|Molecular Vision 2000;
Received 11 January 2000 | Accepted 19 April 2000 | Published 1 May 2000
A possible role for a-crystallins in lens epithelial cell differentiation
Daniel L. Boyle, Larry Takemoto
Division of Biology, Kansas State University, Manhattan, KS
Correspondence to: Daniel L. Boyle, Ph.D., Division of Biology, Ackert Hall, Kansas State University, Manhattan, KS, 66506; Phone: (785) 532-0134; FAX: (785) 532-6799; email: email@example.com
Purpose: To test the hypothesis that crystallin proteins may actively participate in the differentiation of lens epithelial cells into fiber cells.
Methods: Primary epithelial cells from adult bovine lenses were cultured at 37 °C until reaching 95-100% confluency in approximately 4-7 days. Using osmotic lysis of pinocytic vesicles, cells were then loaded with proteins labeled or unlabeled with the fluorescent marker Texas Red (TR). Fetal bovine proteins loaded into cells were lens water soluble fractions, HPLC-purified a-, b-, and g-crystallin fractions, or bovine serum albumin (BSA). Cultures were then monitored for morphological changes over a 7 day period.
Results: Both TR-labeled and unlabeled water-soluble and a-crystallin fractions from bovine lenses resulted in morphological changes to epithelial cells during the first two h postloading. These changes included aggregation of epithelial cells into raised multilayered cell masses, as well as several cells losing attachment to the dish. The initial changes were subsequently followed by elongation of cells within the mass and an increase in size of the mass, so that by 4 days postloading the multilayered, multicellular structures could be visualized with the unaided eye. Differentiation was confirmed within these structures by expression of MIP 26, b- and g-crystallin. These changes did not occur in cultures containing cells originally loaded with b- or g-crystallin fractions, or with cells loaded with BSA.
Conclusions: The results support the hypothesis that a-crystallins may actively participate in the differentiation of lens epithelial cells into fiber cells.
The mammalian lens is an avascular tissue, devoid of innervation, and located posterior to the iris in the eye. This tissue is enclosed by a collagenous capsule that is approximately three times thicker anteriorly as compared to posteriorly . A single layer of cuboidal epithelial cells which are nucleated and contain few organelles, line the anterior inner surface of the capsule. These epithelial cells are arranged into four distinctive zones . Going from anterior central to equatorial regions, these zones consist of a central cuboidal epithelial zone, a zone around the central zone containing relatively smaller epithelial cells, a pre-elongation or germative zone of epithelial cells, and an elongation or transitional zone where epithelial cells elongate and terminally differentiate into fiber cells [2,3]. The process of cell differentiation continues throughout life of the organism, resulting in fiber cells continually being formed at the elongation zone and displaced toward the center of the lens mass.
The molecular and cellular mechanisms responsible for this process are only beginning to be determined. What is known is that the transition from an epithelial cell to a fiber cell is characterized by changes in the expression of specific proteins. The a-crystallins are the most abundant protein in fiber cells [1,2]. In central epithelial cells, aA-crystallin and aB-crystallins are expressed at a molar ratio of 1:3, while in the elongation zone the molar ratio of aA-crystallin to aB-crystallin is 3:1 [1,2]. b- and g-crystallins as well as MIP 26 are not expressed by epithelial cells, but are found in great abundance in fiber cells [2-9]. To date, all these proteins have been used as markers for differentiated lens cells.
Some progress has also been made in determining the underlying molecular and cellular processes of lens epithelial cell differentiation, using anterior lens capsule explants and epithelial cell cultures. Adding various growth factors [3,10-22], or retinal and vitreal extracts [23-32] to culture media results in lens epithelial cells forming multicellular aggregates known as lens lentoid bodies. Cells present in the lentoid body have been characterized as differentiated fiber cells based on the expression of crystallin proteins and MIP 26 [4,10-22]. The initial change in protein content of an epithelial cell differentiating into fiber cells has been characterized by an increased expression of a-crystallins, followed by b-crystallins, then g-crystallins [4,12,14,17]. Based upon this observation, it is possible that expression of these crystallins may not simply be a consequence of differentiation, but instead may also play an important, if not necessary role, in the differentiation process itself. This hypothesis is not new and has been presented in earlier studies . In the present study we have used osmotic lysis of pinocytic vesicles to load lens epithelial cells with fluorescently labeled lens crystallins to test this hypothesis. The results demonstrate that the a-crystallins, but not b- or g-crystallins, cause dramatic changes in cell morphology characteristic of differentiating epithelial cells, suggesting that the a-crystallins may play an active role in the conversion of lens epithelial cells to fiber cells in the intact lens.
Bovine eyes were obtained from a local slaughterhouse and the Department of Animal Sciences and Industry, Kansas State University. Whole eyes were dipped in phosphate buffered saline pH 7.4 containing 10% (v/v) betadine prior to aseptic removal and dissection of the lens. A 360° incision approximately two mm posterior to the ora serrata was made, making the ciliary body, suspensory ligaments and posterior lens accessible. The suspensory ligaments were cut and the lens, with its intact capsule, was removed and placed with anterior surface down in a 40 x 10 mm tissue culture dish (Midwest Scientific, Valley Park, MO) containing two ml of 0.25% (w/v) trypsin-EDTA solution (Sigma, St. Louis, MO) for two min at room temperature (RT). The lens was then added to a 1.5-coverslip clear Delta-TC3 culture dish (Bioptechs, Butler, PA) containing one ml of Trypsin-EDTA. A 360° incision through the capsule at the lens equator was made, followed by removal of the anterior capsule with epithelial cells attached. The lens cortex, nucleus and posterior capsule were removed from the dish. The anterior capsule was then incubated for approximately two min with rotation at RT. One ml of Minimum Essential Medium (MEM) Eagles (Sigma) with non-essential amino acids and Earle's salts, without phenol red, sodium bicarbonate or antibiotics supplemented with L-glutamine (Sigma) and 10% bovine calf serum (Summit Biotechnology, Ft. Collins, CO) was added to the culture dish. Cultures were incubated at 37 °C with 5.0% CO2. After an initial 24-h period, the medium in each culture dish was changed three times per week until cells reached 95-100% confluency.
Isolation of crystallins
Fetal bovine eyes were obtained from Antech, Inc., Tyler, TX. Lenses were dissected from eyes and stored at -75 °C until used. Lens water soluble fraction (WSF) was prepared as previously described . a-, bhigh-, blow-, and g-crystallins were resolved from the WSF using a TSK-3000 column . The bhigh-, blow-, and g-crystallins were collected and used in experiments. The a-crystallin fraction was further resolved on a Biosep-SEC-S4000 gel permeation column into the high molecular weight aggregate fraction and the a-crystallin fraction . This a-crystallin fraction was collected and used in experiments. Collected fractions were dialyzed in Spectra/por (Spectrum Medical Industries, Inc., Los Angeles, CA) dialysis tubing with a 12-14,000 molecular weight cutoff (MWCO) overnight at 4 °C against distilled water. Protein concentrations for each collected fraction were determined in triplicate using the Bio-Rad protein assay based on the Bradford method (Bio-Rad, Hercules, CA), using BSA as the standard. Fractions were lyophilized and stored at -75 °C until used.
aA and aB subunits were purified from the afraction from the TSK-3000 column, in a manner similar to Perry and Abraham  using a C18 reverse phase column (250 x 4.6 mm, 300 angstrom pore size, Rainin Instrument Co., Woburn, MA) with a flow rate of 1 ml/min, and a linear gradient of 30-80% (v/v) acetonitrile in 0.1% (v/v) trifluoroacetic acid, over 60 min. aA and aB peaks were lyophilized, then dissolved in 8 M urea containing 0.1 M Tris, pH 7.4, followed by dialysis against a solution of 0.1 M Tris, pH 7.4.
Labeling proteins with Texas Red
One mg of lyophilized protein or BSA (Sigma) was dissolved in one ml of 0.025 M sodium borate buffer pH 8.8. For each mg of protein dissolved, 10 ml of a stock solution of 10 mg of Texas Red sulfonyl chloride (Molecular Probes, Eugene, OR) dissolved in 100 ml of dimethylformamide was added. This solution was immediately light protected, as well as light protected in all subsequent steps, and allowed mixing at 4 °C for one h. Labeled protein was separated from free fluorophore by dialysis in 12-14,000 MWCO tubing at 4 °C against distilled water overnight. Both labeled and unlabeled proteins were concentrated to one mg of protein per 100 ml of volume using 10,000 MWCO Microcon centrifuge filter devices (Millipore Corporation, Bedford, MA). Label proteins were used immediately or aliquoted and stored at -75 °C until used.
Loading cells with protein
Proteins were introduced into epithelial cells by the method of osmotic lysis of pinosomes [37-39]. Cells were washed twice with 2 ml of Hank's balanced salt solution (HBSS, Sigma) followed by the addition of 1 ml of hypertonic solution containing MEM, 0.5 M sucrose, 10% (v/v) polyethylene glycol (Sigma) and 20 ml (10 mg per ml) of labeled or unlabeled protein for 10 min at 37 °C. The hypertonic solution was than removed and a hypotonic solution of 6 parts MEM to 4 parts sterile, distilled, deionized water was briefly added for approximately one min. The hypotonic solution was replaced by fresh MEM and cultures were followed by confocal microscopy over the next seven days.
Cultures were fixed for 15 min in a 1:1 ratio of MEM and 2% (v/v) paraformaldehyde, 0.2% (v/v) glutaraldehyde in phosphate buffered saline (PBS), pH 7.4, followed by a 2 h fixation in fixative alone. Cultures were then washed with PBS, free aldehydes were quenched and nonspecific binding sites were blocked for one h at RT in Tris buffer (0.05 M Tris, 0.15 M NaCl, pH 7.4) containing 1% (v/v) cold water fish gelatin, 1% (v/v) normal goat serum (blocking buffer), and 0.01% (v/v) Triton X-100. Following blocking, cultures were incubated for one h at RT with one of the following rabbit antisera raised against bovine MIP 26 , b- or g-crystallins  diluted 1:100 in blocking buffer. Cultures were then washed several times and incubated with 10 mg of goat antiserum raised against rabbit IgG Oregon Green (Molecular Probes) per one ml of blocking buffer. Cultures were washed several times with blocking buffer prior to viewing by confocal microscopy.
Live cultures were viewed at 37 °C, while fixed cultures were viewed at RT on a Zeiss laser scanning confocal microscope model LSM 410 equipped with an Axiovert 100 inverted microscope, an Argon-Krypton 488/568/647 laser, a Delta-TC3 culture dish system (Bioptechs), a KP 600 line selection filter, a FT 488/568 Dichroic beam splitter, a LP 590 emission-filter for viewing Texas Red, a BP 515-540 emission filter for viewing Oregon-Green, and the software package LSM 3.993.
No changes in cell morphology were observed in sham- and BSA-loaded cells (Figure 1A,B) 1-2 h postloading. The postloaded cells appeared as a simple epithelial cell layer, which was identical to the appearance prior to loading with protein. The only apparent change noted was an approximate 10% increase in the number of rounded cells not adherent to the dish, which occurred immediately after the hypertonic/hypotonic loading procedures (Figure 1C). Plating of these hypertonic/hypotonic-loading solutions produced attached cells to culture well bottoms within 24 h. Following the sham- and BSA-loaded cells for an additional 4-7 days postloading did not produce any apparent changes in cell morphology, however, the number of rounded cells decreased within a 24 h period.
In initial experiments using WSF proteins from fiber cells loaded into epithelial cells, dramatic changes in cells were noticed very rapidly within 1-2 h postloading (Figure 1D). These changes included elongation of approximately 20% of the cells attached to the dish (Figure 1D, arrow), migration of cells to form multilayered, multicellular structures or aggregates of cells (Figure 1D, arrowhead) which constitutes approximately 20% of the cells, and an increase in cells not attached to the dish (approximately 10%), while other areas of a dish appeared as a monolayer of epithelial cells (approximately 50% of the cells).
To assess if any of these initial changes might be attributed to a specific class of crystallins, cells were loaded with a-, bhigh-, blow-, and g-crystallin fractions. The bhigh-, blow-, and g-crystallin fractions were separated and collected by gel filtration chromatography using a TSK-3000 column. The a-crystallin fraction was collected from the TSK-3000 column and further purified using a Biosep-SEC-S4000 column, to separate the a-crystallin fraction of approximately 800,000 Daltons, from the high molecular weight aggregate fraction that might contain aggregated components that could include b- and/or g-crystallins, and/or growth factors . The purified a-crystallins fraction was determined to consist of greater than 99% aA and aB using a C18 reverse phase column (250 x 4.6 mm, 300 angstrom pore size) with a flow rate of 1 ml/min, and a linear gradient of 5-60% (v/v) acetonitrile in 0.1% (v/v) trifluoroacetic acid, over 40 min (Figure 2). The purified a-crystallins fraction (Figure 1E) induced rapid morphological changes identical to the WSF (Figure 1D), while no apparent changes in cellular morphology were observed with the bhigh-, blow-, and g-crystallin fractions.
To determine if the biological effects observed with the a-crystallins fraction might be due to a-crystallin subunits, purified aA and aB subunits were tested for a biological effect on cell morphology. No apparent changes in initial or long-term (4-7 days) cellular morphology were observed with aA-crystallin. aB-crystallin, however, induced vary rapid changes in cell morphology (Figure 3), identical to that observed with the WSF and a-crystallins fraction.
If cells loaded with WSF and a-crystallins were followed over an additional four to seven days, the size of the multilayered cell regions or aggregates of cells continued to grow (Figure 4A,B). At this time, these multilayered, multicellular structures could be visualized with the unaided eye and measured one to several mm in size. There were between 1-5 of these large multilayered, multicellular structures per dish, and some of these structures were similar in width and length as represented in Figure 4B (approximately 300 to 400 mm x 800 mm), while others were much longer in one dimension as in Figure 4A (approximately 100 mm x 3,300 mm). Within these structures or along the outer edge, long and thin individual cells measuring in excess 100 mm in length and 2-12 mm in width were observed (Figure 4C, arrow, also see Figure 5E). In addition to the large multilayered, multicellular structures, cultures also contained numerous smaller multilayered, multicellular structures similar to those observed within the first four h postloading and large regions of the dish containing stratified cells with no definable edge region. Collectively, the large, small and stratified multicellular regions occupied approximately 80% of the dish area, while the remaining 20% of the dish was occupied by interspersed simple epithelial cells.
If cells loaded with aB-crystallin were followed over an additional four to seven days, with the exception of one out of 8 cultures, cell morphology did not change from those observed during the initial 4 h. In the one exception, one large multilayered, multicellular structure measuring approximately 3 mm by 0.8 mm was observed (Figure 6).
Using the method of osmotic lysis of pinocytic vesicles in combination with fluorescently tagged proteins, percentages of primary lens epithelial cells containing loaded proteins and localization over a 4-7 day period were assessed. Greater than 90% of cells were loaded with proteins by this method, while the fluorescence intensity among a given population of cells varied. Within the first one h postloading, loaded a-crystallins were localized to cytoplasmic structures and within some nuclei (Figure 5A,B). By 4 h postloading, loaded a-crystallins were preferentially localized throughout the cytoplasm of cells specifically found in multilayered, multicellular structures, with much lower fluorescence present in the surrounding monolayer cells, which did not appear to show changes in cell morphology (Figure 5C,D). By 4-7 days postloading, fluorescence was only faintly visible, and was localized diffusely throughout the cytoplasm of the cells with altered, elongated morphology (Figure 5E,F). This image was taken from the edge of a large multilayered, multicellular structure, to observe individual cells. After being fixed, permeablized and stained with the nucleic acid stain, SYTOX GREEN (Molecular Probes), cells in this region showed the presence of condensed and elongated nuclei (Figure 5F, arrowheads), similar to that seen in the bow region of the lens.
To determine if loading with a-crystallins resulted in differentiation of epithelial cells, as determined by expression of specific marker proteins, antiserum to the differentiation markers MIP 26, b-crystallin and g-crystallin were used to localize expression of these proteins in cultures four to seven days postloading (Figure 7). Representative differential interference contrast images of multilayered, multicellular structures are present in Figure 7A,C,E, followed by their corresponding individual optical sections showing the localization of differentiation markers (Figure 7B,D,F), while Figure 7G,H,I were taken from a different multilayered area within the same culture. As seen in these images, MIP 26, b- and g-crystallins were localized to cells throughout the multilayered areas, both in cells within multilayered, multicellular structure (Figure 7B,D,F) and in cells present in stratified cell layers with no definable edge (Figure 7G,H,I). MIP 26, b- and g-crystallins were not localized in monolayer regions or in cells loaded with BSA, and no fluorescence was observed in cells loaded with a-crystallins when the primary antiserum was replaced with normal rabbit serum. Taken together, the results indicate that loading of cells only with a-crystallins, results in the expression of MIP 26, b-crystallin, and g-crystallin components that have been associated with differentiation of epithelial cells.
The results of the current study support the hypothesis that a-crystallins, which show increased expression in epithelial cells at the elongation zone in situ [1-9], may also play an active role in the differentiation process. For this purpose, the method of osmotic lysis of pinocytic vesicles was found to be a relatively fast and efficient method for bulk loading of lens epithelial cells in culture with exogenous proteins. This observation is consistent with similar studies with other cell types [37-39]. The results also indicate that the method and not the protein loaded into cells results in approximately 10% of the cells losing attachment to the dish. In the present study, the experiments using fluorescently labeled proteins verified that protein was indeed loaded into cells. Using this method, however, it was not possible to determine exact concentrations of protein delivered to each cell. The fluorescence intensity of individual cells varied within a given culture. This may indicate that not all cells in a given culture were loaded with equal concentrations of protein or that some cells may have digested some or more of the internalized protein than other cells or that some protein may have been transferred to adjacent cells by intercellular processes. This might explain why the biological effects within a given culture varied and why not all cells became elongated and began to form multilayered, multicellular structures approximately 4 h after loading with a-crystallins. In preliminary studies, we determined that adding less than 20 ml (5, 10, 15 ml, 10 mg protein per ml solution) of the protein sample during the loading procedure did not produce any apparent rapid changes in epithelial cell morphology. This indicates that the concentration of a-crystallins per cell may be an important factor in determining the biological response. Further experiments are needed to determine what concentration of a-crystallins are required to generate a biological response to epithelial cells and to determine if variations in protein concentration in different cells is solely responsible for the variations seen in a given culture.
The present studies show rapid changes in cell morphology and production of lentoid bodies occurs within 4 h postloading with a-crystallins and aB-crystallin. Lentoid body production induced by various growth factors, however, have been reported to require several days to weeks [3,10-22] to produce similar changes as those produced within 4 h with a-crystallins. In addition, these previous studies using growth factors have generally been unable to induce further changes in cell morphology in 4-7 day postinduced cultures as that observed in the present study with a-crystallins. The results of present study show that after 4-7 days postloading with a-crystallins, the epithelial cells further change in morphology, forming multicellular elongated structures that can be seen with the unaided eye. The length and elongated structure of these cells is reminiscent of differentiating cells in the elongation zone of the intact lens. Furthermore, visualization of fluorescently labeled a-crystallins loaded into cells after 4 h clearly shows that these cells undergoing morphological changes were loaded with the fluorescently labeled protein. Loading of cells with a-crystallins does indeed result in cell differentiation, as shown by the subsequent expression of the markers MIP 26, b- crystallin, and g-crystallin. Taken together, the results of the present study suggest an important role, if not a necessary role, for a-crystallins in the differentiation of lens epithelial cells.
It is possible but highly unlikely that the effects on cell morphology observed with a-crystallins were do to a minor contaminant that co-eludes with the a-crystallins fraction. The reverse phase HPLC profile indicated that greater than 99% of the SEC-4000 a-crystallins fraction consisted of aB- and aA-crystallins. Past studies  have shown that this SEC-4000 a-crystallins fraction does not contain immunoreactive b- or g-crystallins. The finding that purified-renatured aB-crystallin produced identical results within the first four h postloading to that of a-crystallins reduces the possibility that the morphological changes observed with a-crystallins were do to a contaminant. In addition, the results were not do to some minor contaminant associated with the fluorochrome Texas red, because identical results were obtained with unlabeled a-crystallins. Taken together, the results of the present study indicate that a-crystallins were responsible for the morphological changes observed in cells.
Based upon previous studies of lens epithelial cells in culture [3,10-32], expression of a-crystallins may not be an initiating event in differentiation, but may rather be an intermediate in a cascade of events that might begin with specific growth factors binding to receptors on lens epithelial cells. Binding of growth factors by cell receptors could generate secondary messages within epithelial cells, resulting in the expression of immediate-early genes [42-44], that would eventually activate expression of genes for the a-, b-, and g-crystallins which would result in differentiation of epithelial cells into fiber cells. In the present study, the finding that b-crystallin and g-crystallin did not result in epithelial cell differentiation does not completely eliminate the possibility that these proteins are without biological effects. Based upon their temporal expression during the differentiation process in vivo [1-9], it is possible that these proteins might play a role in morphological and/or biochemical changes further downstream in the differentiation process, after large amounts of a-crystallin are first expressed.
The exact molecular mechanism of how a-crystallins might be involved in lens epithelial cell differentiation remains to be determined. Previous studies have shown that the a-crystallins can act as molecular chaperones, binding to partially denatured proteins both in vitro [45,46] and probably in vivo , to inhibit further denaturation. The a-crystallins also have hydrophobic domains that have been shown to interact with the lipid bilayer of cell membranes [47-50]. In addition, recombinant aB-crystallin has been shown to ectopically localize to interphase nuclei of transfected CHO cells suggesting a regulatory role of aB in the nucleus . Finally, there is extensive evidence from previous studies, demonstrating that a-crystallins interact with cytoskeletal components. Both aA- and aB-crystallin can bind specifically to actin, both in vitro  and in vivo . In the lens, a-crystallins also form a complex with type III intermediate filament proteins and the beaded filament proteins CP49 and CP115, resulting in filament assembly in the presence of a-crystallins . It is therefore possible that increased synthesis of the a-crystallins in epithelial cells during the differentiation process may have profound effects upon the cytoskeleton, which may in turn profoundly affect cell shape, as seen in the present study. Whatever the mechanism(s) of its interactions with cellular components, the results of the present study suggest that in addition to its roles as a molecular chaperone and as a substance to increase the refractile properties of the lens, the massive amounts of a-crystallin synthesized during elongation of epithelial cells may also play a key role in the differentiation process. Further studies, now in progress, will hopefully ascertain the importance of this role, as well as further characterize the specific interactions of a-crystallins with cellular components that may be necessary events in the functional importance of this protein.
This work is supported by a grant from the National Institutes of Health and supported in part by the Kansas NSF EPSCoR program, by the Kansas NASA EPSCoR program, by University resources, and by the Kansas Agricultural Experiment Station.
1. Rafferty NS. Lens morphology. In: Maisel H, editor. The ocular lens: structure, function, and pathology. New York: Dekker; 1985. p.1-53.
2. Vermorken AJ, Hilderink JM, van de Ven WJ, Bloemendal H. Lens differentiation. Crystallin synthesis in isolated epithelia from calf lenses. J Cell Biol 1978; 76:175-83.
3. Chamberlain CG, McAvoy JW. Fibre differentiation and polarity in the mammalian lens: a key role for FGF. Prog Retin Eye Res 1997; 16:443-8.
4. Beebe DC, Piatigorsky J. Differential synthesis of crystallin and noncrystallin polypeptides during lens fiber cell differentiation in vitro. Exp Eye Res 1976; 22:237-49.
5. McAvoy JW. Cell division, cell elongation and distribution of alpha-, beta-, and gamma-crystallins in the rat lens. J Embryol Exp Morphol 1978; 44:149-65.
6. McAvoy JW. Cell division, cell elongation and the co-ordination of crystallin gene expression during lens morphogenesis in the rat. J Embryol Exp Morphol 1978; 45:271-81.
7. Piatigorsky J. Lens differentiation in vertebrates. A review of cellular and molecular features. Differentiation 1981; 19:134-53.
8. Van Leen RW, Breuer ML, Lubsen NH, Schoenmakers JG. Developmental expression of crystallin genes: in situ hybridization reveals a differential localization of specific mRNAs. Dev Biol 1987; 123:338-45.
9. Yancey SB, Koh K, Chung J, Revel JP. Expression of the gene for main intrinsic polypeptide (MIP): separate spatial distributions of MIP and beta-crystallin gene transcripts in rat lens development. J Cell Biol 1988; 106:705-14.
10. Klok EJ, Lubsen NH, Chamberlain CG, McAvoy JW. Induction and maintenance of differentiation of rat lens epithelium by FGF-2, insulin and IGF-1. Exp Eye Res 1998; 67:425-31.
11. Shang F, Gong X, McAvoy JW, Chamberlain C, Nowell TR Jr, Taylor A. Ubiquitin-dependent pathway is up-regulated in differentiating lens cells. Exp Eye Res 1999; 68:179-92.
12. Richardson NA, McAvoy JW. Age-related changes in fibre differentiation of rat lens epithelial explants exposed to fibroblast growth factor. Exp Eye Res 1990; 50:203-11.
13. Liu J, Chamberlain CG, McAvoy JW. IGF enhancement of FGF-induced fibre differentiation and DNA synthesis in lens explants. Exp Eye Res 1996; 63:621-9.
14. Peek R, McAvoy JW, Lubsen NH, Shoenmakers JG. Rise and fall of crystallin gene messenger levels during fibroblast growth factor induced terminal differentiation of lens cells. Dev Biol 1992; 152:152-60.
15. Ibaraki N, Lin LR, Reddy VN. Effects of growth factors on proliferation and differentiation in human lens epithelial cells in early subculture. Invest Ophthalmol Vis Sci 1995; 36:2304-12.
16. Richardson NA, Chamberlain CG, McAvoy JW. IGF-1 enhancement of FGF-induced lens fiber differentiation in rats of different ages. Invest Ophthalmol Vis Sci 1993; 34:3303-12.
17. Richardson NA, McAvoy JW, Chamberlain CG. Age of rats affects response of lens epithelial explants to fibroblast growth factor. Exp Eye Res 1992; 55:649-56.
18. Lovicu FJ, de Iongh RU, McAvoy JW. Expression of FGF-1 and FGF-2 mRNA during lens morphogenesis, differentiation and growth. Curr Eye Res 1997; 16:222-30.
19. Chamberlain CG, McAvoy JW, Richardson NA. The effects of insulin and basic fibroblast growth factor on fibre differentiation in rat lens epithelial explants. Growth Factors 1991; 4:183-8.
20. McAvoy JW, Chamberlain CG. Fibroblast growth factor (FGF) induces different responses in lens epithelial cells depending on its concentration. Development 1989; 107:221-8.
21. Azuma M, Shearer TR. Induction of elongation in cultured rat lens epithelial cells by FGF and inhibition by selenite. Invest Ophthalmol Vis Sci 1992; 33:2528-31.
22. Lovicu FJ, McAvoy JW. Structural analysis of lens epithelial explants induced to differentiate into fibres by fibroblast growth factor (FGF). Exp Eye Res 1989; 49:479-94.
23. Yamamoto Y. Growth of lens and ocular environment: role of neural retina in the growth of mouse lens as revealed by implantation experiment. Dev Growth Differ 1976; 18:273-8.
24. McAvoy JW. Beta- and gamma-crystallin synthesis in rat lens epithelium explanted with neural retina. Differentiation 1980; 17:85-91.
25. McAvoy JW, Fernon VT. Neural retinas promote cell division and fibre differentiation in lens epithelial explants. Curr Eye Res 1984; 3:827-34.
26. Campbell MT, McAvoy JW. Onset of fibre differentiation in cultured rat lens epithelium under the influence of neural retina-conditioned medium. Exp Eye Res 1984; 39:83-94.
27. Campbell MT, McAvoy JW. A lens fibre differentiation factor from calf neural retina. Exp Cell Res 1986; 163:453-66.
28. McAvoy JW, Campbell MT, Walton J. Influence of neural retina on lens fibre differentiation in rats. Aust N Z J Ophthalmol 1985; 13:103-11.
29. Walton J, McAvoy J. Sequential structural response of lens epithelium to retina-conditioned medium. Exp Eye Res 1984; 39:217-29.
30. McAvoy JW, Richardson NA. Nuclear pyknosis during lens fibre differentiation in epithelial explants. Curr Eye Res 1986; 5:711-5.
31. Richardson NA, McAvoy JW. Analysis of an inductive interaction between lens and neural retina in rat of different ages. Exp Eye Res 1986; 43:1031-42.
32. Richardson NA, McAvoy JW. Age-related changes in fibre differentiation of rat lens epithelial cells in vitro. Exp Eye Res 1988; 46:259-67.
33. Nagineni CN, Bhat SP. Lens fiber cell differentiation and expression of crystallins in co-cultures of human fetal lens epithelial cells and fibroblasts. Exp Eye Res 1992; 54:193-200.
34. Takemoto L, Boyle D. Molecular chaperone properties of the high molecular weight aggregate from aged lens. Curr Eye Res 1994; 13:35-44.
35. McFall-Ngai MJ, Ding LL, Takemoto LJ, Horwitz J. Spatial and temporal mapping of the age-related changes in human lens crystallins. Exp Eye Res 1985; 41:745-58.
36. Perry RE, Abraham EC. High-performance liquid chromatographic separation of lens crystallins and their subunits. J Chromatogr 1986; 351:103-10.
37. Okada CY, Rechsteiner M. Introduction of macromolecules into cultured mammalian cells by osmotic lysis of pinocytic vesicles. Cell 1982; 29:33-41.
38. McElligott MA, Dice JF. Microinjection of cultured cells using red-cell-mediated fusion and osmotic lysis of pinosomes: a review of methods and applications. Biosci Rep 1984; 4:451-66.
39. Lee G, Delohery TM, Ronai Z, Brandt-Rauf PW, Pincus MR, Murphy RB, Weinstein IB. A comparison of techniques for introducing macromolecules into living cells. Cytometry 1993; 14:265-70.
40. Boyle DL, Takemoto LJ. Localization of MIP 26 in nuclear fiber cells from aged normal and age-related nuclear cataractous human lenses. Exp Eye Res 1999; 68:41-9.
41. Boyle D, Takemoto L, Charaterization of the alpha-gamma and alpha-beta complex: evidence for an in vivo functional role of alpha-crystallin as a molecular chaperone. Exp Eye Res 1994; 58:9-15.
42. Nath P, Getzenberg R, Beede D, Pallansch L, Zelenka P. c-myc mRNA is elevated as differentiating lens cells withdraw from the cell cycle. Exp Cell Res 1987; 169:215-22.
43. Cvekl A, Kashanchi F, Brady JN, Piatigorsky J. Pax-6 interactions with TATA-box-binding protein and retinoblastoma protein. Invest Ophthalmol Vis Sci 1999; 40:1343-50.
44. Kawauchi S, Takahashi S, Nakajima O, Ongino H, Morita M, Nishizawa M, Yasuda K, Yamamoto M. Regulation of lens fiber cell differentiation by transcription factor c-Maf. J Biol Chem 1999; 274:19254-60.
45. Horwitz J. Alpha crystallin can function as a molecular chaperone. Proc Natl Acad Sci U S A 1992; 89:10449-53.
46. Gopalakrishnan S, Boyle D, Takemoto L. Preferential interaction of alpha crystallin with denatured forms of gamma crystallin. Invest Ophthalmol Vis Sci 1994; 35:382-7.
47. Spector A, Garner MH, Garner WH, Roy D, Farnsworth P, Shyne S. An extrinsic membrane polypeptide associated with high-molecular-weight protein aggregates in human cataract. Science 1979; 204:1323-6.
48. Bloemendal H, Hermsen T, Dunia I, Benedetti EL. Association of crystallins with the plasma membrane. Exp Eye Res 1982; 35:61-7.
49. Ifeanyi F, Takemoto L. Specificity of alpha crystallin binding to the lens membrane. Curr Eye Res 1990; 9:259-65.
50. Boyle DL, Takemoto L. EM immunolocalization of alpha crystallins: association with the plasma membrane from normal and cataractous human lenses. Curr Eye Res 1996; 15:577-82.
51. Bhat SP, Hale IL, Matsumoto B, Elghanayan D. Ectopic expression of alpha B-crystallin in Chinese hamster ovary cells suggests a nuclear role for this protein. Eur J Cell Biol 1999; 78:143-50.
52. Gopalakrishnan S, Boyle D, Takemoto L. Association of actin with alpha crystallin. Trans Kans Acad Sci 1993; 96:7-12.
53. Del Vecchio PJ, MacElroy KS, Rosser MP, Church RL. Association of alpha-crystallin with actin in cultured lens cells. Curr Eye Res 1984; 3:1213-9.
54. Carter JM, Hutcheson AM, Quinlan RA. In vitro studies on the assembly properties of the lens proteins CP49, CP115: coassembly with alpha-crystallin but not with vimentin. Exp Eye Res 1995; 60:181-92.