segments, 982 epon-embedded sections, encompassing 319, 475 u2 of SC endothelium. (C) Representative histologic section. In the flow enabling configuration cells of SCE are elongated and attenuated. The juxtacanalicular space of the TM is large. Primate red blood cells (prbc) have refluxed into Schlemm’s canal (SC) and are in uniform contact with Schlemm’s canal endothelium along the inner wall (el) lining of the canal. Toluidine blue stains plasma intensely providing an excellent tracer. No plasma or red blood cells cross the endothelial lining and the juxtacanalicular space of the trabecular meshwork (TM) is free of blood in all sections studied providing no tracer evidence of transcellular pores. Corneoscleral wall (CSW), light micrograph (×2050).

Monkey Red Blood Cell Tracers at Low Intraocular Pressure (IOP < EVP) Non-Flow Configuration

Lowering IOP below EVP in vivo before fixation establishes the non-flow configuration (Fig. 13). Because hydrostatic pressure in the canal is higher than in the AC, blood completely fills SC and collapses the TM. Primate red blood cells (RBCs) at times reflux into the lumen of the SC valves (Fig. 13), typically near the distal end adjacent to the SC valves external wall attachment. At times however, primate RBCs fill the entire length of the lumen of SC valves, some reaching the juxtacanalicular space at the origin of the SC valve lumen (Figs. 6 and 13). Primate RBCs refluxing from SC into the lumen of SC valves indicates the presence of a communication between the lumen of SC valves and the lumen of SC.

Monkey red blood cells (prbc) introduced into SC in living monkeys. Based on 1764 epon embedded sections from 44 segments in six macaca mulatta eyes. (A) Illustration depicting slow in vivo reduction of IOP to zero mm Hg before fixation. From left to right (A.1, A.2, A.3), primate red blood cells and plasma reflux from episcleral veins into collector channels and then into SC. Primate RBC’s enter the lumen of SC valves. (B) Two serial histologic longitudinal sections through a SC valve. The endothelial lining (ET) of the SC valve encloses a lumen. SC valves span Schlemm’s canal (SC) from the trabecular meshwork (TM) to the external or corneoscleral wall (CSW). Primate red blood cells are present within SC valve lumen and reflux from SC as far as the juxtacanalicular space of the TM. Collector channel ostia (cco) opens into a collector channel (cc). (C) Histologic cross-section through SC valve in which endothelial lining (ET) encloses the lumen of SC valve. Primate red blood cells (prbc) and a subendothelial cell (SEC) are present within the valve lumen. Endothelial cells comprising the walls of SC valves are thicker and their nuclei are more rounded with increased notches and folds compared with the appearance seen at physiologic pressure.

Avian Red Blood Cell Tracers in the Anterior Chamber in Monkey and Human Eyes

Avian RBCs are an excellent tracer because, unlike RBCs of primates, they contain a nucleus that stains intensely. Avian RBCs introduced into the AC enter SC valves, at times filling the lumen, indicating a free communication between the anterior chamber and SC valves (Figs. 10, 14, and 15).

Avian red blood cells (arbc) introduced into AC in living monkeys. (A.1) Avian red blood cell infusion into anterior chamber and trabecular meshwork, followed by arbc entry into SC valves (A.2). (B.1) Initial IOP reduction below episcleral venous pressure to allow blood to reflux into SC, thus dilating the canal. (B.2) Progressive reduction of IOP below episcleral venous pressure causes progressive canal dilation, straightening and stretching SC valves between SC walls. (C) Illustration of how radial histologic sections from B.2 now permit longitudinal sections through length of valve illustrated in Fig. 15. From: Johnstone et al.
Avian red blood cell (arbc) tracers introduced into anterior chamber of living monkey eyes by protocol in Fig. 14. Experiments in two macaca mulatta eyes. Unlike nucleus-deficient primate RBC’s, avian RBC’s have darkly staining nuclei, and therefore provide a good tracer. (A,B,C,D) From left to right representative serial histologic sections encompass the entire width of SC valve as depicted in Fig. 14C. The endothelial lining (ET) of the SC valve is continuous with the inner wall endothelium of the trabecular meshwork (TM) and spans across Schlemm’s canal (SC) to attach to the corneoscleral (CSW). Avian red blood cells (arbc’s) are present in the trabecular meshwork, the juxtacanalicular space and fill the lumen of the aqueous valve in the central section through SC valve in C. Note the two collagenous supporting structures (css) at SC valve distal end with a narrow space where they meet (white arrows). (E) Enucleated human eyes. Avian red blood cells infused into anterior chamber of two eyes. Radial segments of limbus stretched by tension on scleral spur to dilate canal. Avian RBC’s are in SC valve lumen as in the monkey eye. From: Johnstone et al.

Anatomic Correlates to Schlemm’s Canal Valves

Direct return of another extravascular fluid, lymph, to the venous system is by means of an extrinsic pumping mechanism involving valves76 rather than intracellular or transcellular pores. Venous blood returns through an extrinsic pumping mechanism77,78 involving valves. Cerebrospinal fluid returns directly to the vascular system via valves in the model of Welsh.79 Aqueous returns directly to the venous system in the canine eye as demonstrated by Van Buskirk80 (Fig. 16).

Canine eye. Scanning electron microscopy of casts of the outflow system. Trabecular meshwork (a) connects collecting vessel (b) with vein of Hovius (c) present in the corneoscleral wall (d). The vein of Hovius is analogous to Schlemm’s in the primate eye. Rigid sclera surrounds the collecting vein and Hovius vein; therefore casting is possible. Van Buskirk’s work is extremely important because it clearly demonstrates that aqueous humor discharges directly into the vascular system by way of collector vessels. A minor modification in the primate eye is the traverse of the collecting vessel across the venous sinus (Schlemm’s canal), before discharge of aqueous. By virtue of this modification, collecting vessels (SC valves), are capable of behaving as a one-way valvular device. Van Buskirk EM. The canine eye: The vessels of aqueous drainage. Invest Ophthalmol 1979;18:223–30. Copyright © (1979) IOVS. All rights reserved.


Laboratory evidence of pressure-sensitive TM tissues and SC valves, coupled with clinical knowledge of constant pressure gradient changes within the eye predicts pump-like behavior of the outflow system. Identification of in vivo correlates provides a measure of the robustness of the laboratory-derived aqueous outflow pump model. A recent search of the literature related to pulsatile flow in aqueous veins provides evidence consistent with the prediction of an aqueous outflow pump mechanism, as does recent clinical evidence of pulsatile flow into collector channels and SC.

Pulsatile Aqueous Flow into Aqueous Veins

Pulsatile aqueous discharge into the aqueous veins is well characterized.12,81–88 For example, Kleinert observes 196 pulsatile aqueous veins in 111 normal eyes, and in each of 18 eyes with a large ocular pulse resulting from aortic regurgitation.87 Pulsatile aqueous flow is synchronous with the cardiac pulse.81,83–85,88–90 Interestingly, Ascher observes that “each systolic wave reaching the eye will increase IOP, the pressure increase is translated to SC, and the contents of the canal can give way in only one direction ... Schlemm’s canal will then be rhythmically filled with more fluid.”12 Ascher points out that the observations can not be easily reconciled with the traditional aqueous outflow model as a rigid, passive structure, which would not allow rapid translation of pressure gradients across the TM to SC12; as a result, these observations have languished for over 50 years.

Vries characterizes aqueous vein pulsation in remarkable detail.89 Tributaries of episcleral veins join aqueous veins creating a mixing vein (Fig. 17). Linear stratification occurs in the mixing vein because of differences in viscosity and specific gravity.91 During systole a rapidly moving modified parabolic wave of aqueous originating from the aqueous vein enters the mixing vein and the aqueous stratum widens. As the systolic wave dissipates during diastole, venous blood enters the mixing vessel causing a narrowing of the aqueous stratum.

Change in stroke volume with IOP increase following drinking of 1 liter of H2O. Fifty-nine-year-old Caucasian male with low-tension glaucoma. (A) and (B) Appearance before H2O drinking (IOP 11) and 2 hours after H2O drinking (IOP 11). (C) and (D) Representative appearance 30 minutes following H2O drinking (IOP 13). (E) and (F) Appearance 60 minutes after H2O drinking (IOP 16). (A) and (B) are areas indicated by green box in (C). Aqueous enters base of aqueous-episcleral vein mixing system (black arrows) and small amount of aqueous discharges into episcleral vein with each systole (dashed arrow). (C) and (D) With each systolic stroke, aqueous continues to fill the episcleral vein in A and B, but in addition, aqueous travels up the episcleral mixing vein (black arrows) to fill more than half its length with aqueous during systole. (E) and (F) Areas of aqueous mixing vein and episcleral vein in A–D fill with aqueous. Each systolic stroke has sufficient volume to drive a column of aqueous as far as a more distal episcleral vein, recreating a new vasculature as a mixing vein, thus increasing the aqueous discharge field. Linear stratification in the newly recruited episcleral vein (black arrows) oscillates between blood and clear aqueous with the cardiac pulse. The recruited mixing vein passes under an episcleral vein between the black and white arrows. A far greater stroke volume and more forceful pulse wave is present with the higher IOP of (E) and (F). The greater stroke volume is illustrated by the oscillating pulse wave passage to the end of the visible episcleral mixing vein in (F) (white arrows) in contrast to passage only to the base the aqueous-episcleral mixing vein system in (A) and (B).

Pulse wave origination from an aqueous source rather from episcleral venous pulsation is clear from the character of the rapidly moving aqueous pulse wave. The aqueous pulse wave starts at the entrance of the emissary aqueous vessels in the sclera and displaces the slower moving blood in the mixing vessels as it advances.89 Influx of aqueous resulting from the moving aqueous wave front in systole occurs more rapidly than the influx of blood during diastole.89 Compression of recipient episcleral veins, which precludes a retrograde venous pulse, causes an increase in pulsations.81,83,84 Furthermore, a complex pulsatile parabolic wave of aqueous originates at the aqueous vein scleral emissary. The complex parabolic wave propagates in aqueous and mixing veins at the same time movement of blood in adjacent episcleral veins is much slower and pulsatile behavior is absent. Compression of the homolateral carotid artery causes arrest of pulsations.90 Enhanced pulsatile aqueous discharge results from increasing IOP caused by pressure on the eye83,84 or water drinking92 (Fig. 17).

Pulsatile Aqueous Flow into Collector Channels

Stegmann uses an operating room microscope with 80-power magnification and high-resolution videographic capability to document his microsurgery techniques.63 Gonioscopically, he observes pulsatile aqueous flow into collector channels92 by compressing episcleral veins with the flange of a goniolens to cause slight blood reflux into collector channels; aqueous in the collector channels is blood-tinged permitting visualization of aqueous movement. Synchrony of pulsations with the cardiac pulse is evident because head movement and associated image movement is apparent with each cardiac cycle.

Pulsatile Aqueous Flow into Schlemm’s Canal

Circumferential flow of refluxed blood in SC is observed by Stegmann.92 In Stegmann’s videotapes of eyes with circumferential blood flow, I observed that circumferential flow results from cyclic ejection of a column of clear fluid into the blood in SC (Fig. 18).92 The pattern of clear fluid mixing with blood in SC is explained by aqueous entry into the lumen of SC since aqueous is the only clear fluid available to induce the phenomenon. The ejection of aqueous into SC occurs in synchrony with the cardiac pulse. The ejected aqueous column in SC initially appears as a complete clear column against a background of blood followed by turbulent swirling aqueous eddies and subsequent replacement of the aqueous with blood before ejection of the next aqueous column.

Courtesy Robert Stegmann. Gonioscopic view of anterior chamber angle of 6-year-old African female without glaucoma under anesthesia before undergoing cataract surgery. Episcleral venous pressure raised by creating pressure on episcleral veins opposite to goniolens mirror resulting in filling Schlemm’s canal with blood. Representative images from video sequence at 30 frames per second encompassing pulsatile aqueous discharge sequence during cardiac cycle. Intervening segments provide appearance of smooth transition clearly outlining funnel and cylindrical structures. Synchrony with cardiac pulse is apparent; at high magnification cardiac pulse-induced head movement causes corresponding image movement. Blood is visible in Schlemm’s canal above the base of the iris. (A) Beginning of systole. Funnel base (fb) and iris marking provide reference points. Funnel apex (fa) faintly seen below arrow (fa) with minimal evidence of clear aqueous. Clear aqueous not apparent in proximal cylindrical area (pca) or distal cylindrical area (dca). Turbulent bolus of clear aqueous swirling at site of aqueous ejection column (aec). (B) Systole. Funnel apex has moved upward and contains aqueous. Proximal cylindrical area contains aqueous. Aqueous ejection column (aec) dissipating. (C) Early diastole. Funnel apex aqueous decreased, proximal cylindrical area aqueous increased. Aqueous ejection column in Schlemm’s canal barely visible. (D) Late diastole. Distal end of cylindrical area contains aqueous and aqueous ejection column in SC is maximal. Faint outline of clear aqueous in funnel and proximal cylinder has shifted downward. Structure has straightened slightly compared with shape in B during systole. The cycle begins again with A.

The origin of the aqueous column in SC is a clear funnel of aqueous originating proximally with a base at the level of the posterior TM just above the scleral spur. The funnel apex extends upward toward Schwalbe’s line. The funnel of aqueous is continuous with a more distal clear cylindrical area that curves so that a portion of the cylinder runs circumferentially in SC. The recurring pattern of aqueous ejection into SC originates from the distal cylindrical area. The same phenomenon occurs consistently in the 30 cardiac cycles recorded. The salient features are present at each cycle, with the appearance varying slightly from cycle to cycle, never appearing exactly the same.

The sequence of aqueous propulsion originates when the funnel of aqueous enlarges rapidly with enlargement progressing from base to apex. Following complete filling the funnel becomes progressively smaller beginning at the base and progressing to the apex; the clear funnel of aqueous completely collapses during some cycles. As the funnel-shaped area becomes smaller during diastole the more distal cylindrical area progressively enlarges with the enlargement starting at the apex of the funnel creating the appearance of a pulse wave of aqueous progressively entering the cylindrical region from the funnel.

The cylindrical area containing aqueous enlarges in a progressive fashion beginning proximally at the apex of the funnel until the cylindrical area achieves the appearance of a complete column. Reduction in size of the cylindrical area starts proximally at the apex of the funnel before enlargement of the distal cylindrical area is completed. Progressive reduction in size of the aqueous containing cylinder from the proximal to distal region culminates in the ejection of aqueous into the blood-filled SC. The cylindrical region collapses at completion of the ejection phase.

Aqueous is constrained to a specific path as it flows into SC in a propulsive wave. The funnel-shaped and cylindrical areas define a lumen that changes shape in synchrony with the cardiac cycle. The constraining structures appear highly compliant because the funnel-shaped and cylindrical areas rapidly undergo a change in shape in response to IOP transients. The pattern of flow suggests that the structural tissues guiding flow accommodate or aid in causing the propulsive wave. The size, shape, and movement of aqueous through the structures are consistent with behavior predicted from the laboratory characterization of SC valve geometry7,66 and responses of both SCE8,21,26 and SC valves7,23,24,66,93,94 to tissue loading induced by pressure gradient changes.


The proposed mechanism is based first on consideration of laboratory evidence of tissue geometry, tissue composition, tissue loading responses, and tracer studies. Second, the laboratory evidence is correlated with in vivo evidence of pulsatile flow into aqueous veins from collector channels, into collector channels from SC, and into SC from the AC. Third, a series of animations depict various scenarios of tissue movement and aqueous flow patterns during systole and diastole92 that place constraints on possible alternatives.

At the homeostatic setpoint or attractor state,30,95 the entire TM faces a mean IOP-induced load that results in tensional integration of structures throughout the meshwork. During systole, increasing IOP causes SCE endothelium to distend further outward into SC inducing further tension on the trabecular lamellae. Movement of SCE into the canal causes a rise in SC pressure. Aqueous cannot pass backward toward the anterior chamber because SC valves and their lumens suspended in the canal experience a pressure as high as that in the canal. Because aqueous can move only one way, increasing pressure in SC with systole causes pulsatile discharge of aqueous into the collector channels and aqueous veins. Outward movement of SCE enlarges the funnel at the entrance of SC valves causing the funnel portion of the valves to fill with aqueous at the same time aqueous is being forced out of SC by the outward-moving SCE.

During diastole, IOP decreases and the IOP-induced pressure load on SCE and the trabecular lamellae also decreases. Elastic energy stored in the trabecular tissues during systole induces elastic recoil in diastole. Continuing recoil of the trabecular lamellae and SCE reduces SC pressure causing aqueous to enter SC from SC valves, representing a form of diastolic “suction”. In this model, the pressure gradient across SCE need not be large. Because elastic recoil of the TM causes diastolic filling of SC from SC valves, at the initiation of the next systole the end diastolic volume of SC is sufficiently large that pressure gradients across SCE are small or absent. Accordingly, during the next systole, the outward movement of SCE caused by the systolic IOP increase also causes a corresponding increase in SC pressure, effectively pushing fluid out of the canal without requiring a large pressure gradient across the wall of SCE. Similarly, SC valve transmural pressure gradients are modest due to the Starling resistor effect in a collapsible tube connected to 2 reservoirs and enclosed in a chamber with changing pressures.15,96,97

Tissue Biomechanics Optimize Short-Term Intraocular Pressure Regulation

Relationships of tension to length and stress to stretch are common to many soft tissues, are prevalent throughout the cardiovascular system,13 and are present in the aqueous outflow system.98 In this aqueous outflow model, at the homeostatic IOP setpoint, trabecular tissue distention is at the lower end of its length-tension curve. Systole induces modest additional tension and correspondingly with a fall in IOP during diastole, modest recoil occurs leading to limited SC filling. If IOP rises beyond the homeostatic setpoint, the trabecular tissues move up the length-tension curve. A rise in IOP during systole induces greater tension. The resulting recoil during diastole is more forceful increasing the aqueous volume entering SC. During the next systole, additional aqueous is discharged from SC thus increasing stroke volume. The increased stroke volume in turn reduces IOP, returning the trabecular tissues to their homeostatic length-tension relationship92 (Fig. 17).

Aqueous Outflow and Vascular System Parallels

Aqueous entry to the AC and egress from the AC to SC occurs down a pressure gradient set up by the heart. Principles of cardiac hydrodynamic tissue loading followed by recoil of elastic elements is a physiologic mechanism extensively employed throughout the cardiovascular system.13,78,99 Tissue loading and recoil begins in the heart where ventricular filling during diastole is initially rapid because the ventricle produces a diastolic “suction” as the extracellular elements of the myocardium recoil elastically from their contracted systolic configuration.13,78,99 A feature of the large elastic arteries is that the stretch during systole results in elastic recoil during diastole augmenting the smooth flow of blood.13,78,99 Left coronary artery flow occurs primarily during diastole due to the elastic recoil of the systolic pressure-distended aorta.13,78,99 Skeletal muscle contraction followed by elastic recoil of extracellular elements of the contiguous muscles and vein wall creates a diastolic suction driving the skeletal muscle pump in veins.78,99 Similarly, non-muscular lymphatic vessels are in series with the muscular vessels permitting extrinsic propulsion of lymph in which pumping is associated with recoil of the vessel walls in response to changing pressure transients.78 End diastolic volume and stretch regulate stroke volume in the larger lymphatics, thus linking stroke volume to pressure in the lumen.78


Mechanotransduction Mechanisms Couple Intraocular Pressure and Flow to Regulatory Pathways

Intrinsic regulation of vascular wall structure is the primary mechanism optimizing vascular wall stress to control pressure and flow. Extrinsic forces such as neural and hormonal factors then secondarily augment pressure and flow regulation. Through biomechanical coupling or mechanotransduction, cells alter their functional characteristics in response to externally applied loads.18 The first systematic tissue-loading studies of the aqueous outflow system8 demonstrated a tight biomechanical coupling between IOP and TM architecture, a coupling emphasized in subsequent reports (Fig. 4).7,23,24,66,93,94 The coupling of IOP and TM architecture led us to predict that SCE and associated resistance elements of the TM “behave functionally as a pressure-sensing and pressure-regulatory arrangement.”8 Since that time understanding of biomechanical coupling principles has led to a unifying model of intrinsic pressure and flow regulation in the vascular system.13,100 Shear and wall stresses integral to the aqueous pump model predict the presence of parallel intrinsic regulatory pathways in the aqueous outflow and vascular regulatory systems.

Limitations of the Passive Aqueous Outflow Model

Pressure and flow-mediated biomechanical coupling provides a regulatory framework in the vascular system, yet in the literature of the aqueous outflow system, no analogous unifying regulatory framework has been proposed.101 The prevailing paradigm in the aqueous outflow system literature posits a sufficiently rigid syncytium of ECM material in the juxtacanalicular space to sustain a steady passive resistance that regulates both pressure and flow.5,6,102,103 The paradigm precludes biomechanical coupling because resistance resides in a syncytium of ECM material. The pressure gradient dissipates on passage through the ECM material and flow occurs across low- resistance pores in SCE.104–106 The ECM resistance paradigm precludes both meaningful tissue loading at SCE to create wall stress and insufficient flow rates to induce shear stress.101 Pore frequency in SCE markedly decreases or is absent when fixation duration is short106 (eg, 30- or 60- minute in vivo fixation duration (Fig. 12)) suggesting that pores result from fixation.106 In the aqueous outflow pump model all aqueous flow is through SC valves.

Unifying Principles of Mechanotransduction Govern Optimization of Aqueous and Blood Flow

Intrinsic trabecular and vascular endothelial cell deforming forces that permit biomechanical coupling or mechanotransduction include stresses perpendicular to the wall lumen called normal or wall stress and those tangential to the lumen called shear stress, while intermediate vectors cause torsional stress.13,15,78 Force-dependent changes in cell surface receptors and cytoskeletal scaffold geometry transduce the forces into biochemical responses.44,107 Endothelial cells sense pressure and flow, transduce the flow through biomechanical linkage, and then initiate responses that effect changes in cellular and extracellular structural components of the walls of the outflow system and vasculature.13 In the aqueous pump model, wall stress, shear stress (resulting from high flow in SC valves and at their discharge sites in SC), and oscillatory stresses that induce tissue and cellular deformation are integral to function. Unifying principles of intrinsic pressure and flow regulation identified in the systemic vascular system13 predict comparable regulatory pathways in the aqueous outflow system. The presence of any such aqueous outflow system regulatory pathways strengthens the model.

The unifying pressure regulatory principle of vascular biomechanics is that vessel walls adapt to maintain a local wall shear stress set point that is a function of the optimal local transmural pressure.13,108,109 Vascular wall stress (tension/unit thickness) is similar from capillaries to the aorta.78 Vessel radius is determined by wall stress and vessel radius alterations impact both transmural pressure and flow.78 Minimizing work cost of blood transport with respect to radius has led to an evolutionarily optimized vessel radius.110,111 Shear stress resulting from flow governs lumen radius and causes adaptive changes in wall thickness, changing tissue mass and volume.13 Different energy cost functions apply depending on vessel location and function, but once differentiated to serve that function, vessels maintain constant shear stress that in turn maintains local transmural pressure relationships.78,108

Cellular Biomechanics Optimize Long-Term Intraocular Pressure Regulation

Mechanisms of tissue biomechanics allow the aqueous outflow pump to undergo short-term pressure-dependent stroke volume changes to maintain short-term homeostasis. However, long- term homeostasis requires modulation of the cellular biomechanics that define pump performance. Wall stresses modulate cellular biomechanics by intrinsic regulation of cellular constituents of the load-bearing tissues. In the aqueous pump model, load-bearing elements control mean lumen size as well as cyclic pressure-induced tissue distension and recoil. Cellular load-bearing constituents include SCE, juxtacanalicular cells, the endothelium lining the lumen of SC valves, and endothelium lining trabecular lamellae. Extracellular load-bearing constituents include the glycocalyx78,112–119 and ECM of the tendon-like31 trabecular lamellae.

Trabecular and vascular endothelial cells are mechanosensors13,44,78 that through mechanotransduction direct higher-level vessel wall self-organization13,44,78 focused around optimization of wall and shear stress. Pressure and shear stress-mediated signals in endothelia initiate a remarkable array of responses at the cellular, molecular, and genetic levels, causing both rapid responses and slow adaptive changes that regulate pressure and flow.13,44,78,120 These processes are not linear but are part of a highly complex interactive network44 in which an alteration in any component requires a contemporaneous adjustment of numerous other components in an iterative fashion described by Boolean networks.30,95 Shear and wall stress serve as evolutionarily optimized Boolean attractor states108,110 defining the regulatory end point (homeostatic setpoint) toward which all inputs are directed.

Regulatory Pathways Mediated by Mechanotransduction of Pressure and Flow

In the aqueous outflow pump model, the outflow tissues are characterized as a specially adapted vascular wall experiencing shear and wall stress like vasculature elsewhere. Pressure regulatory networks in the systemic vasculature are governed by universal principles of biomechanical coupling.16,30,120,121 Several reports show that TM cells can sense mechanical stretching forces,101,122–127 but regulatory mechanisms are not understood.101 However, in the aqueous pump model universal principles that organize pressure and flow-mediated regulatory behavior in the systemic vasculature44,120 provide a unifying framework for interpretation of regulatory behavior identified in the aqueous outflow system literature.

Shear stress changes cause endothelial cells to reorient in the direction of flow and involves rearrangement of the cell membrane, cytoskeletal elements, nuclear location, organelle rearrangement including the microtubule organizing center and Golgi, changes in focal adhesion alignment, and cell stiffening.120 Intermediate filament reorientation begins within seconds and actin within minutes of the onset of increased shear stress.120 Intercellular adhesion molecules in cell junctions such as the occludin/ZO-1 complex adapt their structure concurrently with the actin cytoskeleton.120,128,129

Shear stress responses include activation of stretch sensitive ion channels, inositol phosphate, diacylglyerol, and G proteins.120 An increase in cell turgor by aquaporin78,130,131 alters surface topography and cytoskeletal prestress16 thus altering shear120 and wall stress responses.16 Additional shear stress regulatory pathways include the GTPase Rho involved in cytoskeletal reorganization132–136 and Raf,137 kinases such as protein kinase C,138,139 FAK,140,141 Ikappa B,142 and map kinases (ERK, JNK, P38, BMK-1).139 Shear stress also modulates transcription factor families such as c-fos, NFKB, and AP-1.143–145

Wall pressure and shear stress changes in endothelial cells induce alterations in integrin attachments to the load-bearing ECM elements.120 In the case of the TM, SCE stresses transmit through cell processes to the endothelium lining the trabecular lamellae and to their corresponding basement membrane via integrins. In endothelial cells exposed to oscillatory shear stress or hydrostatic pressure, fibronectin fibrils increase; there is also a clustering of alpha-5 beta-1 integrin receptors and focal adhesion proteins.146

Shear and pulsatile stretch mediated through integrins regulate ECM deposition147 that alters load-bearing characteristics of the vascular wall.44,107 Stress-mediated signals alter NO and ET-1 release,13 alter growth factors such as FGF, PDGF, VEGF,13,148 as well as metalloproteinases149–153 and TIMPs.151 Shear stress responses that regulate the glycocalyx154–156 (a luminal negatively charged fiber matrix composed of heparin and chondroitin/dermatan sulfate112,119,157) maintain normal resistance characteristics of endothelium ensuring retention of normal load-bearing properties.

Shear stress and inflammation share regulatory pathways because shear stress changes represent an insult that initiates inflammatory cell adhesion13 and shear stress plays a protective role in vascular homeostasis by inhibiting endothelial responses to cytokine stimulation.158 Inflammation in turn causes endothelial lining changes that alter shear stress as a means to allow cell adhesion.13 Inflammatory pathways that perturb optimized setpoints of shear stress response networks include cytokines such as interleukins, TGFB, and TNF alpha13,158 as well as inflammation-related adhesion molecules such as E selectin (ELAM),159,160 CD44,161 VCAM-1,162–164 and MCP.158,165,166 The sterol response element exhibits shear stress responses165 comparable to high-dose glucocorticoid exposure167 providing a mechanism to link shear stress and steroid responses in trabecular tissues.125,168–172


This report examines evidence supporting a model of the aqueous outflow system as a mechanical pump. Laboratory evidence demonstrates the presence of valves in SC. Elastic and contractile properties of the TM and SC valves permit pressure transients to cause pulsatile fluid movement through the outflow system. Clinical evidence of pulsatile flow into SC, from SC into the collector channels, and from aqueous veins into episcleral veins supports the model. In this model alterations in the stroke volume of aqueous discharged to the episcleral veins maintain short-term homeostasis. Mechanotransduction through biomechanical coupling of IOP, flow and wall stresses optimizes tissue biomechanics resulting in long-term homeostasis.


1. Bill A, Maepea O. Mechanisms and routes of aqueous humor drainage. In: Albert DM, Jacobiec FA, eds. Principles and Practice of Ophthalmology. Philadelphia: W.B. Saunders; 1994:206–225.
2. Grant WM. Further studies on facility of flow through the trabecular meshwork. Arch Ophthalmol. 1958;60:523–533.
3. Johnstone MA, Grant WM. Microsurgery of Schlemm’s canal and the aqueous outflow system. Am J Ophthalmol. 1973;76:906–917.
4. Grant WM. Experimental Aqueous Perfusion in Enucleated Human Eyes. Arch Ophthalmol. 1963;69:783–801.
5. Kaufman PL. Pressure-dependent outflow. In: Ritch R, Shields MB, Krupin T, eds. The Glaucomas. St. Louis: Mosby; 1996:307–333.
6. Millar JC, Gabelt BT, Kaufman PL. Aqueous humor dynamics. In: Tasman W, Jaeger EA, eds. Duane’s Clinical Ophthalmology. Philadelphia: Harper & Row; 2002:1–34.
7. Johnstone MA. Pressure-dependent changes in configuration of the endothelial tubules of Schlemm’s canal. Am J Ophthalmol. 1974;78:630–638.
8. Johnstone MA, Grant WM. Pressure-dependent changes in structure of the aqueous outflow system in human and monkey eyes. Am J Ophthalmol. 1973;75:365–383.
9. Coleman DJ, Trokel S. Direct-recorded intraocular pressure variations in a human subject. Arch Ophthalmol. 1969;82:637–640.
10. Hedges TR, Baron EM, Hedges TR, et al. The retinal venous pulse: Its relation to optic disc characteristics and choroidal pulse. Ophthalmol. 1994;101:542–547.
11. Phillips CI, Tsukahara S, Hosaka O, et al. Ocular pulsation correlates with ocular tension: the choroid as piston for an aqueous pump? Ophthalmic Res. 1992;24:338–343.
12. Ascher KW. The Aqueous Veins. Vol. 1. Springfield: Charles C. Thomas; 1961.
13. Humphrey JD. Cardiovascular Solid Mechanics: Cells, Tissues, and Organs, 1st ed. New York: Springer-Verlag; 2002.
14. Sramek BB, Valenta J, Klimes F. Biomechanics of the Cardiovascular System, 1st ed. Prague-Irvine: Czech Technical University Press; 1995.
15. Fung YC. Biomechanics: Circulation. New York: Springer-Verlag; 1996.
16. Ingber DE. Tensegrity I. Cell structure and hierarchical systems biology. J Cell Sci. 2003;116:1157–1173.
17. Wendling S, Canadas P, Chabrand P. Toward a generalised tensegrity model describing the mechanical behaviour of the cytoskeleton structure. Comput Methods Biomech Biomed Engin. 2003;6:45–52.
18. Wendling S, Canadas P, Oddou C, et al. Interrelations between elastic energy and strain in a tensegrity model: contribution to the analysis of the mechanical response in living cells. Comput Methods Biomech Biomed Engin. 2002;5:1–6.
19. Flocks M. The anatomy of the trabecular meshwork as seen in tangential section. Arch Ophthalmol. 1957;56:708–718.
20. Grierson I, Lee WR. Junctions between the cells of the trabecular meshwork. Albrecht Von Graefes Arch Klin Exp Ophthalmol. 1974;192:89–104.
21. Grierson I, Lee WR. Pressure-induced changes in the ultrastructure of the endothelium lining Schlemm’s canal. Am J Ophthalmol. 1975;80:863–884.
22. Grierson I, Lee WR, Abraham S, et al. Associations between the cells of the walls of Schlemm’s canal. Albrecht Von Graefes Arch Klin Exp Ophthalmol. 1978;208:33–47.
23. Johnstone MA. Pressure-dependent changes in nuclei and the process origins of the endothelial cells lining Schlemm’s canal. Invest Ophthalmol Vis Sci. 1979;18:44–51.
24. Johnstone MA. The morphology of the aqueous outflow channels. In: Drance SM, ed. Glaucoma: Applied Pharmacology in Medical Treatment of Glaucoma. New York: Grune & Stratton; 1984:87–109.
25. Lutjen-Drecoll E. Functional morphology of the trabecular meshwork in primate eyes. Prog Retin Eye Res. 1998;18:91–119.
26. Grierson I, Lee WR. The fine structure of the trabecular meshwork at graded levels of intraocular pressure. (1) Pressure effects within the near-physiological range (8–30 mmHg). Exp Eye Res. 1975;20:505–521.
27. Alberts B, Bray D, Lewis J, et al. The cytoskeleton. In: Alberts B, Bray D, Lewis J, et al., eds. Molecular Biology of the Cell. New York: Garland Publishing, Inc.; 1994:788–847.
28. Darnell J, Lodish H, Baltimore D. Actin, myosin and intermediate filaments: Cell movements & cell shape. In: Darnell J, Lodish H, eds. Molecular Cell Biology. New York: WH Freeman & Company; 1990:859–899.
29. Ingber DE. Integrins, tensegrity, and mechanotransduction. Gravit Space Biol Bull. 1997;10:49–55.
30. Ingber DE. Tensegrity II. How structural networks influence cellular information processing networks. J Cell Sci. 2003;116:1397–1408.
31. Hernandez MR, Gong H. Extracellular matrix of the trabecular meshwork and optic nerve head. In: Ritch R, Shields MB, Krupin T, eds. The Glaucomas, 2nd ed. St. Louis: Mosby; 1996: v. 1.
32. Hogan MJ, Alvarado J, Weddell JE. Histology of the Human Eye, and Atlas and Textbook. Philadelphia: Saunders; 1971.
33. Holmberg AS. Schlemm’s canal and the trabecular meshwork. an electron microscopic study of the normal structure in man and monkey (Cercopithecus Ethiops). Documentia Ophthalmologica. 1965;29:339–373.
34. Inomata H, Bill A, Smelser GK. Aqueous humor pathways through the trabecular meshwork and into Schlemm’s canal in the cynomolgus monkey (Macaca irus). An electron microscopic study. Am J Ophthalmol. 1972;73:760–789.
35. Grierson I, Lee WR. The fine structure of the trabecular meshwork at graded levels of intraocular pressure. (2) Pressures outside the physiological range (0 and 50 mmHg). Exp Eye Res. 1975;20:523–530.
36. Raviola G. Effects of paracentesis on the blood-aqueous barrier: An electron microscope study on Macaca mulatta using horseradish peroxidase as a tracer. Invest Ophthalmol. 1974;13:828–858.
37. Zhou L, Zhang SR, Yue BY. Adhesion of human trabecular meshwork cells to extracellular matrix proteins. Roles and distribution of integrin receptors. Invest Ophthalmol Vis Sci. 1996;37:104–113.
38. Zhou L, Maruyama I, Li Y, et al. Expression of integrin receptors in the human trabecular meshwork. Curr Eye Res. 1999;19:395–402.
39. Tervo K, Paallysaho T, Virtanen I, et al. Integrins in human anterior chamber angle. Graefes Arch Clin Exp Ophthalmol. 1995;233:291–295.
40. Grierson I, Lee WR. Changes in the monkey outflow apparatus at graded levels of intraocular pressure: a qualitative analysis by light microscopy and scanning electron microscopy. Exp Eye Res. 1974;19:21–33.
41. Kayes J. Pressure gradient changes on the trabecular meshwork of monkeys. Am J Ophthalmol. 1975;79:549–556.
42. Grierson I, Lee WR. Pressure effects on the distribution of extracellular materials in the rhesus monkey outflow apparatus. Albrecht Von Graefes Arch Klin Exp Ophthalmol. 1977;203:155–168.
43. Van Buskirk EM. Anatomic correlates of changing aqueous outflow facility in excised human eyes. Invest Ophthalmol Vis Sci. 1982;22:625–632.
44. Ingber DE. Mechanical signaling and the cellular response to extracellular matrix in angiogenesis and cardiovascular physiology. Circ Res. 2002;91:877–887.
45. Gills JP, Roberts BC, Epstein DL. Microtubule disruption leads to cellular contraction in human trabecular meshwork cells. Invest Ophthalmol Vis Sci. 1998;39:653–658.
46. Mertts M, Garfield S, Tanemoto K, et al. Identification of the region in the N-terminal domain responsible for the cytoplasmic localization of Myoc/Tigr and its association with microtubules. Lab Invest. 1999;79:1237–1245.
47. Weinreb RN, Ryder MI. In situ localization of cytoskeletal elements in the human trabecular meshwork and cornea. Invest Ophthalmol Vis Sci. 1990;31:1839–1847.
48. de Kater AW, Shahsafaei A, Epstein DL. Localization of smooth muscle and nonmuscle actin isoforms in the human aqueous outflow pathway. Invest Ophthalmol Vis Sci. 1992;33:424–429.
49. Ringvold A. Actin filaments in trabecular endothelial cells in eyes of the vervet monkey. (Cercopithecus aethiops). Acta Ophthalmol (Copenh). 1978;56:217–225.
50. Tamm ER, Siegner A, Baur A, et al. Transforming growth factor-beta 1 induces alpha-smooth muscle-actin expression in cultured human and monkey trabecular meshwork. Exp Eye Res. 1996;62:389–397.
51. Grierson I, Millar L, De Yong J, et al. Investigations of cytoskeletal elements in cultured bovine meshwork cells. Invest Ophthalmol Vis Sci. 1986;27:1318–1330.
52. Weinreb RN, Ryder MI, Polansky JR. The cytoskeleton of the cynomolgus monkey trabecular cell. II. Influence of cytoskeleton-active drugs. Invest Ophthalmol Vis Sci. 1986;27:1312–1317.
53. Ryder MI, Weinreb RN. The cytoskeleton of the cynomolgus monkey trabecular cell. I. General considerations. Invest Ophthalmol Vis Sci. 1986;27:1305–1311.
54. Ryder MI, Weinreb RN, Alvarado J, et al. The cytoskeleton of the cultured human trabecular cell. Characterization and drug responses. Invest Ophthalmol Vis Sci. 1988;29:251–260.
55. Iwamoto Y, Tamura M. Immunocytochemical study of intermediate filaments in cultured human trabecular cells. Invest Ophthalmol Vis Sci. 1988;29:244–250.
56. Tamura M, Iwamoto Y, Nakatsuka K, et al. Immunofluorescence studies of the cytoskeletal and contractile elements in cultured human trabecular cells. Jpn J Ophthalmol. 1989;33:95–102.
57. Grierson I, Rahi AH. Microfilaments in the cells of the human trabecular meshwork. Br J Ophthalmol. 1979;63:3–8.
58. Tomarev SI, Wistow G, Raymond V, et al. Gene expression profile of the human trabecular meshwork: NEIBank sequence tag analysis. Invest Ophthalmol Vis Sci. 2003;44:2588–2596.
59. Ellingsen BA, Grant WM. The relationship of pressure and aqueous outflow in enucleated human eyes. Invest Ophthalmol. 1971;10:430–437.
60. Moses RA. The effect of intraocular pressure on resistance to outflow. Surv Ophthalmol. 1977;22:88–100.
61. Moses RA. Circumferential flow in Schlemm’s canal. Am J Ophthalmol. 1979;88:585–591.
62. Moses RA, Grodzki WJ Jr, Etheridge EL, et al. Schlemm’s canal: the effect of intraocular pressure. Invest Ophthalmol Vis Sci. 1981;20:61–68.
63. Stegmann R, Pienaar A, Miller D. Viscocanalostomy for open-angle glaucoma in black African patients. J Cataract Refract Surg. 1999;25:316–322.
64. Smit BA, Johnstone MA. Effects of viscocanalostomy on the histology of Schlemm’s canal in primate eyes. Invest Ophthalmol Vis Sci. 2000;41:S578.
65. Smit BA, Johnstone MA. Effects of viscoelastic injection into Schlemm’s canal in primate and human eyes: potential relevance to viscocanalostomy. Ophthalmology. 2002;109:786–792.
66. Johnstone MA, Tanner D, Chau B. Endothelial tubular channels in Schlemm’s canal. Invest Ophthalmol Vis Sci. 1980;19:123.
67. Grierson I, Lee WR. Acid mucopolysaccharides in the outflow apparatus. Exp Eye Res. 1975;21:417–431.
68. Acott TS, Kingsley PD, Samples JR, et al. Human trabecular meshwork organ culture: morphology and glycosaminoglycan synthesis. Invest Ophthalmol Vis Sci. 1988;29:90–100.
69. Knepper PA, Goossens W, Hvizd M, et al. Glycosaminoglycans of the human trabecular meshwork in primary open-angle glaucoma. Invest Ophthalmol Vis Sci. 1996;37:1360–1367.
70. Lerner LE, Polansky JR, Howes EL, et al. Hyaluronan in the human trabecular meshwork. Invest Ophthalmol Vis Sci. 1997;38:1222–1228.
71. Knepper PA, McLone DG. Glycosaminoglycans and outflow pathways of the eye and brain. Pediatr Neurosci. 1985;12:240–251.
72. Alvarado JA, Yun AJ, Murphy CG. Juxtacanalicular tissue in primary open angle glaucoma and in nonglaucomatous normals. Arch Ophthalmol. 1986;104:1517–1528.
73. Murphy CG, Yun AJ, Newsome DA, et al. Localization of extracellular proteins of the human trabecular meshwork by indirect immunofluorescence. Am J Ophthalmol. 1987;104:33–43.
74. Svedbergh B. Protrusions of the inner wall of Schlemm’s canal. Am J Ophthalmol. 1976;82:875–882.
75. Johnstone MA, Tanner D, Chau B, et al. Concentration-dependent morphologic effects of cytochalasin B in the aqueous outflow system. Invest Ophthalmol Vis Sci. 1980;19:835–841.
76. Rudbeck O. Nova Excercit at 10 Anatomica, Exhibens Ductus Hepaticos Aquosos, & Vasa Glandularum Serosa, nunc primum inventa, 1st ed. Vol. 1. Uppsala: Eucharius Lauringerus; 1653. Contributions from the Karolinska Institute Library and Museum Collections–Volume III. Produced by Hagelin Antikvariat AB; Stockholm 1992.
77. Harvey W. Exercitatio Anatomica de Motu Cordis et Sanguinis in Animalibus; 1628. An English Translation with Annotations. Springfield: Charles C. Thomas; 1970.
78. Levick JR. Cardiovascular Physiology, 3rd ed. London; co-published New York: Arnold; co-published by Oxford University Press; 2000:416.
79. Welch K, Friedman V. The cerebrospinal fluid valves. Brain. 1960;83:454–469.
80. Van Buskirk EM. The canine eye: The vessels of aqueous drainage. Invest Ophthalmol. 1979;18:223–230.
81. Ascher KW. Physiologic importance of the visible elimination of intraocular fluid. Am J Ophthalmol. 1942;25:1174–1209.
82. De Vries S. De Zichtbare Afvoer Van Het Kamerwater, 1st ed. Amsterdam: Drukkerij Kinsbergen; 1947.
83. Goldmann H. Abfluss des Kammerwassers beim Menschen. Ophthalmologica. 1946;111:146152.
84. Goldmann H. Weitere Mitteilung über den Abfluss des Kammerwassers beim Menschen. Ophthalmologica. 1946;112:344–346.
85. Thomassen TL. On aqueous veins. Acta Ophth. 1947;25:369–378.
86. Ascher KW. Glaucoma and the aqueous veins. Am. J. Ophth. 1942;25:1309–1315.
87. Kleinert H. Der sichtbare Abfluss des Kammerwassers in den epiibulbären Venen. II. Mitteilung. Die pulsiereden Kammerwassergefässe. von Graefes ArchOphth. 1952;152:587–608.
88. Kleinert H. Der Sichtbare Abfluss des Kammerwassers in den epibulbären Venen. von Graefes ArchOphth. 1951;152:278–299.
89. Vries S. De zichtbare Afvoer von het Kamerwater. Amsterdam; Drukkerij Kinsbergen; 1947.
90. Trantas NG. La pression des vaisseaux retiniens sous la compressoin du sinus carotidien et la circulation oculaire. Bull Soc Opht France. 1950;63:64–71.
91. Gartner S. Blood vessels of the conjunctiva. Arch Ophthalmol. 1944;32:464–469.
92. Johnstone MA. The aqueous outflow system as a mechanical pump. International Glaucoma Review. 2003;5:14.
93. Johnstone MA. Glaucoma and the aqueous outflow channels. Transactions of the Pacific Coast OtoOphthalmological Society. 1979;60:153.
94. Johnstone MA. Stretching and compression of trabecular tissue. Inter’l Soc. for Eye Research Abstract, Fifth International Congress of Eye Research; 1980:90.
95. De Jong H. Modeling and simulation of genetic regulatory systems: a literature review. J Comput Biol. 2002;9:67–103.
96. Knowlton FP, Starling EH. The influence of variations in temperature and blood pressure on the performance of the isolated mammalian heart. Physiology. 1912;44:206–219.
97. Conrad WA. Pressure—flow relationships in collapsible tubes. IEEE Trans Biomed Eng. 1969;16:284–295.
98. Wiederholt M. Direct involvement of trabecular meshwork in the regulation of aqueous humor outflow. Curr Opin Ophthalmol. 1998;9:46–49.
99. Berne RM, Levy MN. Cardiovascular Physiology, 8th ed. St. Louis, MO; Mosby 2001.
100. Rodbard S. Vascular caliber. Cardiology. 1975;60:4–49.
101. Bradley JM, Kelley MJ, Zhu X, et al. Effects of mechanical stretching on trabecular matrix metalloproteinases. Invest Ophthalmol Vis Sci. 2001;42:1505–1513.
102. Johnson DH, Johnson M. How does nonpenetrating glaucoma surgery work? Aqueous outflow resistance and glaucoma surgery. J Glaucoma. 2001;10:55–67.
103. Johnson DH, Johnson M. Glaucoma surgery and aqueous outflow: how does nonpenetrating glaucoma surgery work? Arch Ophthalmol. 2002;120:67–70.
104. Bill A, Svedbergh BB. Scanning electron microscopic studies of the trabecular meshwork and the canal of Schlemm—an attempt to localize the main resistance to outflow of aqueous humor in man. Acta Ophthalmol. 1972;50:295–320.
105. Maepea O, Bill A. Pressures in the juxtacanalicular tissue and Schlemm’s canal in monkeys. Exp Eye Res. 1992;54:879–883.
106. Ethier CR. The inner wall of Schlemm’s canal. Exp Eye Res. 2002;74:161–172.
107. Alenghat FJ, Ingber DE. Mechanotransduction: all signals point to cytoskeleton, matrix, and integrins. Sci STKE 2002:PE6.
108. Pries AR, Secomb TW, Gaehtgens P. Design principles of vascular beds. Circ Res. 1995;77:1017–1023.
109. Pries AR, Secomb TW, Gaehtgens P. Structure and hemodynamics of microvascular networks: heterogeneity and correlations. Am J Physiol. 1995;269:H1713–H1722.
110. Murray CD. The physiological principle of minimum work. I. The vascular system and the cost of blood volume. Proc Natl Acad Sci. 1926;12:207–214.
111. Taber LA. An optimization principle for vascular radius including the effects of smooth muscle tone. Biophys J. 1998;74:109–114.
112. Curry FE, Michel CC. A fiber matrix model of capillary permeability. Microvasc Res. 1980;20:96–99.
113. Michel CC. Filtration coefficients and osmotic reflexion coefficients of the walls of single frog mesenteric capillaries. J Physiol. 1980;309:341–355.
114. Michel CC, Phillips ME. The effects of bovine serum albumin and a form of cationised ferritin upon the molecular selectivity of the walls of single frog capillaries. Microvasc Res. 1985;29:190–203.
115. Fu BM, Shen S. Structural mechanisms of acute VEGF effect of microvessel permeability. Am J Physiol Heart Circ Physiol. 2003;284:H2124–H2135.
116. Hu X, Adamson RH, Liu B, et al. Starling forces that oppose filtration after tissue oncotic pressure is increased. Am J Physiol Heart Circ Physiol. 2000;279:H1724–H1736.
117. Fraser WD, Baines AD. Application of a fiber-matrix model to transport in renal tubules. J Gen Physiol. 1989;94:863–879.
118. Epstein DL, Rohen JW. Morphology of the trabecular meshwork and inner-wall endothelium after cationized ferritin perfusion in the monkey eye. Invest Ophthalmol Vis Sci. 1991;32:160–171.
119. Ethier CR, Chan DW. Cationic ferritin changes outflow facility in human eyes whereas anionic ferritin does not. Invest Ophthalmol Vis Sci. 2001;42:1795–1802.
120. Davies PF, Barbee KA, Volin MV, et al. Spatial relationships in early signaling events of flow-mediated endothelial mechanotransduction. Annu Rev Physiol. 1997;59:527–549.
121. Ingber D. The architecture of life. Sci Am. 1998;278:48–57.
122. Stamer WD, Roberts BC, Epstein DL. Hydraulic pressure stimulates adenosine 3′,5′-cyclic monophosphate accumulation in endothelial cells from Schlemm’s canal. Invest Ophthalmol Vis Sci. 1999;40:1983–1988.
123. Tumminia SJ, Mitton KP, Arora J, et al. Mechanical stretch alters the actin cytoskeletal network and signal transduction in human trabecular meshwork cells [see comments]. Invest Ophthalmol Vis Sci. 1998;39:1361–1371.
124. Gonzalez P, Epstein DL, Borras T. Genes upregulated in the human trabecular meshwork in response to elevated intraocular pressure. Invest Ophthalmol Vis Sci. 2000;41:352–361.
125. Tamm ER, Russell P, Epstein DL, et al. Modulation of myocilin/TIGR expression in human trabecular meshwork. Invest Ophthalmol Vis Sci. 1999;40:2577–2582.
126. Mitton KP, Tumminia SJ, Arora J, et al. Transient loss of alphaB-crystallin: an early cellular response to mechanical stretch. Biochem Biophys Res Commun. 1997;235:69–73.
127. Sato Y, Matsuo T, Ohtsuki H. A novel gene (oculomedin) induced by mechanical stretching in human trabecular cells of the eye. Biochem Biophys Res Commun. 1999;259:349–351.
128. Schnittler HJ. Structural and functional aspects of intercellular junctions in vascular endothelium. Basic Res Cardiol. 1998;93(Suppl 3):30–39.
129. DeMaio L, Chang YS, Gardner TW, et al. Shear stress regulates occludin content and phosphorylation. Am J Physiol Heart Circ Physiol. 2001;281:H105–H113.
130. Stamer WD, Snyder RW, Smith BL, et al. Localization of aquaporin CHIP in the human eye: implications in the pathogenesis of glaucoma and other disorders of ocular fluid balance. Invest Ophthalmol Vis Sci. 1994;35:3867–3872.
131. Stamer WD, Seftor RE, Snyder RW, et al. Cultured human trabecular meshwork cells express aquaporin-1 water channels. Curr Eye Res. 1995;14:1095–1100.
132. Ishida T, Takahashi M, Corson MA, et al. Fluid shear stress-mediated signal transduction: how do endothelial cells transduce mechanical force into biological responses? Ann N Y Acad Sci. 1997;811:12–23.
133. Li S, Chen BP, Azuma N, et al. Distinct roles for the small GTPases Cdc42 and Rho in endothelial responses to shear stress. J Clin Invest. 1999;103:1141–1150.
134. Tzima E, del Pozo MA, Shattil SJ, et al. Activation of integrins in endothelial cells by fluid shear stress mediates Rho-dependent cytoskeletal alignment. EMBO J. 2001;20:4639–4647.
135. Shyy JY, Chien S. Role of integrins in endothelial mechanosensing of shear stress. Circ Res. 2002;91:769–775.
136. Rao PV, Deng PF, Kumar J, et al. Modulation of aqueous humor outflow facility by the Rho kinase-specific inhibitor Y-27632. Invest Ophthalmol Vis Sci. 2001;42:1029–1037.
137. Li YS, Shyy JY, Li S, et al. The Ras-JNK pathway is involved in shear-induced gene expression. Mol Cell Biol. 1996;16:5947–5954.
138. Girard PR, Nerem RM. Endothelial cell signaling and cytoskeletal changes in response to shear stress. Front Med Biol Eng. 1993;5:31–36.
139. Traub O, Berk BC. Laminar shear stress: mechanisms by which endothelial cells transduce an atheroprotective force. Arterioscler Thromb Vasc Biol. 1998;18:677–685.
140. Berk BC, Corson MA, Peterson TE, et al. Protein kinases as mediators of fluid shear stress stimulated signal transduction in endothelial cells: a hypothesis for calcium-dependent and calcium-independent events activated by flow. J Biomech. 1995;28:1439–1450.
141. Ishida T, Peterson TE, Kovach NL, et al. MAP kinase activation by flow in endothelial cells. Role of beta 1 integrins and tyrosine kinases. Circ Res. 1996;79:310–316.
142. Bhullar IS, Li YS, Miao H, et al. Fluid shear stress activation of IkappaB kinase is integrin-dependent. J Biol Chem. 1998;273:30544–30549.
143. Nagel T, Resnick N, Dewey CF Jr, et al. Vascular endothelial cells respond to spatial gradients in fluid shear stress by enhanced activation of transcription factors. Arterioscler Thromb Vasc Biol. 1999;19:1825–1834.
144. Chien S, Li S, Shyy YJ. Effects of mechanical forces on signal transduction and gene expression in endothelial cells. Hypertension. 1998;31:162–169.
145. Du W, Mills I, Sumpio BE. Cyclic strain causes heterogeneous induction of transcription factors, AP-1, CRE binding protein and NF-kB, in endothelial cells: species and vascular bed diversity. J Biomech. 1995;28:1485–1491.
146. Thoumine O, Nerem RM, Girard PR. Oscillatory shear stress and hydrostatic pressure modulate cell-matrix attachment proteins in cultured endothelial cells. In Vitro Cell Dev Biol Anim. 1995;31:45–54.
147. Simon BC, Noll B, Maisch B. Endothelial dysfunction—assessment of current status and approaches to therapy. Herz. 1999;24:62–71.
148. Gloe T, Sohn HY, Meininger GA, et al. Shear stress-induced release of basic fibroblast growth factor from endothelial cells is mediated by matrix interaction via integrin alpha(v)beta3. J Biol Chem. 2002;277:23453–23458.
149. Lehoux S, Trone F, Tedgui A. Mechanisms of blood flow-induced vascular enlargement. Biorheology. 2002;39:319–324.
150. Yamaguchi S, Yamaguchi M, Yatsuyanagi E, et al. Cyclic strain stimulates early growth response gene product 1-mediated expression of membrane type 1 matrix metalloproteinase in endothelium. Lab Invest. 2002;82:949–956.
151. Sho E, Sho M, Singh TM, et al. Arterial enlargement in response to high flow requires early expression of matrix metalloproteinases to degrade extracellular matrix. Exp Mol Pathol. 2002;73:142–153.
152. Bongrazio M, Baumann C, Zakrzewicz A, et al. Evidence for modulation of genes involved in vascular adaptation by prolonged exposure of endothelial cells to shear stress. Cardiovasc Res. 2000;47:384–393.
153. Magid R, Murphy TJ, Galis ZS. Expression of matrix metalloproteinase-9 in endothelial cells is differentially regulated by shear stress. Role of c-Myc. J Biol Chem. 2003;278:32994–32999.
154. Arisaka T, Mitsumata M, Kawasumi M, et al. Effects of shear stress on glycosaminoglycan synthesis in vascular endothelial cells. Ann N Y Acad Sci. 1995;748:543–554.
155. Suarez J, Rubio R. Regulation of glycolytic flux by coronary flow in guinea pig heart. Role of vascular endothelial cell glycocalyx. Am J Physiol. 1991;261:H1994–H2000.
156. Damiano ER. The effect of the endothelial-cell glycocalyx on the motion of red blood cells through capillaries. Microvasc Res. 1998;55:77–91.
157. Turner MR, Clough G, Michel CC. The effects of cationised ferritin and native ferritin upon the filtration coefficient of single frog capillaries. Evidence that proteins in the endothelial cell coat influence permeability. Microvasc Res. 1983;25:205–222.
158. Chiu JJ, Lee PL, Lee CI, et al. Shear stress attenuates tumor necrosis factor-alpha-induced monocyte chemotactic protein-1 expressions in endothelial cells. Chin J Physiol. 2002;45:169–176.
159. Chappell DC, Varner SE, Nerem RM, et al. Oscillatory shear stress stimulates adhesion molecule expression in cultured human endothelium. Circ Res. 1998;82:532–539.
160. Kraiss LW, Alto NM, Dixon DA, et al. Fluid flow regulates E-selectin protein levels in human endothelial cells by inhibiting translation. J Vasc Surg. 2003;37:161–168.
161. Ando J, Kamiya A. Flow-dependent regulation of gene expression in vascular endothelial cells. Jpn Heart J. 1996;37:19–32.
162. Korenaga R, Ando J, Kosaki K, et al. Negative transcriptional regulation of the VCAM-1 gene by fluid shear stress in murine endothelial cells. Am J Physiol. 1997;273:C1506–C1515.
163. Ando J, Tsuboi H, Korenaga R, et al. Shear stress inhibits adhesion of cultured mouse endothelial cells to lymphocytes by downregulating VCAM-1 expression. Am J Physiol. 1994;267:C679–C687.
164. Ando J, Tsuboi H, Korenaga R, et al. Down-regulation of vascular adhesion molecule-1 by fluid shear stress in cultured mouse endothelial cells. Ann N Y Acad Sci. 1995;748:148–156.
165. Shyy JY, Li YS, Lin MC, et al. Multiple cis-elements mediate shear stress-induced gene expression. J Biomech. 1995;28:1451–1457.
166. Jalali S, Li YS, Sotoudeh M, et al. Shear stress activates p60src-Ras-MAPK signaling pathways in vascular endothelial cells. Arterioscler Thromb Vasc Biol. 1998;18:227–234.
167. Ji JY, Jing H, Diamond SL. Shear stress causes nuclear localization of endothelial glucocorticoid receptor and expression from the GRE promoter. Circ Res. 2003;92:279–285.
168. Weinreb RN, Bloom E, Baxter JD, et al. Detection of glucocorticoid receptors in cultured human trabecular cells. Invest Ophthalmol Vis Sci. 1981;21:403–407.
169. Polansky JR, Kurtz RM, Alvarado JA, et al. Eicosanoid production and glucocorticoid regulatory mechanisms in cultured human trabecular meshwork cells. Prog Clin Biol Res. 1989;312:113–138.
170. Underwood JL, Murphy CG, Chen J, et al. Glucocorticoids regulate transendothelial fluid flow resistance and formation of intercellular junctions. Am J Physiol. 1999;277:C330–C342.
171. Clark AF, Wilson K, McCartney MD, et al. Glucocorticoid-induced formation of cross-linked actin networks in cultured human trabecular meshwork cells. Invest Ophthalmol Vis Sci. 1994;35:281–294.
172. Polansky JR, Fauss DJ, Zimmerman CC. Regulation of TIGR/MYOC gene expression in human trabecular meshwork cells. Eye. 2000;14:503–514.

glaucoma; Schlemm’s canal; trabecular meshwork; endothelium; valve; pump; shear stress

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