Prelude to corneal tissue engineering - gaining control of collagen organization

Abstract

By most standard engineering practice principles, it is premature to credibly discuss the "engineering" of a human cornea. A professional design engineer would assert that we still do not know what a cornea is (and correctly so), therefore we cannot possibly build one. The proof resides in the fact that there are no clinically viable corneas based on classical tissue engineering methods available. This is possibly because tissue engineering in the classical sense (seeding a degradable scaffolding with a population synthetically active cells) does not produce conditions which support the generation of organized tissue. Alternative approaches to the problem are in their infancy and include the methods which attempt to recapitulate development or to produce corneal stromal analogs de novo which require minimal remodeling. Nonetheless, tissue engineering efforts, which have been focused on producing the fundamental functional component of a cornea (organized alternating arrays of collagen or "lamellae"), may have already provided valuable new insights and tools relevant to development, growth, remodeling and pathologies associated with connective tissue in general. This is because engineers ask a fundamentally different question (How can that be done?) than do biological scientists (How is that done?). The difference in inquiry has prompted us to closely examine (and to mimic) development as well as investigate collagen physicochemical behavior so that we may exert control over organization both in cell culture (in vitro) and on the benchtop (de novo). Our initial results indicate that reproducing corneal stroma-like local and long-range organization of collagen may be simpler than we anticipated while controlling spacing and fibril morphology remains difficult, but perhaps not impossible in the (reasonably) near term.

Figure 1

The cornea (approximately 500 microns thick in humans) is a layered structure comprising the epithelium (E), Bowman’s layer (Bw), stroma (St), Descemet’s membrane (De), and the endothelium (En). (with permission from Klyce and Beuerman, 1997)

Figure 2

A. Proposed model of fibril orientation in the cornea based on X-Ray synchrotron data. B. Variation in corneal tensile strength as a function of direction. Though the cornea is typically considered a simple nematic stack of lamellae comprising aligned collagen which alternate in orientation by 90 degrees, recent investigations demonstrate an array of fibrils which run circumferentially around the periphery. Preferred fibril orientation and aligned fibril concentration is reflected in the tensile strength or modulus. The figure shows the tensile modulus found in test strips excised and loaded in the direction of the arrows. (A) with permission from (Meek and Boote, 2004) and B with permission from Ruberti et al. (Ruberti et al., 2007).

Figure 3

Bowman’s layer in the dogfish. There is no discernible “organization” to the the fibrils in this layer yet this portion of the cornea is more transparent than the underlying more “organized” stroma proper. With permission from (Goldman and Benedek, 1967)

Figure 4

Transmission Electron Micrograph of collagen fibril arrangement in cornea. Corneal collagen fibrils have a highly monodisperse diameter distribution and are arranged in oriented arrays within lamellae which are approximately 1–2 microns thick. (Section is normal to tangent plane). Image courtesy of Dr. Haiyan Gong.

Figure 5

Light micrograph of stratified corneal epithelium grown using air-lift technique and in organotypic, co-culture with endothelium and keratocytes. Bar is 50 microns. with permission from Zieske et al 1994 (Zieske et al., 1994)

Figure 6

Developing chick cornea. Epithelium, primary stroma and endothelium which is continuous with the lens are depicted. The acellular primary stroma is 10 microns thick at this stage of development (stage 27; day 5). With permission from Trelstad and Coulombre (Trelstad and Coulombre, 1971)

Figure 7

Primary stroma in developing chick cornea. A) TEM of primary stromal collagen showing orthogonal and discontinuous fibrils. B) QFDE image of orthogonal primary stromal collagen. With permission from Trelstad and Coulombre (Trelstad and Coulombre, 1971) and Hirsch et al. (Hirsch et al., 1999).

Figure 8

Critical stages in corneal development. Images depicts corneal collagen organization in transverse section (large rectangles) and en face section (small circles). At stage 27 (day 5) the epithelium has generated a primary stroma that strictly orthogonal and approximately 10 microns thick. Following a rapid expansion at the end of day 5, secondary stroma is produced posteriorly by invading mesenchymal cells. Note the shift in the register of the orthogonal collagen in the anterior uninvaded primary stroma. At the end of this stage (stage 33 (day 7)) the nascent stroma is 170 microns thick with the anterior 10 microns uninvaded. By stage 35 (day 9) the stroma is 190 microns thick with the posterior 180 microns invaded my mesenchymal cells. The anteriormost 40 microns demonstrate an angular shift between orthogonal layers of collagen, while the posterior 150 microns are in register. With permission from Trelstad and Coulombre (Trelstad and Coulombre, 1971)

Figure 9

Light micrographs of developing (A) rabbit and (B) human cornea at the initiation of mesenchymal invasion. Note the similarity in the geometry. Both rabbit and human do not have discernible endothelium at the time of mesenchymal invasion, while some primates and avians do. With permission from (Cintron et al., 1983; Sevel and Isaacs, 1988).

Figure 10

Light micrographic sequence of the developing rabbit stroma from Day 14 through birth to post-natal day 21. The depicted progression is generally representative of mammalian stromal development (sans Bowman’s). The sequence demonstrates prospective stroma formation (day 14), mesenchymal cell invasion (day 15), strom0l synthesis and expansion (days 16 to 23), increase in posterior stromal organization cell flattening (days 16–21), stromal compaction/deturgescence (days 23-birth), Anterior stromal cell flattening (days 26-birth). With permission from Cintron et al (Cintron et al., 1983).

Figure 11

Transmission electron micrograph of 21 day rabbit fetal cornea demonstrating the density of cells in the developing tissue. With permission from (Cintron et al., 1983)

Figure 12

(A) SEM of an adult human corneal keratocyte (K) showing the complex dendritc morphology of the cell body and processes. (B) Light micrographs of keratocyte network in the feline. Keratocytes are connected by broad cellular processes (open arrows) extending from main cell body which contains nucleus (arrows) With permission from Muller et al 1995 (Muller et al., 1995) (A) and Jester et al 1994 (Jester et al., 1994) (B)

Figure 13

Phase contrast optical micrographs of dendritic keratocytes (A) versus more fibroblastic (B-repair type fibroblasts) cells in culture. With permission from (Musselmann et al., 2005).

Figure 14

(A) Confocal Scanning Laser Microscope image of myofibroblast stress fibers aligned adjacent to the wound margin (dark area) in a linear incision wound in a rabbit cornea. (B) Schematic of the possible “shoestring” wound contracture model in the rabbit. In vivo confocal 3-D imaging revealed that wound contracture in corneal wounds my occur in a manner similar to pulling closed the top of a shoe with shoestings. With permission from Petroll et al (Petroll et al., 1993)).

Figure 15

Phase contrast image of a multilayered primary human corneal fibroblast culture after one week in culture. Modified from Guo et al (Guo et al., 2007) with permission. Bar is 20 microns

Figure 16

Confocal reflectance microscopy of human corneal fibroblasts in a reconstituted type I collagen gel that has been mechanically loaded by fibroblast contractile force. The cells produce organization in bundles of collagen fibrils between their “ends” when they are induced to align by the mechanical constraints on the matrix. With permission from Karmichos et al (Karamichos et al., 2007)

Figure 17

Photomicrograph of Giemsa-stained preparation illustrating the alignment of fibroblasts with orthogonal layers of shear-aligned collagen. Collagen was aligned during polymerization by draining collagen monomer solution on inclined plate. Cells were introduced onto first layer of aligned collagen. A second layer of aligned collagen was produced orthogonal to the first followed by plating a second population of cells. Note resulting orthogonal arrangement of cells. With permission from Elsdale and Bard (Elsdale and Bard, 1972).

Figure 18

Corneal construct of Griffith et al. (A) Demonstration of construct clarity compared to post-mortem human cornea. Scale bar 10 mm (B) Cultured eye bank cornea (C) cultured corneal equivalent. Scale bar 100 microns. Epithelium – EP; Bowman’s membrane – Bm; Stroma – S; Descemet’s – Dm; Endothelium – En. With permission from Griffith et al., 1999

Figure 19

Photographic comparison of the TERP5-collagen composite of Li et al.,2003 (A) and a reconstituted type I collagen gel (B). With permission from Li et al. (Li et al., 2003))

Figure 20

H&E stained stromal analog of Orwin and Hubel. (A) By day 10 of culture the cells have migrated through the stromal sponge. (B) By day 22 of culture cells have elongated along the collagen fibers. With permission from Orwin and Hubel (Orwin and Hubel, 2000).

Figure 21

SEM micrographs of primary human corneal cells seeded onto dense collagen films. A) Rounded cells after one day on the film. B) Confluent cells after one week on the film. Asterisk indicates film top surface. With permission From Crabb et al. (Crabb et al., 2006)

Figure 22

SEM micrographs of ECM synthesized by primary human corneal cells seeded onto dense collagen films after 5 weeks in culture. Arrows indicate smooth “fibrils” which are approximately 100 nm in diameter while the bracket indicates smaller diameter fibrils (47 nm) with a relatively large dispersity (± 17 nm). With permission from Crabb et al (Crabb et al., 2006)

Figure 23

Scanning electron micrographs of a reconstituted type I collagen gel (A), cell-assembled collagen in native human corneal stroma (B), cell-assembled collagen in human hip skin (C). In the native cornea, arrays of highly-organized fibrils split and pass above and below lamellae with different orientations. In skin, the collagen is more loosely organized and does not have to be transparent. It appears simpler to transform the structure from A to C than it is to transform the structure from A to B. Image in (B) with permission from Radner et al., 1998 (Radner et al., 1998). Image in (C) with permission from Fligiel et al (Fligiel et al., 2003).

Figure 24

Standard TEM micrographs of pHCSCs stimulated to produce collagen. (A) low magnification image of cells and matrix in the constructs. The collagen fibrils clearly alternate in direction in neighboring lamellae. (B) Higher magnification showing at least 9 direction changes in collagen lamellar orientation. (C) Higher magnification image showing detailed fibril organization. Bars (A,B) 2 μm and (C) 1μm. With permission from Guo et al. (Guo et al., 2007)

Figure 25

Contrast-Enhanced Optical DIC image of cell-assembled collagenous ECM. Location of image section is approximately 8 microns above and parallel to the transwell membrane. Image texture is suggestive of alternating arrays of fibrillar structures which persist over large length scales (order 10s of microns). The construct in this image is 4 weeks old. Bar is 10 microns. Image courtesy of Nima Saeidi. (unpublished data)

Figure 26

Transmission electron micrographs of 4 week old stromal construct produced by untransformed human corneal fibroblasts in culture. Note synthetically active stromal cells with prominent RER and orthogonal arrangement of fibrils in lamellae. The architecture of the construct organization is very similar to that found in the developing mammalian corneal stroma. Arrows indicate prominent dilated RER. Bar is 2 microns. With permission from Guo et al. (Guo et al., 2007)

Figure 27

Immunofluorescence and TEM micrographs of HCSSCs in pellet culture. (A) Fluorescence image of connexin and cadherin staining in pellet culture showing prominent staining for these junctional proteins in the periphery. (B) Electron Micrograph demonstrating organized arrangement of fibrils in the area with the increased staining for junctional proteins. With permission from Du et al. (Du et al., 2007)

Figure 28

DIC optical micrograph. Collagen fibrils assembled on clean borosilicate glass in the shear chamber. The fibrils were assembled in the presence of a 18 sec⁻¹ shear rate (arrow indicates flow direction). The kinetics of the assembly process suggest that collagen begins aggregating very rapidly on the glass surface (within 2 minutes) and that the alignment of the fibrils quickly becomes compromised by the formation of fibillar loops (inset in figure). (Bar is 10 microns). Image courtesy of Nima Saeidi (unpublished data – manuscript in preparation)

Figure 29

Quick Freeze Deep Etch image of extracted collagen, aligned and re-assembled in shear chamber. Morphology of the fibrils does not match native D-periodic banding. Individual monomers can be observed interacting with both the fibril and the substrate surface. This multiple interaction is likely the cause of the disrupted structure. Bar is 50 nm. Photograph courtesy of Nima Saeidi (unpublished data – manuscript in preparation)

Figure 30

Aligned collagen layer on glass substrate produced by introducing cold collagen solution onto warm spinning disk. Note the degree of alignment is excellent. However, even during spincoating, there is upstream polymerization instability which causes fibrils to turn, disrupting the alignment (highlighted circle). Black arrows indicate flow direction. Spin rate 1000 rpm; collagen flow rate 0.5 ml/min (3.0 mg/ml Purecol); 12 minute run. Bar is 10 microns. With permission from (Ruberti et al., 2007) Photo courtesy of Nima Saeidi (unpublished data – manuscript in preparation)

Figure 31

(A) Offset disk concept where collagen is deposited onto glass substrate offset in spinning disk. The collagen flows with a slight radial spread over substrate surface. Substrate may be rotated between coating runs to produce layered aligned structures (above). (B) SEM of collagen orthogonal layers produced in successive runs. Flow rate 0.25 ml/min, 1000 rpm, 15 min per run. Modified from Braithwaite and Ruberti US Pat 7,048,963

Figure 32

(A) Phase contrast optical microscopy of human corneal fibroblasts cultured on shear-aligned collagen from spin coater. Within six hours, fibroblasts had aligned in the direction of the collagen on the glass (center region). Fibroblasts on glass which was scraped clean of shear-aligned collagen were arranged randomly. (B) Confocal image of human corneal fibroblasts grown on denuded rabbit stroma after 24 hours in culture. Fibroblasts align (most likely with the fibril orientation in the exposed stroma). Images courtesy of Xiaoqing Guo.

Figure 33

Model of Birk and Trelstad (Birk and Trelstad, 1984) in which collagen fibrils are produced and aligned in compartments or surface crypts. In this model the cells are responsible for both the synthesis and organization of each fibril at the local level. Note that the same cell secretes collagen fibrillar arrays at right angles to one another. Thus the cell cannot move while depositing the fibils (and still maintain orthogonal fibril orientation). 1, 2, 3 types of surface invaginations producing collagen forming extracellular spaces. SV - secretory vesicle, With permission from (Birk and Trelstad, 1984)

Figure 34

(A) Model of fibripositor in cell. Procollagen is released from the secretory vesicle into the surface crypt where monomer orientation is controlled. One or both of the propeptides are cleaved to allow fibrillogenesis and the resulting collagen fibrils are “vectorially discharged” into the ECM. (B) Single reactor (fibripositor analog) in collagen nanoloom. Atelo, acid soluble collagen monomers (2.8 mg/ml) are kept cold (4°C) and at low pH on one side of polycarbonate track-etched membrane populated with 80 nm pores. The collagen is driven down the pore with a pressure gradient where conditions at the outlet promote fibrillogenesis (high pH, 37°C, high NaCl concentration). Fibrils are bound to a substrate which moves past the membrane, drawing assembling collagen from the pore. Image courtesy Northeastern University Capstone Design Course.

Figure 35

(A) Collagen Nanoloom was constructed in 2004 and comprises a 3-D nanomotion gantry, a peltier cooled collagen printer head (highlighted circle) and a heated copper substrate on which to print. At the heart of the design is a biomimetic array of fibripositors which are 80 nm in diameter and 6 microns long and contained in a polycarbonate track-etched membrane. (B) DIC optical image of a single broken strip of aligned “collagen” found after an attempted printer run. Three years of effort on the device have raised some very sophisticated engineering problems. Nonetheless it will likely provide some level of control over collagen organization when optimized. Bar is 20 microns. Images courtesy Kathryn Portale (unpublished data-manuscript in preparation).

Figure 36

Collagen fibrils formed by neutralizing concentrated collagen monomer solutions. Patterns of the resulting fibrils mimic patterns found in ligament (a) and in compact bone (b). Bars are 1 micron (a) and 500 nm (b). With permission from (Giraud-Guille et al., 2003)

Figure 37

Transmission Electron Micrographs of cell-synthesized organized collagen. (A) Standard TEM of embryonic chicken stroma at developmental day 14. Collagen fibrils appear to be “extruded” from cell. Note the large electron lucent areas. (B) standard TEM of pHCSC in scaffold-free development model at 11 weeks. Note discontinuous collagen and “white” space. (C) QFDE of 11 week pHCSC in development model. Aligned fibrils are present (lower right). Note that there is no “white” space detected by QFDE which does not depend on stains. Image in (A) with permssion from (Young et al., 2007). Images in B and C courtesy Drs. Ruberti and Zieske (manuscript under review at Developmental Dynamics)

Figure 38

(A) Schematic of the general approach to concentrate and confine collagen monomers. Step 1 – place low concentration collagen between two coverslips inside dialysis tube and immersed in osmicant (PEG in this case). Step 2 - Allow dialysis to continue until desired concentration is achieved. Step 3 – induce fibrillogenesis of the concentrated monomer solution. (B) DIC optical micrograph of the results of similar experiment. Concentration and confinement of monomers between two featureless coverslips produced an organized array of collagen fibrils. Image in (B) courtesy of Nima Saeidi (unpublished data – manuscript in preparation)

Figure 39

DIC optical image of collagen fibrillar arrays. The method of production of this collagen construct is similar to that of Figure 38. Low concentration collagen monomer solution was dialyzed against PEG and placed between two coverslips. The resulting constructs contained multiple layers of organized collagen alternate in direction. Image courtesy of Nima Saeidi. Bar is 20 microns (unpublished data – manuscript in preparation)

Figure 40

TEM of collagen fibrils produced from liquid crystalline type I atelo-collagen monomers. The fibrils are small (about 20 nm) and well-aligned. Banding is not entirely clear, but can be seen in some areas. Bar 100 nm. Image courtesy of Nima Saeidi (unpublished data – manuscript in preparation)