Using poly(lactide-co-glycolide) electrospun scaffolds to deliver cultured epithelial cells to the cornea
The loss of limbal corneal epithelial stem cells reduces the transparency of the cornea [1,2]. To replace these lost stem cells, several methods have been developed to transplant autologous or allogeneic cultured limbal epithelial cells. To transplant cells onto the cornea, a carrier is necessary as the cultured cells are unlikely to survive if placed directly on this challenging wound bed [3]. Cells have been transferred by culturing them into integrated sheets that were wrapped around contact lenses or petrolatum gauze for transplantation onto the eye [4] or by placing the cells on fibrin [1], but the most com-monly used method is the use of the human amniotic membrane as a delivery dressing [5,6]. Cultured cells are placed on small pieces of the prepared amniotic membrane and this is sutured, stromal-side-down, onto the eye. The membrane degrades over a few weeks leaving the cells in place.
However, the response to the amniotic mem-brane is dependent on the host response to the donor tissue [7,8] and even though the amnio-tic membrane has been used for many years without any transmission of disease, there have been some cases of immunorejection following multiple treatments using membranes from different donors [9,10]. Furthermore, despite the tissue-banked amniotic membrane undergoing vigorous screening for approximately 6 months [11], transmission of diseases is only minimized and occult diseases may not be not detected [12]. Another problem in practice is that human
amnion needs to be prepared for use, and cell performance on the amnion can vary as a function of this [13,14].
An ideal delivery carrier for corneal cells must be biocompatible with minimal risk of disease transmission (preferably none), achieve good cell adhesion and have appropriate tensile strength to secure to the cornea, and undergo biodegradation leaving the transplanted cells in place [15].
Synthetic scaffolds that can achieve the former will minimize the very slight chance of viral dis-ease transmission, which cannot be fully elimi-nated with the use of donated human tissue. They will also remove the inherent variability seen with donor amnion.
We recently reported on developing electro-spun polymers as 3D scaffolds to replace the human dermis [16]. In this study, we now seek to develop a rapidly degradable synthetic carrier membrane for delivering cultured corneal cells to damaged corneas.
Electrospinning is simple and reproducible, and can produce fibers with diameters ranging from 3 nm to 6 µm [17,18]. Mats of electrospun scaffolds placed together can roughly mimic the extracellular matrix of tissues [19] and sup-port cell growth. Furthermore, it is relatively simple to tune the rate of breakdown of these scaffolds by varying the ratio of polylactide acid (PLA) to polyglycolide acid (PGA) as reported by Blackwood et al [16]. In this study, scaffolds of 100% PLA survived, intact, for a year post-implantation in rats. Scaffolds of 75% PLA and
Aims: To assess the potential of electrospun poly(lactide-co-glycolide) membranes to provide a biodegradable cell carrier system for limbal epithelial cells. Material & methods: 50:50 poly(lactide-co-glycolide) scaffolds were spun, sterilized and seeded with primary rabbit limbal epithelial cells. Cells were cultured on the scaffolds for 2 weeks and then examined by confocal microscopy, cryosectioning and scanning-electron microscopy. The tensile strength of scaffolds before and after annealing and sterilization was also studied. Results: The limbal cells had formed a continuous multilayer of cells on either side of the scaffold. Scaffolds with cells showed signs of the onset of degradation within 2 weeks in culture media at 37°C. Scaffolds that were annealed resulted in a more brittle and stiff mat. Conclusions: We suggest this carrier membrane could be used as a replacement for the human amniotic membrane in the treatment of limbal stem cell deficiency, lowering the risk of disease transmission to the patient.
KEYWORDS: cell carrier n cornea n electrospun n limbal n poly(lactide-co-glycolide) Pallavi Deshpande1, Rob McKean2, Keith A Blackwood1, Richard A Senior1, Adekemi Ogunbanjo1, Anthony J Ryan2
& Sheila MacNeil†1
1Department of Engineering Materials, Kroto Research Institute, University of Sheffield, North Campus, Broad Lane, Sheffield, S3 7HQ, UK 2Department of Chemistry, University of Sheffield, Sheffield S3 7HF, UK †Author for correspondence: Tel.: +44 114 222 5995 Fax: +44 114 222 5943 [email protected]
ReseaRch aRticle Deshpande, McKean, Blackwood et al.
Regen. Med. (2010) 5(3)396 future science group
25% PGA underwent a 50% reduction after 3 months, while scaffolds of 50% PLA and 50% PGA started to lose structural integrity after only 2 weeks. These latter scaffolds degraded too fast to be used for dermal scaffolds (which will ideally be replaced by new dermal collagen, produced by the tissue-engineered skin cells, over several months) but their relatively rapid degradation seems a reasonable time scale for delivery of cul-tured corneal cells with progressive dissolution of the underlying membrane.
Accordingly, the aim of the current study was to begin to evaluate the use of a 50:50 poly(lactide-co-glycolide) (PLGA) electrospun fiber mat as a potential carrier for limbal epi-thelial cells as a lower-risk clinical option to the use of human donor amnion. We investigated the effect of heat annealing and sterilization on the mechanical properties of the scaffold, and assessed how well the corneal epithelial cells attached to the scaffold. We also assessed how well scaffolds with cells could be handled and sutured, and looked at the initial signs of scaffold breakdown within the first 2 weeks of placing cells on these membranes.
Methods n Rabbit limbal epithelial cells
Rabbit limbal epithelial cells were isolated from commercial white rabbits derived from the New Zealand white rabbit. The rabbits typi-cally weighed 2.4–2.6 kg. As described earlier [20], the limbal rim of the cornea was excised into four segments and placed into 2.5 mg/ml of dispase II (in 10% Dulbecco’s modified Eagle medium) for 45 min. The limbal cells were then scraped off the tissues into phosphate-buffered saline using a pair of blunt forceps. The solution was collected and centrifuged at 1000 rpm for 5 min; the supernatant was discarded and the cells were resuspended in fresh media.
The cells were seeded into flasks along with growth-arrested 3T3 murine fibroblasts as a feeder layer. The culture media for the rabbit lim-bal epithelial cells was Green’s media [21], consist-ing of Dulbecco’s modified Eagle medium and Ham’s F12 medium (Gibco®, Invitrogen Life Technologies, CA, USA) in a 3:1 ratio supple-mented with 10% foetal calf serum, 10 ng/ml EGF (R&D Systems, MN, USA), 0.4 mg/ml hydrocortisone, 1.8 × 10-4 mol/l adenine, 5 mg/ml insulin, 5 mg/ml transferrin, 2 × 10-3 mol/l glutamine, 2 × 10-7 mol/l triiodothyronine (Sigma-Aldrich, UK), 0.625 mg/ml amphoter-icin B, 100 IU/ml penicillin and 100 mg/ml streptomycin (Gibco).
n Fabrication of PLGA scaffoldsPoly(lactide-co-glycolide) (50:50; Sigma-Aldrich) of molecular weight 40,000–75,000 was electrospun as previously described [16]. Briefly, the polymers were dissolved in dichlo-romethane at a concentration of 25% (wt/wt). The polymer was passed through a 2 ml syringe with a stainless steel needle with an internal diameter of 0.8 mm. The solution was delivered at a constant flow rate of 1.5 ml/h using a syringe pump. The polymer was electrospun at a volt-age of 10 kV, supplied by a high-voltage source (Alpha series III, Brandenburg, Germany). Sheets of the scaffold were collected onto alu-minum foil sheets wrapped around an earthed aluminum rotating collector (300 rpm). Jet formation was assisted by placing an aluminum ring at 10 kV approximately 5 mm behind the tip of the syringe needle.
n Sterilization of scaffoldsScaffolds were sterilized by either immersion in 70% cold ethanol for 10 min or 0.01% perace-tic acid for 3 h using a protocol we previously assessed for human donor skin [22].
n Culture of cells on the PLGA scaffoldsPrimary rabbit limbal epithelial cells were seeded onto a double-layered sheet of the nontreated scaffold sterilized by immersion into 70% ethanol.
The cells were seeded onto the scaffolds (two single sheets of scaffold placed together) using a stainless steel ring, and after 24 h in culture, the ring was removed. After 2 days under submerged conditions, the scaffolds were placed on a steel-mesh grid, bringing the scaffolds to an air–liquid interface with a media change every 2 days. On day 14, the cells on the scaffolds were fixed using 10% buffered formaldehyde. The rabbit limbal epithelial cells were stained with CellTracker™ Red (Invitrogen, Paisley, UK) according to the manufacturer’s protocol. The cells were visual-ized using the Zeiss LSM 510 META (Zeiss, Welwyn, UK) confocal microscope histology and scanning-electron microscopy (SEM).
n Preparation of scaffolds for histologyAfter fixation in 10% buffered formaldehyde for 24 h, samples were cut in half with a scalpel and mounted in optimal cutting temperature compound (VWR International, West Sussex, UK) to expose the center of the scaffold. Samples were sectioned using a Leica CM3050 S cryostat
Using PLGA electrospun scaffolds to deliver epithelial cells to the cornea ReseaRch aRticle
www.futuremedicine.com 397future science group
(Leica Microsystems, Germany) at a thickness of 8 µm. Sections were mounted on 3-aminopro-pyltriethoxysilane (Sigma-Aldrich)-coated slides that were prepared by a brief dip in 3% 3-amino-propyltriethoxysilane (in acetone), washed three times in water and allowed to dry. For histologi-cal staining, slides were washed gently in water for 1 min to remove optimal cutting temperature compound, followed by staining in hematoxylin for 1 min 30 s and washed in water again for 2 min. Sections were then immersed briefly in water and a coverslip was placed on the section using aqueous mountant. Sections were examined under light microscopy.
n Preparation of scaffolds for SEMScaffolds were prepared for SEM, as described previously [16]. Approximately 25 nm of gold was sputter-coated onto the scaffolds and then the samples were examined using a SEM (FEI XL-20 SEM, Philips, Guildford, UK) at an accelerating voltage of 10–15 kV.
n Mechanical testing of the scaffoldsScaffolds (heat treated at 60°C for 3 h or non-heat treated) were cut into dog-bone shapes as shown in Figure 1. Using the Bose Electroforce® 3100 tensile test machine (Bose, MN, USA), the scaffolds were clamped in place via stainless steel clamps and pulled apart at a rate of 1 mm/10 s over a distance of 6 mm. The force exerted was measured via a 22 N small-load cell. A total of six samples were used for each experimental con-dition. Scaffolds were also tested poststeriliza-tion in 0.01% peracetic acid for 3 h compared with dry scaffolds presterilization.
StatisticsThe difference between the mechanical proper-ties of scaffolds was assessed using analysis of variance (ANOVA) and p < 0.05 were considered significant.
ResultsPrimary rabbit limbal epithelial cells were used in this study as in the UK there is a scarcity of human donor corneas for experimental research. Investigating corneal cell growth on the scaf-folds by confocal microscopy showed a con-tinuous layer of rhodamine-stained epithelial cells on the surface of the scaffold (Figure 2A–C). Cryosectioning, and hematoxylin staining of the scaffolds after 2 weeks of cell culture dem-onstrated that the control scaffolds without cells measured approximately 100 µm in thickness and there was, as expected, no hematoxylin staining
(Figure 3A). Scaffolds with epithelial cells cultured on them for 14 days showed clear hematoxylin nuclei staining (Figure 3B) and the thickness of the scaffold was approxiamtely 150 µm. A multilayer of cells was observed on both the top and bottom surfaces of the scaffold with a lower density of cells throughout the scaffold.
Scanning-electron microscopy of the scaffolds without cells showed random fibers of approxi-mately 3–8 µm (Figure 4A) and that a cross-section of the scaffolds measured approximately 100 µm in thickness after 2 weeks in media (Figure 4B). The cultured corneal cells formed a layer on the surface of the scaffold (Figure 4C & D), confirm-ing both the histology and confocal microscopy results, and the thickness of the scaffold was now approximately 70 µm. The reduction in the thickness of the scaffold assessed postcell culture by SEM (but not seen with hematoxylin staining where it apparently increased ) requires further investigation. A comparison of SEM sections of scaffolds, with and without cells, suggested that the fibers without cells appeared more loosely
15 mm
20 mm
8 mm
5 mm
5 mm
Figure 1. Dimensions of the ‘dog-bone’ used to assess scaffold mechanical properties.
100 µm
100 µm
100 µm
Figure 2. (A) Front view and (B & C) side view of confocal images of the rabbit limbal epithelial cells stained with CellTracker™ Red and cultured on the electrospun scaffolds.
ReseaRch aRticle Deshpande, McKean, Blackwood et al.
Regen. Med. (2010) 5(3)398 future science group
arranged than the fibers of the scaffolds with cells (Figure 4B & D) but the preparation of samples for SEM may have introduced fixation artefacts, so caution is needed in interpreting these results and further work will be needed to examine the fate of the scaffold versus time.
Further investigations looked at the handling and initial breakdown of scaffolds with cells on them. Scaffolds that were freshly sterilized and seeded with cells were transparent, easy to handle and could be readily sutured to metal grids the fol-lowing day (Figure 5A). Degradation of the scaffolds was then studied by culturing the rabbit limbal epithelial cells on the scaffolds at an air–liquid interface. After 2 weeks in culture, the scaffolds started to degrade and reduce in area. By this time
they appeared to be slightly opaque (Figure 5C). During the second week of culture, the scaffolds with cells on them were slightly brittle to handle (and difficult to suture) compared with the first week when they were easy to handle and suture.
It has been reported that heat treating the scaf-fold reduces the shrinkage of PLGA scaffolds [23]. In addition to this, peracetic acid was used to sterilize the scaffolds in the tensile study as it has been previously used in sterilization of dermis [22].
Final experiments focused on the physical properties of the scaffolds after heat annealing and sterilization with peracetic acid. Heat annealing is a simple way to put together several layers of thin scaffold to make a thicker membrane and steriliza-tion is necessary in preparing to take biomaterials such as these to the clinic.
Wet-annealed scaffolds led to a more brittle sheet of fibers that broke on the application of a smaller force compared with the dry-annealed scaffolds (p < 0.05) as summarized in TABle 1. Annealing also increased the stiffness (Young’s modulus) of the dry scaffolds (p < 0.05); however, there was no significant difference in Young’s modulus on wetting the scaffolds. Sterilizing the scaffolds with peracetic acid also led to a significant reduction in the tensile strength of the heat-annealed scaffolds (p < 0.01), as shown in TABle 1, but reduced the stiffness. It was also observed that the nonannealed scaffolds were easier to handle compared with the annealed scaffolds, which became brittle by day 14.
DiscussionThe aim of this study was to begin to assess the potential of rapidly degrading PLGA scaffolds as an alternative to the human amniotic membrane to treat corneal diseases. Increasingly, patients who have lost limbal epithelial stem cells are being treated with laboratory-expanded limbal stem cells (from the contralateral eye where this is unaffected or allogeneic cells where both eyes are affected). When it comes to delivering these cells back to the cornea, the most popular meth-odology is that of growing them on small pieces of human donor amniotic membrane and then suturing the membrane back onto the eye. Many hundreds of patients have now been treated using this approach with very encouraging results overall [24–26].
Once sutured onto the eye, the amniotic membrane usually disintegrates within 1 month after surgery. However, it has been reported that, owing to donor variations and handling before surgery, treatment outcomes using the amniotic membrane are not consistent [13,14]. Another
50 µm
50 µm
Figure 3. Cryosection of electrospun scaffolds (A) without cells and (B) after 2 weeks culture with cells.
Using PLGA electrospun scaffolds to deliver epithelial cells to the cornea ReseaRch aRticle
www.futuremedicine.com 399future science group
issue is risk management. Blood samples are taken from donors at the point of delivery of the baby and then again some 6 months later to screen for the presence of the HIV. During this time, the amniotic membrane must be banked but not released, hence, it requires careful bank-ing procedures to reduce the risk of viral disease transmission. This can be achieved with good tissue banking practices but the real costs of these are substantial. In addition, reports suggest that the amniotic membrane not only functions as a carrier, but also provides growth factors and cytokines, which may be important in epithe-lial–stromal interactions on the ocular surface, as well as maintenance of the microenvironment of the limbal cell niche [27].
In view of these issues, many groups are working towards an alternative to the amniotic membrane, and materials such as recombinant
collagen gels [28], synthetic polymer gels [29], contact lenses [20,30] and temperature-sensitive synthetic polymers [31] have all been developed as cell carriers.
In this study, we propose a very simple approach of using a PLGA scaffold that degrades rapidly. PLGA scaffolds undergo a hydrolytic chain scission of the ester bonds in their back-bone, eventually leading to the production of carbon dioxide and water [32]. The rate of break-down of the scaffolds can be controlled to a use-ful degree by varying the ratio of PLA (which has a hydrophobic methyl backbone and tends to degrade very slowly) to PGA (more hydrophilic and degrades more rapidly). In general, cells attach well to these scaffolds. It is thought that, in the presence of serum, the fibronectin and vitronectin adhere to the polymer surface and the cells, in turn, adhere to the proteins [33,34].
100 µm
100 µm 20 µm
20 µm
Figure 4. Scanning-electron microscopy images of scaffolds. (A) Front view and (B) cross-sectional view of cell-free scaffolds. (C) Front view and (D) cross-sectional view of scaffolds cultured with limbal epithelial cells for 14 days.
Figure 5. Images of the scaffolds with limbal epithelial cells on (A) day 0 sutured to a metal grid, (B) day 1 and (C) day 14.
ReseaRch aRticle Deshpande, McKean, Blackwood et al.
Regen. Med. (2010) 5(3)400 future science group
Recent studies have also demonstrated the use of PGA scaffolds in corneal tissue engineering [35,36], where PGA has been used to replace the stroma of the cornea. Our results from cultur-ing skin cells on PLGA scaffolds demonstrated that the cells became well organized, and both keratinocytes and fibroblasts rapidly (within 7 days) produced new collagen on these fibers [16]. Accordingly, our aim in this study was to evaluate whether scaffolds of 50% PLA and 50% PGA would be suitable candidates to replace amniotic membrane.
We used rabbit corneal epithelial cells, which attached well to the scaffolds, and we looked at the handling of the scaffolds with cells at an early time point (within the first few days of seeding of the cells) and after 2 weeks. The scaffolds supported good growth of corneal epithelial cells, achieving a confluent layer within 2 weeks and the scaffolds could be readily sutured (in this experimental pro-tocol to metal mesh grids) within the first week of culture. This would be ideal for clinical use – cells could be seeded on the scaffolds and then a day or two later, the scaffolds could be sutured to the eye.
The next issue was the breakdown of the scaf-folds and it was obvious that there were major changes happening by week 2, the scaffolds became opaque and more brittle in their handling and appeared to be visibly shrinking. We inter-pret these changes as evidence that the scaffolds are beginning to break down.
Initially, we used ethanol sterilization for our experimental work but, while convenient for laboratory experiments, this is not a recognized sterilization methodology that can be taken to
the clinic. We then introduced peracetic acid steri-lization, which is clinically approved and is also a wet-sterilization methodology. This also reduced scaffold strength but improved pliability and did not affect the interaction of cells with the scaf-fold fibers. A recent study from our laboratory has demonstrated that there is no significant differ-ence in the viability of epithelial cells and fibrob-lasts on scaffolds sterilized with either peracetic acid, g-irradiation or ethanol [37].
Tensile studies looked at the pros and cons of annealing thin layers of electrospun scaffolds to make a thicker material for handling. While this worked well in terms of producing a thicker scaffold, the mechanical properties and flexibility of the scaffold were compromised making these harder to handle.
In conclusion, these initial in vitro results are very encouraging for using simple electro-spun scaffolds that could be stored sterilized and dry as an off-the-shelf product and then seeded with cells and sutured (or fibrin glued) onto the patient’s cornea within a few days. The next challenge will be to assess the perform-ance of these scaffolds with cells in a cornea model (ex vivo, as we have recently used [20], or in vivo) to assess the fate of the corneal cells as the membrane breaks down.
Financial & competing interests disclosureThe authors have no relevant affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript. This includes employment, consultancies, honoraria, stock ownership or options, expert testimony, grants or patents received or pending, or royalties.
No writing assistance was utilized in the production of this manuscript.
Ethical conduct of research The authors state that they have obtained appropriate institutional review board approval or have followed the principles outlined in the Declaration of Helsinki for all human or animal experimental investigations. In addition, for investi gations involving human subjects, informed consent has been obtained from the participants involved.
Table 1. Mechanical testing of poly(lactide-co-glycolide) scaffolds.
Treatment Maximum yield(×106) (N/m2)
Maximum strain Young’s Modulus(×106)
Nonheat treated
Dry 0.53 ± 0.19 26 ± 14% 5.1 ± 2.3
Wet 0.29 ± 0.08 17.7 ± 6.5% 3.8 ± 1.32
Heat treated (60°C for 3 h)
Dry 0.88 ± 0.28 22.4 ± 3.2% 10.5 ± 2.2
Wet 0.38 ± 0.29 9 ± 4.7% 7.6 ± 2.9
Dry: No treatment; Wet: Sterilized with 0.01% peracetic acid for 3 h prior to testing.
Executive summary
� Simple, rapidly degrading electrospun scaffolds of 50% polylactide acid and 50% polyglycolide acid were used to support the culture of rabbit corneal epithelial cells.
� A continuous layer of corneal cells formed within 2 weeks, as assessed by cryosectioning, confocal microscopy and scanning-electron microscopy.
� Scaffolds could be readily handled and sutured within the first week of culture with cells. � These synthetic off-the-shelf scaffolds could be a lower risk replacement for the amniotic membrane in delivering limbal epithelial cells to
treat corneal diseases.
Using PLGA electrospun scaffolds to deliver epithelial cells to the cornea ReseaRch aRticle
www.futuremedicine.com 401future science group
Bibliography1 Rama P, Bonini S, Lambiase A et al.:
Autologous fibrin-cultured limbal stem cells permanently restore the corneal surface of patients with total limbal stem cell deficiency. Transplantation 72(9), 1478–1485 (2001).
2 Daniels JT, Harris A, Mason C: Corneal epithelial stem cells in health and disease. Stem Cell Rev. 2(3), 247–254 (2006).
3 Dua HS, Azuara-Blanco A: Autologous limbal transplantation in patients with unilateral corneal stem cell deficiency. Br. J. Ophthalmol 84(3), 273–278 (2000).
4 Pellegrini G, Traverso CE, Franzi AT, Zingirian M, Cancedda R, De Luca M: Long-term restoration of damaged corneal surfaces with autologous cultivated corneal epithelium. Lancet 349(9057), 990–993 (1997).
5 Gomes JAP, dos Santos MS, Cunha MC, Mascaro VLD, Barros JN, de Sousa LB: Amniotic membrane transplantation for partial and total limbal stem cell deficiency secondary to chemical burn. Ophthalmol. 110(3), 466–473 (2003).
6 Tseng SCG, Prabhasawat P, Barton K, Gray T, Meller D: Amniotic membrane transplantation with or without limbal allografts for corneal surface reconstruction in patients with limbal stem cell deficiency. Arch. Ophthalmol. 116(4), 431–441 (1998).
7 Dua H, Gomes J, King A, Maharajan V: The amniotic membrane in ophthalmology. Surv. Ophthalmol. 49(1), 51–77 (2004).
8 Akle C, Welsh K, Adinolfi M, Leibowitz S, Mccoll I: Immunogenicity of human amniotic epithelial cells after transplantation into volunteers. Lancet 2(8254), 1003–1005 (1981).
9 Wang M, Yoshida A, Kawashima H, Ishizaki M, Takahashi H, Hori J: Immunogenicity and antigenicity of allogeneic amniotic epithelial transplants grafted to the cornea, conjunctiva, and anterior chamber. Invest. Ophthalmol. Vis. Sci. 47(4), 1522–1532 (2006).
10 Gabler B, Lohmann C: Hypopyon after repeated transplantation of human amniotic membrane onto the corneal surface. Ophthalmol. 107(7), 1344–1346 (2000).
11 Adds PJ, Hunt CJ, Dart JKG: Amniotic membrane grafts, “fresh” or frozen? A clinical and in vitro comparison. Br. J. Ophthalmol. 85(8), 905–907 (2001).
12 Otero J, Fresno MF, Escudero D, Seco M, González M, Peces R: Detection of occult disease in tissue donors by routine autopsy. Transplant Int. 11(2), 152–154 (1998).
13 Hopkinson A, McIntosh RS, Tighe PJ, James DK, Dua HS: Amniotic membrane for ocular surface reconstruction: donor variations and the effect of handling on TGF-b content. Invest. Ophthalmol. Vis. Sci. 47(10), 4316–4322 (2006).
14 Rahman I, Said DG, Maharajan VS, Dua HS: Amniotic membrane in ophthalmology: indications and limitations. Eye 23(10), 1954–1961 (2009).
15 Ahmed TAE, Dare EV, Hincke M: Fibrin: a versatile scaffold for tissue engineering applications. Tissue Eng. Part B Rev. 14(2), 199–215 (2008).
16 Blackwood KA, McKean R, Canton I et al.: Development of biodegradable electrospun scaffolds for dermal replacement. Biomaterials 29(21), 3091–3104 (2008).
18 Pham QP, Sharma U, Mikos AG: Electrospinning of polymeric nanofibers for tissue engineering applications: a review. Tissue Eng. 12(5), 1197–1211 (2006).
19 Srouji S, Kizhner T, Suss-Tobi E, Livne E, Zussman E: 3-D nanofibrous electrospun multilayered construct is an alternative ECM mimicking scaffold. J. Mater. Sci. Mater. Med. 19(3), 1249–1255 (2008).
20 Deshpande P, Notara M, Bullett N, Daniels JT, Haddow DB, MacNeil S: Development of a surface-modified contact lens for the transfer of cultured limbal epithelial cells to the cornea for ocular surface diseases. Tissue Eng. Part A 15(10), 2889–2902 (2009).
21 Rheinwald JG, Green H: Serial cultivation of strains of human epidermal keratinocytes: the formation of keratinizing colonies from single cells. Cell 6(3), 331 (1975).
22 Huang Q, Dawson R, Pegg D, Kearney J, MacNeil S: Use of peracetic acid to sterilize human donor skin for production of acellular dermal matrices for clinical use. Wound Repair Regen. 12(3), 276–287 (2004).
23 Zong X, Ran S, Kim KS, Fang D, Hsiao BS, Chu B: Structure and morphology changes during in vitro degradation of electrospun poly(glycolide-co-lactide) nanofiber membrane. Biomacromol. 4(2), 416–423 (2003).
24 Sangwan V, Matalia H, Vemuganti G et al.: Clinical outcome of autologous cultivated limbal epithelium transplantation. Indian J. Ophthalmol. 54(1), 29–34 (2006).
25 Sangwan VS, Matalia HP, Vemuganti GK et al.: Early results of penetrating keratoplasty after cultivated limbal epithelium transplantation. Arch. Ophthalmol. 123(3), 334–340 (2005).
26 Sangwan VS, Vemuganti GK, Singh S, Balasubramanian D: Successful reconstruction of damaged ocular outer surface in humans using limbal and conjuctival stem cell culture methods. Biosci. Reports 23(4), 169–174 (2003).
27 Grueterich M, Espana EM, Tseng SCG: Ex vivo expansion of limbal epithelial stem cells: amniotic membrane serving as a stem cell niche. Surv. Ophthalmol. 48(6), 631–646 (2003).
28 Dravida S, Gaddipati S, Griffith M et al.: A biomimetic scaffold for culturing limbal stem cells: a promising alternative for clinical transplantation. J. Tissue Eng. Regen. Med. 2(5), 263–271 (2008).
29 Sudha B, Madhavan H, Sitalakshmi G et al.: Cultivation of human corneal limbal stem cells in mebiol gel® – a thermo-reversible gelation polymer. Indian J. Med. Res. 124, 655–664 (2006).
30 Pino CJ, Haselton FR, Chang MS: Seeding of corneal wounds by epithelial cell transfer from micropatterned pdms contact lenses. Cell Transplant. 14, 565–571 (2005).
31 Nishida K, Yamato M, Hayashida Y et al.: Corneal reconstruction with tissue-engineered cell sheets composed of autologous oral mucosal epithelium. N. Engl. J. Med. 351(12), 1187–1196 (2004).
32 Loo S, Ooi CP, Wee SH, Boey YC: Effect of isothermal annealing on the hydrolytic degradation rate of poly(lactide-co-glycolide) (PLGA). Biomaterials 26(16), 2827–2833 (2005).
33 Horbett TA: The role of adsorbed proteins in animal cell adhesion. Colloid Surf. B Biointer. 2(1–3), 225–240 (1994).
34 Evans MDM, Steele JG: Multiple attachment mechanisms of corneal epithelial cells to a polymer – cells can attach in the absence of exogenous adhesion proteins through a mechanism that requires microtubules. Exp. Cell Res. 233(1), 88–98 (1997).
35 Hu X, Lui W, Cui L, Wang M, Cao Y: Tissue engineering of nearly transparent corneal stroma. Tissue Eng. 11(11–12), 1710–1717 (2005).
36 Zhang YQ, Zhang WJ, Liu W et al.: Tissue engineering of corneal stromal layer with dermal fibroblasts: Phenotypic and functional switch of differentiated cells in cornea. Tissue Eng. Part A 14(2), 295–303 (2008).
37 Selim M, Bullock A, Blackwood K, Chapple C, MacNeil S: Developing biodegradable scaffolds for tissue engineering of the urethra. Br. J. Urol. (2010) (In Press).
