A novel serum-free method for culturing human prenatal retinal pigment epithelial cells.
ABSTRACT Established techniques for culturing primary human retinal pigment epithelial (RPE) cells have facilitated the laboratory investigation of this multipurpose retinal cell layer. However, most culture methods involve the use of animal serum to establish and maintain RPE monolayers, which can complicate efforts to define and study factors involved in the maturation and function of these cells. Therefore, this study was conducted to develop a simple, serum-free system to propagate and sustain human RPE in vitro.
RPE was dissected from human prenatal donor eyes and cultured in serum-free defined medium containing the commercially formulated supplement B27 or N2. Cultures were grown initially as adherent tissue sections or suspended spherical aggregates and later expanded and maintained as monolayers. PCR, Western blot analysis, and immunocytochemistry were used to monitor gene and protein expression in established cultures, followed by examination of secretory products in RPE conditioned medium by ELISA and mass spectrometric analysis.
In medium supplemented with B27, but not N2, RPE could be expanded up to 40,000-fold over six passages and maintained in culture for more than 1 year. In long-term cultures, typical cellular morphology and pigmentation were observed, along with expression of characteristic RPE markers. RPE monolayers also retained proper apical-basal orientation and secreted multiple factors implicated in the maintenance of photoreceptor health and the pathogenesis of age-related macular degeneration.
Monolayer cultures of human prenatal RPE can be grown and maintained long term in the total absence of serum and still retain the phenotype, gene and protein expression profile, and secretory capacity exhibited by mature RPE cells.
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ABSTRACT: The retinal pigment epithelium contains three major types of pigment granules; melanosomes, lipofuscin and melanolipofuscin. Melanosomes in the retinal pigment epithelium (RPE) are formed during embryogenesis and mature during early postnatal life while lipofuscin and melanolipofuscin granules accumulate as a function of age. The difficulty in studying the formation and consequences of melanosomes and lipofuscin granules in RPE cell culture is compounded by the fact that these pigment granules do not normally occur in established RPE cell lines and pigment granules are rapidly lost in adult human primary culture. This review will consider options available for overcoming these limitations and permitting the study of melanosomes and lipofuscin in cell culture and will briefly evaluate the advantages and disadvantages of the different protocols.Experimental Eye Research 09/2014; 126:61–67. · 3.02 Impact Factor
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ABSTRACT: Invadopodia and podosomes, collectively referred to as invadosomes, are F-actin-rich basal protrusions of cells that provide sites of attachment to and degradation of the extracellular matrix. Invadosomes promote the invasion of cells, ranging from metastatic cancer cells to immune cells, into tissue. Here, we show that neuronal growth cones form protrusions that share molecular, structural and functional characteristics of invadosomes. Growth cones from all neuron types and species examined, including a variety of human neurons, form invadosomes both in vitro and in vivo. Growth cone invadosomes contain dynamic F-actin and several actin regulatory proteins, as well as Tks5 and matrix metalloproteinases, which locally degrade the matrix. When viewed using three-dimensional super-resolution microscopy, F-actin foci often extended together with microtubules within orthogonal protrusions emanating from the growth cone central domain. Finally, inhibiting the function of Tks5 both reduced matrix degradation in vitro and disrupted motoneuron axons from exiting the spinal cord and extending into the periphery. Taken together, our results suggest that growth cones use invadosomes to target protease activity during axon guidance through tissues. © 2015. Published by The Company of Biologists Ltd.Development (Cambridge, England). 01/2015;
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ABSTRACT: The retinal pigment epithelium (RPE) comprises a monolayer of polarized pigmented epithelial cells that is strategically interposed between the neural retina and the fenestrated choroid capillaries. The RPE performs a variety of vectorial transport functions (water, ions, metabolites, nutrients and waste products) that regulate the composition of the subretinal space and support the functions of photoreceptors (PRs) and other cells in the neural retina. To this end, RPE cells display a polarized distribution of channels, transporters and receptors in their plasma membrane (PM) that is remarkably different from that found in conventional extra-ocular epithelia, e.g. intestine, kidney, and gall bladder. This characteristic PM protein polarity of RPE cells depends on the interplay of sorting signals in the RPE PM proteins and sorting mechanisms and biosynthetic/recycling trafficking routes in the RPE cell. Although considerable progress has been made in our understanding of the RPE trafficking machinery, most available data have been obtained from immortalized RPE cell lines that only partially maintain the RPE phenotype and by extrapolation of data obtained in the prototype Madin-Darby Canine Kidney (MDCK) cell line. The increasing availability of RPE cell cultures that more closely resemble the RPE in vivo together with the advent of advanced live imaging microscopy techniques provides a platform and an opportunity to rapidly expand our understanding of how polarized protein trafficking contributes to RPE PM polarity.Experimental Eye Research 09/2014; 126C:5-15. · 3.02 Impact Factor
A Novel Serum-Free Method for Culturing Human
Prenatal Retinal Pigment Epithelial Cells
David M. Gamm,1,2J. Nicholas Melvan,1Rebecca L. Shearer,1Isabel Pinilla,3
Grzegorz Sabat,4Clive N. Svendsen,1,5and Lynda S. Wright1
PURPOSE. Established techniques for culturing primary human
retinal pigment epithelial (RPE) cells have facilitated the labo-
ratory investigation of this multipurpose retinal cell layer. How-
ever, most culture methods involve the use of animal serum to
establish and maintain RPE monolayers, which can complicate
efforts to define and study factors involved in the maturation
and function of these cells. Therefore, this study was con-
ducted to develop a simple, serum-free system to propagate
and sustain human RPE in vitro.
METHODS. RPE was dissected from human prenatal donor eyes
and cultured in serum-free defined medium containing the
commercially formulated supplement B27 or N2. Cultures
were grown initially as adherent tissue sections or suspended
spherical aggregates and later expanded and maintained as
monolayers. PCR, Western blot analysis, and immunocyto-
chemistry were used to monitor gene and protein expression
in established cultures, followed by examination of secretory
products in RPE conditioned medium by ELISA and mass spec-
RESULTS. In medium supplemented with B27, but not N2, RPE
could be expanded up to 40,000-fold over six passages and
maintained in culture for more than 1 year. In long-term cul-
tures, typical cellular morphology and pigmentation were ob-
served, along with expression of characteristic RPE markers.
RPE monolayers also retained proper apical–basal orientation
and secreted multiple factors implicated in the maintenance of
photoreceptor health and the pathogenesis of age-related mac-
CONCLUSIONS. Monolayer cultures of human prenatal RPE can be
grown and maintained long term in the total absence of serum
and still retain the phenotype, gene and protein expression
profile, and secretory capacity exhibited by mature RPE cells.
(Invest Ophthalmol Vis Sci. 2008;49:788–799) DOI:10.1167/
organization and function that is critical for the preservation of
outer retinal health and activity.1In addition, mounting evi-
dence points to an important role for the RPE in the develop-
ment of the neurosensory retina.2,3The study of the RPE has
been aided by advancements in culture techniques that encour-
age the adoption of characteristic RPE properties found in vivo.
Such properties include, among others, formation of a pig-
mented, compact monolayer of polygonal cells connected by
tight junctions, expression of specialized proteins, preserva-
tion of cellular orientation, and an ability to secrete multiple
factors.4–11However, the demonstration of some or all of these
features in vitro has necessitated the use of animal serum
and/or intricate combinations of chemical and protein constit-
uents not available in prefabricated media supplements.
The presence of serum in cell culture medium can be
problematic, because it contains numerous partially character-
ized or undefined factors that vary in concentration between
commercial preparations.7,12–15The existence of a defined,
serum-free growth and maintenance medium would be of
particular value for the straightforward identification of factors
released by the RPE in vitro. It would also provide a simplified,
reproducible environment for studying the maturation and
physiology of the RPE layer and its response to pharmacologic
treatments. Last, it would facilitate the use of cultured human
RPE in clinical studies by reducing exposure of cells to animal
products before transplant. Some protocols allow for the re-
duction or removal of serum from RPE medium preparations
after initial attachment or once the cells reach confluence.16–19
However, this practice may alter cell survival and function and
does not eliminate the exposure of proliferating RPE cell pop-
ulations to serum.14,20To address this problem, Tezel and Del
Priore12developed a chemically defined, serum-free medium
that supported the initial attachment and growth to confluence
of adult human RPE cells. However, production of the custom-
ized serum-free medium required the separate addition of nu-
merous components and did not use commercially formulated
supplements. Furthermore, the cultured RPE cells underwent
limited characterization in vitro. Thus, the impact of the serum-
free environment on the expression of critical RPE proteins
Sources of human RPE for experimentation and clinical use
have grown to include primary prenatal6,8,17,21and postna-
tal7,12,22,23tissue, transformed cell lines,24–27and embryonic
stem (ES) cells.28These sources differ widely in their expan-
sion potential, degree of differentiation, and propensity to
display RPE-like characteristics. For example, the commonly
used ARPE-19 cell line can be passaged indefinitely but does
not remain pigmented or reliably express certain RPE-selective
proteins, such as bestrophin.9,27,28Postnatal human RPE, on
the other hand, has limited expansion potential, but retains a
relatively mature phenotype in culture.7,12Human ES cells are
theoretically in limitless supply and can produce cells with a
he homogenous, relatively nondescript appearance of RPE
on the light microscopic level belies a complex cellular
From the1Waisman Center, and the Departments of2Ophthalmol-
ogy and Visual Sciences,
Biotechnology Center, University of Wisconsin School of Medicine and
Public Health, Madison, Wisconsin; and the3Department of Ophthal-
mology, Miguel Servet University Hospital and Aragones de Ciencias de
la Salud, Zaragoza, Spain.
Supported by National Eye Institute Grant K08EY015138, the
Walsh Foundation, Lincy Foundation, the Foundation Fighting Blind-
ness, the Retina Research Foundation, the Kinetics Foundation, the
U.S. Department of Defense (CNS), and Instituto Aragones de Ciencias
de la Salud (IP). DMG is a recipient of a Research To Prevent Blindness
Robert E. McCormick Scholar Award.
Submitted for publication June 23, 2007; revised October 2 and
30, 2007; accepted December 14, 2007.
Disclosure: D.M. Gamm, None; J.N. Melvan, None; R.L.
Shearer, None; I. Pinilla, None; G. Sabat, None; C.N. Svendsen,
None; L.S. Wright, None
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be marked “advertise-
ment” in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Corresponding author: David M. Gamm, Department of Ophthal-
mology and Visual Sciences, University of Wisconsin School of Medi-
cine and Public Health, T607 Waisman Center, 1500 Highland Avenue,
Madison, WI 53705; firstname.lastname@example.org.
5Anatomy and Neurology, and
Investigative Ophthalmology & Visual Science, February 2008, Vol. 49, No. 2
Copyright © Association for Research in Vision and Ophthalmology
gene and protein expression pattern and appearance similar to
primary RPE cultures with little contamination from other cell
types.28However, the functional competence of ES cell-de-
rived RPE is yet to be established29and, similar to other
sources of RPE, they are grown in the presence of serum.
Cultures of primary human prenatal RPE have the benefit of
possessing a greater growth capacity than adult RPE while still
exhibiting many of the known features of mature RPE in
vitro.6,17There are also disadvantages to the culture of human
prenatal RPE, including supply limitations and ethical concerns
regarding its use.
With these issues in mind, we sought to develop a simple,
serum-free method for culturing human prenatal RPE by using
commercially formulated medium supplements. B27, a supple-
ment that contains numerous factors of potential importance
for RPE growth and maturation,6,8,12,18,30supported the adher-
ence, propagation, and passaging of RPE on laminin-coated
tissue culture plastic. In contrast, RPE cultures exhibited min-
imal expansion in the presence of N2 supplement, which
includes only a fraction of the components present in B27.30,31
Primary attachment and outgrowth of RPE cells was improved
by implementing a method in which explants were cultured
initially as suspended spheroids. Resulting pigmented sphe-
roids consistently adhered to laminin-coated tissue culture plas-
tic and produced high yields of RPE cells. The morphology,
gene and protein expression profile, polarity, and secretory
capacity of cells cultured in this manner were similar to that
reported for RPE grown in the presence of serum.6,11
Serum-Free RPE Monolayer Culture
RPE was isolated from human eyes between 10 and 16 weeks of
gestation according to protocols approved by the NIH, the Institutional
Review Board at the University of Wisconsin-Madison, and the Univer-
sity of Washington. Eyes were shipped from the University of Wash-
ington Birth Defects Laboratory overnight at 4°C in sterile-filtered
transport medium (30 mM KCl, 5 mM glucose, 0.24 mM MgCl2, 1.95
mM NaH2PO4? 6H2O, 20 mM Na2HPO4? 2H2O, and 20 mM lactic acid,
followed by adjustment to pH 7.2 with KOH pellets and 300 mOsM
with approximately 140 millimoles sorbitol; all chemicals obtained
from Sigma-Aldrich, St. Louis, MO) and dissected on arrival. Whole eyes
were maintained in ice-cold dissection medium (DM; 70% DMEM
containing 4.5 g/L D-glucose (catalog no. 11965; Invitrogen, Carlsbad,
CA), 30% F12 nutrient mixture containing L-glutamine (catalog no.
11765; Invitrogen), and 1% antibiotic–antimycotic solution (catalog no.
14240; Invitrogen), whereas the attached periocular tissues were care-
fully removed and discarded. The remaining intact globes were rinsed
twice in DM, and the anterior segment of the eye and the vitreous were
removed. The eye cup was then washed two times with DM, and the
detached neural retina was removed in its entirety with forceps. If the
neural retina had not spontaneously detached after removal of the
vitreous, a 1-mL syringe with a 25-gauge,5⁄8-in. needle was used to
irrigate with DM gently under the peripheral retina, to separate it fully
from the underlying RPE. After two more DM washes, the remaining
RPE and attached choroid were peeled from the underlying sclera with
forceps, being careful to avoid the ciliary margin region. At this point,
the RPE–choroid explants were treated in one of three ways.
Method 1: Isolated RPE Sheets. Some explants were incu-
bated in DM containing 2% Dispase (catalog no. 17105-041; Invitrogen)
for 30 minutes at 37°C and washed twice in DM before small sheets of
RPE were teased away from the choroid with forceps and irrigation.
The isolated RPE sheets were collected and further chopped into
200-?m sections with a McIlwain automated tissue chopper (Mickle
Laboratory Engineering Co., Ltd., Guildford, UK), as described previ-
ously for primary cortical and retinal tissue.32,33After the tissue sec-
tions were chopped, they were placed on laminin-coated (0.001%,
catalog no. L2020; Sigma-Aldrich) tissue culture plastic (flasks or wells)
in a minimal volume of serum-free RPE medium (SFRM), consisting of
DM supplemented with either 2% B27 (50? solution, catalog no.
17504; Invitrogen) or 1% or 2% N2 (100? solution, catalog no. 17502;
Invitrogen), designated SFRM-B27 or SFRM-N2, respectively.
Method 2: RPE–Choroid Explants. Alternatively, RPE–cho-
roid explants were immediately chopped into 200-?m sections with-
out exposing them to proteolytic enzymes or manipulating them fur-
ther with forceps. After examining the chopped explant sections
under a dissection microscope, only those containing prominent
sheets of attached RPE were subsequently placed onto laminin-coated
tissue culture plastic. As described earlier, chopped explants were
cultured exclusively in SFRM-B27 or SFRM-N2.
Method 3: Pigmented Spheroids. Based on observations of
cell cultures established from the previous two methods, a modifica-
tion of method 2 was devised in which chopped RPE–choroid explants
were initially placed in suspension culture in SFRM-B27. Nonpig-
mented and pigmented spherical tissue aggregates formed within
hours, with many partially pigmented spheroids becoming uniformly
pigmented over 2 to 4 weeks in culture. At various times, darkly
pigmented spheroids were removed from suspension culture by using
a glass Pasteur pipette and were plated onto laminin-coated tissue
culture plates in SFRM-B27 or SFRM-N2.
Regardless of the method used, all cultures were maintained at
37°C and 5% CO2, and 50% to 75% of the cell culture medium was
exchanged every 1 to 2 days. Within 24 to 48 hours of initial plating,
expanding monolayers of RPE cells were observed to emanate from
attached tissue sections or pigmented spheroids. With method 2,
occasional colonies also arose that contained dense collections of
discrete, multilayered choroidal fibroblasts. The contaminating colo-
nies were marked on the outside of the flask and manually removed
with a Pasteur pipette under a dissecting microscope. After approxi-
mately 4 to 7 days, the remaining tissue sections or spheroids were
easily detached with simple irrigation and collected, leaving the sur-
rounding monolayer colonies of RPE cells attached to the flasks. De-
tached pigmented spheroids could be replated on laminin-coated plas-
tic at least three times with no observable decrease in initial RPE
production, or stored in liquid nitrogen in serum-free, cell-freezing
medium (catalog no. C6295; Sigma-Aldrich).
After removal of the tissue sections or spheroids, the remaining
adherent RPE monolayer colonies were treated with cell-detachment
medium (Accutase, catalog no. SCR005; Chemicon, Temecula, CA) for
10 to 15 minutes at 37°C until all cells had rounded up and begun to
lift off the plastic. Dissociated RPE cells were harvested by gently
irrigating the remaining loosely attached cells to lift them off the plastic
with a glass Pasteur pipette and transferring the cell suspension to a
15-mL conical tube. After a 3-minute centrifugation at 1000 rpm, the
cell detachment solution overlying the cell pellet was removed, and
the cells were resuspended in 10 mL of SFRM-B27 or SFRM-N2. The
cells were pelleted and resuspended a second time, counted on a
hemocytometer, and plated on laminin-coated tissue culture plastic at
a density of ?50,000 cells/cm2. When cultures reached confluence
(3–7 days after plating), they could be passaged until a growth plateau
was reached, and evidence of active cell division was no longer evident
by light microscopy (up to six passages). At each passage, a portion of
the cells were designated for long-term observation and study or for
serial passaging to establish growth curves. Excess RPE cells were
stored as aliquots in liquid nitrogen after dissociation and later re-
thawed to establish additional cultures as needed.
RPE Growth Determination
Growth curves were obtained by serially passaging RPE cultures within
24 hours of reaching confluence and determining the total number of
cells present per tissue culture flask or well using a hemocytometer
and trypan blue dye exclusion. Initial cell counts at day 0 (i.e., passage
1) were obtained after dislodging and removing adherent primary
tissue sections or pigmented spheroids and harvesting the surrounding
IOVS, February 2008, Vol. 49, No. 2
Serum-Free Human RPE Culture 789
monolayer skirt of RPE cells for subsequent passage. All counts were
performed in triplicate, and the mean was used to establish the in-
crease (n-fold change) in cell quantity for each passage. Doubling times
were calculated from exponential growth curves plotted for each
culture (Prism ver. 3.02; GraphPad Software, San Diego, CA). Cumu-
lative cell counts at later passages were extrapolated to account for the
fraction of cells stored in liquid nitrogen.
Acutely dissociated RPE cells from three separate cultures were plated
onto poly-L-lysine (0.01%; Sigma-Aldrich) and laminin-coated glass cov-
erslips (50,000 cells/coverslip) and incubated in SFRM-B27, with or
without 20 ng/mL FGF2 (catalog no. GF003; Chemicon). After 2 days
in culture, bromodeoxyuridine (BrdU; Sigma-Aldrich) was added to the
media to a final concentration of 0.2 ?M. Twenty-four hours later, the
RPE cells were fixed for 10 minutes in ice-cold methanol, washed in
PBS, and prepared for immunocytochemistry.
RNA Isolation and cDNA Synthesis
Total RNA was isolated (RNeasy MiniKit; Qiagen, Valencia, CA) using
the company’s protocol, which included an optional DNase I treat-
ment step. Individual RNA samples were treated a second time with
amplification grade DNase I (Invitrogen) before cDNA synthesis, which
was performed with a kit (Superscript III First-Strand Synthesis System
for RT-PCR; Invitrogen). All reactions included control samples in
which reverse transcriptase enzyme was not added to the reaction.
PCR reactions were performed by combining master mix (Promega,
Madison, WI), 10 ?m each of the appropriate forward and reverse
primers (Table 1), and 1:40 diluted cDNA template. Samples were
initially denatured at 95°C for 5 minutes followed by 35 cycles of PCR
amplification (95°C for 15 seconds, 60°C for 30 seconds, 72°C for 1
minute) and a final extension at 72°C for 10 minutes. PCR products
were visualized on a 1.5% agarose gel containing 0.1% ethidium bro-
Quantitative PCR was performed on a thermocycler (Opticon 2; Bio-
Rad, Hercules, CA, with SYBR Green 2? master mix; Applied Biosys-
tems, Foster City, CA) and 300 picomoles of the following forward and
reverse primers in 20-?L reactions: ?-actin: (forward) 5?-GCGAGAA-
GATGACCCAGATC-3?, (reverse) 5?-CCAGTGGTACGGCCAGAGG-3?;
bestrophin: (forward) 5?-CAA GCT GCT ATA TGG CGA GTT-3?, (re-
verse) 5?-GCC AGC CTA TAA ATA AAG CGG AT-3?. Reverse transcrip-
tion reactions were diluted 1:40, and 4 ?L were used per reaction.
Samples from five individual cultures were initially denatured at 95°C
for 15 minutes followed by 40 cycles of PCR amplification (95°C for 20
seconds, 55°C for 30 seconds, 72°C for 30 seconds) with a plate
reading after every extension. Melting curves were determined to
confirm amplification of the expected fragment. In addition, standard
curves were measured for all primer sets by serial dilution of reverse
transcriptase reactions, and efficiencies were calculated to ensure that
all primer sets had similar efficiencies of amplification. Each reaction
was performed in triplicate and CTvalues were obtained by averaging
the results. The relative differences (n-fold) in gene expression of a
characteristic marker of mature RPE, bestrophin, was determined
across cultures by using ?-actin as an internal control for each reaction
set (to determine ?CTvalues) and arbitrarily designating one culture as
a reference (to determine ??CTvalues). Data are presented as the
mean relative change ? SEM of three separate qPCR amplification
reactions for each culture.
Cell Lysate Preparation
RPE cells were solubilized in modified RIPA buffer (50 mM Tris-HCl
[pH 7.4], 150 mM NaCl, 2 mM EDTA, 0.1% SDS, 0.7% Triton X-100,
0.3% Nonidet P-40, 1% deoxycholate, 1 mM PMSF, 2 mM NaVO3) and
1:100 protease inhibitor cocktail for use with mammalian cell and
tissue extracts (catalog no. P8340; Sigma-Aldrich) containing 104 mM
4-(2-aminoethyl)benzenesulfonyl fluoride, 80 ?M aprotinin, 2 mM leu-
peptin, 4 mM bestatin, 1.5 mM pepstatin A, and 1.4 mM E-64. Lysates
were sonicated and total protein was quantified (DC Protein Assay;
TABLE 1. Primers Used for RT-PCR
AmplifiedForward ReverseSize (bp)
GCC CTC CTG CAC
AAG TTT GAC TTT
ATT TAT AGG CTG
GCC CTC ACG GAA
TTC CGC ATG GTA
CCT GAA GAG GAA
AGC CTG AGA GCA
TGA ATG TCA CCA
AAA GAG TCT CCA
TGC CCT ATG CCT
AAT CCA TCA TTC
ACC GGG CTC TCT
AGC GTC CGG ACC
CAA TAA CAG TTT
TGA GCT GGT GGA
TGC TCT TCA GTT
AGG GCA GAA TCA
TCA CGA AGT GGT
ATG AGC AAT TGG
TGG TGG ATG CTG
AAG AGC GAC CCT
CAC ATC AAG CTA
TGC TAA TGG TGG
AAA CCC ACA ACG
TCC ACT GAG CAA
AGC CGA ACT TCT
AGT TGG TCT CTG
TGC AAG CGT AGT
TGT TCT GCC GGA
GTC ATA AAG CCT
ACT GCA GCC GGA
AAT TCA CAT AGC
TGT TGA TCT GCA
CTC CCT TGG ACA
TTC ACA CCA CTG
TTC TCC ACT GCT
TGC ACC CAG TTG
TTG ATC TCT TGC
GCA GTG TCA AGG
GAA TGC TGA AGT
TAC TTC CTT CTG
GGT CTT GGG CAT
TCT GCA TGG TGA
TGT TGG ACT CCT
TTC CAG CCA CCT
CTG AAG ACC AAA
ATA CTG CCC AGT
TCG TTT CAG TGC
ATT TCT GGT ACA
GCT CCA CGT GCT
TTG CCT TGT CCG
TGG ACG TTT ACT
790Gamm et al.
IOVS, February 2008, Vol. 49, No. 2
RPE-Conditioned Medium Preparation
Tissue culture flasks containing confluent RPE monolayers were
washed three times with DM followed by a 24-hour incubation in DM.
Thereafter, RPE-conditioned medium (RPE CM) was collected, and
1:200 protease inhibitor cocktail for use in tissue culture medium
(catalog no. P1860; Sigma-Aldrich) was added. Conditioned medium
was then concentrated (up to 80-fold) using centrifugal filter devices
(Amicon; Millipore, Billerica, MA) with a 3-kDa cutoff according to the
manufacturer’s instructions. Total protein was then quantified (DC
Protein Assay; Bio-Rad).
Twenty micrograms of protein from RPE cell lysates or concentrated
RPE-conditioned medium was collected from RPE monolayers. Protein
samples were separated on 4% to 20% gradient Tris-Cl gels (Bio-Rad),
electroblotted onto PVDF membranes and stained with Ponceau red to
confirm transfer. Membranes were blocked with 5% nonfat dry milk
and 2.5% BSA in TBST for 1 hour at room temperature followed by
consecutive 1 hour incubations at room temperature with primary
antibody in TBST?1.5% BSA and HRP-conjugated secondary antibody
in TBST?1% nonfat dry milk. Primary antibodies used for Western blot
analysis were directed against RPE65 (mouse monoclonal, 1:2000;
Chemicon), bestrophin (mouse monoclonal, 1:500; Chemicon),
CRALBP (mouse monoclonal, 1:50,000, gift of John Saari, University of
Washington, Seattle), ezrin (rabbit polyclonal, 1:1000; Cell Signaling
Technologies, Danvers, MA), occludin (rabbit polyclonal, 1:250;
Zymed, Carlsbad, CA), claudin-10 (rabbit polyclonal, 1:250; Zymed),
PEDF (mouse monoclonal, 1:500; Chemicon), VEGF (rabbit polyclonal,
1:200, Santa Cruz Biotechnology, Santa Cruz, CA), and FGF2 (mouse
monoclonal, 1:500; Upstate Biotechnology, Charlottesville, VA). Pro-
tein bands were visualized by chemiluminescence (ECL or ECL Plus
Western Blot Analysis Detection Kit; GE Healthcare, Chalfont St.
Enzyme-Linked Immunosorbent Assay
During routine passaging of RPE cultures, 106cells were plated into
each laminin-coated well of a six-well plate in SFRM-B27. Cultures were
maintained with daily medium exchange for 4 to 6 weeks until cultures
displayed mature RPE cell morphology and pigmentation. Cultures
were then washed three times with DM followed by a 24-hour incu-
bation in SFRM-B27. Thereafter, RPE CM was collected and 1:200
protease inhibitor cocktail (catalog no. P1860; Sigma-Aldrich) was
added. Conditioned medium was then concentrated threefold with the
centrifugal filter devices (Amicon Centriplus; Millipore) with a 3-kDa
cutoff according to the manufacturer’s instructions. Levels of VEGF
and FGF2 protein were quantified in triplicate by ELISA (R&D Systems,
Minneapolis, MN) using the manufacturer’s protocols. PEDF levels
were similarly quantified by ELISA (Chemicon), with the exception
that the concentrated RPE CM was first treated with urea, per the
manufacturer’s protocol. The plated RPE cells from each well were
then dissociated and counted by using a hemocytometer to express
results as nanograms of growth factor produced per day per million
cells (mean ? SEM). A minimum of three assays were performed for
each growth factor.
During routine passaging of proliferating RPE cultures, 30,000 to
50,000 cells were plated in SFRM-B27 onto poly-L-lysine and laminin-
coated glass coverslips in 24-well plates. Cultures were maintained in
SFRM-B27 with daily medium exchange for 4 to 8 weeks until cultures
displayed mature RPE cell morphology and pigmentation. Cells were
fixed with 4% paraformaldehyde for 20 minutes and washed with
phosphate-buffered saline (PBS). Fixed cell cultures were permeabil-
ized for 10 minutes in 0.2% Triton X-100 in PBS, blocked with 5%
normal goat serum and 0.2% Triton X-100 in PBS for 30 minutes, and
processed for immunocytochemistry with primary antibodies to be-
strophin (1:100), CRALBP (1:2000), ezrin (1:100), occludin (1:100),
RPE65 (1:100), and ZO-1 (rabbit polyclonal, 1:100, Zymed). After they
were rinsed with PBS, the cells were incubated for 30 minutes with
secondary antibodies conjugated to Alexa 488 or Alexa 546 (1:1000;
Invitrogen-Molecular Probes, Eugene OR). Thereafter, Hoechst 33258
(1:10,000 in PBS) was added for 5 minutes to visualize nuclei, followed
by mounting in antifade reagent (Prolong Gold; Invitrogen-Molecular
Probes). Serial confocal image sections were collected with a laser
scanning fluorescence confocal microscope (model C1; Nikon Corp.,
Tokyo, Japan) and the resulting imaging data were used to reconstruct
cross-sectional z scans and rendered volume images (EZ-C1 software;
Nikon Corp.). For BrdU immunostaining, coverslips were exposed to
primary antibody directed against BrdU (rat monoclonal, 1:300; Accu-
rate Chemical, Westbury, NY), washed, and incubated with 1:2000
anti-rat secondary antibody conjugated to Cy3 (Jackson Laboratories,
West Grove, PA), followed by Hoechst 33258. To count BrdU-positive
cells, digital images of immunostained cells were taken using a high-
resolution camera (SPOT RT Slider; Diagnostic Instruments, Sterling
Heights, MI). Five to 10 random fields (?40 magnification) were
counted from at least three separate coverslips per experiment. The
percentage of BrdU incorporation was expressed as the mean ? SEM
for three separate experiments, and the results were analyzed using
Student’s two-tailed, unpaired t-test.
Two-Dimensional Gel Electrophoresis
Protein samples from RPE CM were concentrated to 919 ?g/mL and
mixed with five volumes of precooled (?20°C) precipitation solution
(10% trichloroacetic acid in acetone). Proteins were precipitated over-
night at ?20°C, pelleted by centrifugation for 15 minutes at 12,000g
(4°C), washed twice with ice-cold acetone, pelleted again, and washed
once more with 50% ice-cold acetone in water to remove any residual
salt contamination. The pellet was dried to remove excess acetone but
was prevented from completely drying, to facilitate subsequent resolu-
bilization. The sample was reconstituted in isoelectric focusing (IEF)
buffer (8 M urea, 4% CHAPS, 40 mM Tris-HCl [pH 9.0], 100 mM
dithiothreitol [DTT], and 0.2% ampholyte [Bio-Lyte 3/10; Bio-Rad],
with trace bromophenol blue).
For isoelectric focusing, 17 cm, pH 3 to 10 IPG strips (ReadyStrip;
Bio-Rad) were rehydrated with 500 and 1200 ?g of protein sample
overnight at room temperature. The isoelectric focusing was per-
formed in an IEF cell (Protean; Bio-Rad) at 20°C with a current limit of
30 ?A/strip for a total of 55,000 V/h (1-hour equilibration at 250 V,
3-hour slow ramp at 3000 V, 4-hour slow ramp at 10,000 V, and final
fast ramp at 10,000 V for 45 V/h). Focused IPG strips were stored at
?80°C until second-dimension analysis.
Before second-dimension analysis, the strips were equilibrated for
30 minutes in SDS equilibration solution (6 M Urea, 2% SDS, 30%
glycerol, 50 mM Tris-HCl [pH 8.8], with trace bromophenol blue)
containing 1% wt/vol DTT, then transferred to SDS equilibration solu-
tion containing 2.5% IEF and incubated for an additional 30 minutes.
The second dimension was performed overnight (Protean II xi system;
Bio-Rad) with 1-mm-thick, precast, 8% to 16% gradient SDS polyacryl-
amide gels (Bio-Rad) as follows: 1-hour stacking at 25 V (start: 8 mA,
end: 7 mA), 1-hour stacking at 50 V (start: 16 mA, end: 15 mA), 2-hour
resolving at 100 V (start: 31 mA, end: 25 mA), 16-hour resolving at 80
V (start: 19 mA, end: 10 mA), 0.5-hour resolving at 300 V (start: 40 mA,
end: 38 mA).
After electrophoresis, the gels were rinsed in water, fixed for 3
hours in MeOH/H2O/CH3COOH (50%:45%:5%) and stained overnight
with colloidal Coomassie blue (SimplyBlue SafeStain; Invitrogen). After
they were stained, the gels were washed in water three times for 30
minutes to remove any background staining, and the image was cap-
tured on a GS-800 calibrated densitometer (Bio-Rad). Spots of interest
were manually excised with a scalpel and transferred to siliconized
tubes for subsequent proteolytic digestion and mass spectrometric
IOVS, February 2008, Vol. 49, No. 2
Serum-Free Human RPE Culture791
Enzymatic Digestion and Mass
In gel digestion and mass spectrometric analysis were performed at the
Mass Spectrometry Facility (Biotechnology Center, University of Wis-
consin-Madison). Coomassie G-250 (colloidal)–stained gel pieces were
destained completely in 1:1 methanol:H2O containing 100 mM
NH4HCO3, dehydrated once for 10 minutes in 1:1 acetonitrile (ACN):
H2O with 25 mM NH4HCO3and again for 1 minute in 100% ACN, and
dried for 5 minutes (Speed-Vac). The dried gel pieces were rehydrated
and reduced in 25 mM DTT (in 25 mM NH4HCO3buffer) for 30
minutes at 56°C, alkylated with 55 mM iodoacetamide (in 25 mM
NH4HCO3buffer) in darkness at room temperature for 30 minutes,
washed twice in H2O for 1 minute each, equilibrated in 25 mM
NH4HCO3for 1 minute, dehydrated once for 10 minutes in 1:1 ACN:
H2O containing 25 mM NH4HCO3and again for 1 minute in 100% ACN,
dried, and rehydrated with 20 ?L of 20 ng/?L trypsin solution (Se-
quence Grade Modified; Promega) containing 25 mM NH4HCO3. The
digestion was conducted overnight (18 hours) at 37°C and subse-
quently terminated by acidification with an equal volume of 2.5%
trifluoroacetic acid (TFA). Peptides generated from the digestion were
extracted with an equal volume of 0.1% TFA and vigorous vortexing for
15 minutes, followed by the addition of an identical volume of 70:25:5
ACN:H2O:TFA and repeated vortexing. The peptide solution was col-
lected and dried completely in a Speed-Vac, resuspended in 50 ?L of
0.1% TFA, and solid phase extracted (ZipTip C18 pipette tips; Milli-
pore). Peptides were eluted off the C18 column with 60:40:0.2 ACN:
H2O:TFA directly onto 384-well plates (Opti-TOFApplied Biosystems)
and recrystalized with 0.75 ?L of matrix (10 mg/mL ?-cyano-4-hydroxy-
cinnamic acid in 50:35:0.1 acetone/CAN/TFA). Peptide map finger-
print result-dependent MS/MS analysis was then performed (4800 Ma-
trix-Assisted Laser Desorption/Ionization-Time of Flight-Time of Flight
[MALDI/TOF-TOF] mass spectrometer; Applied Biosystems). Peptide
fingerprints were generated by scanning a 700- to 4000-Da mass range
using 1000 shots acquired from 20 randomized regions of the sample
spot at 3600 intensity of an on-axis laser in positive reflectron mode
(OptiBeam; Applied Biosystems). The 10 most abundant precursors
(excluding trypsin autolysis peptides and sodium/potassium adducts)
were selected for subsequent tandem MS analysis, during which 2000
total shots were taken with 4200 laser intensity. Postsource decay
(PSD) fragments from the precursors of interest were isolated by
timed-ion selection and reaccelerated into the reflectron to generate
the MS/MS spectrum. Raw data was deconvoluted (GPS Explorer soft-
ware; Applied Biosystems) and submitted for peptide mapping and
MS/MS ion search analysis against a nonredundant National Center for
Biotechnology Information (NCBI; Bethesda, MD) database with an
in-house licensed Mascot search engine (Matrix Science, London, UK).
Role of Serum-Free Medium Containing B27
Supplement in Primary Human RPE Culture
Expansion and Maintenance
Human prenatal RPE was dissected as isolated sheets (method
1) or RPE–choroid explants (method 2), chopped into 200-?m
sections, and placed in laminin-coated tissue culture plastic in
serum-free RPE medium (SFRM) containing DMEM with high
glucose, Ham’s F12, and either B27 (SFRM-B27) or N2 (SFRM-
N2) supplement. B27 supplement was chosen for this study
because it is commercially formulated and contains many con-
stituents present in customized RPE medium preparations
and/or deemed important for optimal RPE growth and mainte-
nance (Table 2).6,8,12,18,30N2 supplement, which contains a
limited subset of factors present in B27,30,31was used for
With either method, initial outgrowth of cells was observed
in the presence of SFRM-B27 or SFRM-N2, but on subsequent
passages, substantial culture expansion occurred with SFRM-
B27 only (Fig. 1). No cell expansion was seen with cultures
maintained in SFRM without B27 or N2 (data not shown). The
doubling time of these RPE cell cultures in SFRM-B27 over the
first three passages was 1.84 ? 0.30 days (n ? 4), and growth
could be maintained for a maximum of six passages over 3 to
4 weeks, producing an approximately 40,000-fold increase in
the number of cells. After passaging and plating at a density of
50,000 cells/cm2, the cells became confluent within 5 days,
adopted a characteristic RPE morphology, and became increas-
TABLE 2. Composition of B27, N2, and Customized Serum-Free
Tezel and Del
Bovine serum albumin
Vitamin A derivative
DL-?-Tocopherol (vitamin E)
Follicle stimulating hormone
Epidermal growth factor
Basic fibroblast growth factor
‡ Retinyl acetate.
§ all-trans Retinoic acid.
free medium containing either B27 (filled symbols) or N2 (open sym-
bols) supplement. Cultures were derived from isolated RPE sheets
(method 1; circles), RPE–choroid explants (method 2; squares) or first
(up triangles) or second (down triangles) platings of pigmented sphe-
roids (method 3). Data points reflect cumulative cell counts obtained
after each culture passage.
Growth potential of representative RPE cultures in serum-
792Gamm et al.
IOVS, February 2008, Vol. 49, No. 2
ingly compacted and pigmented over the ensuing 6 weeks
(Figs. 2A–C). RPE monolayers were difficult to dissociate at this
late stage and could not be expanded further.
Although similar growth characteristics were observed in
RPE cultures established using method 1 or 2, disadvantages
were also encountered with both approaches. In method 1,
chopped sections of isolated, primary RPE would frequently
fail to adhere to the laminin-coated plastic, while in method 2,
contaminating choroidal cell colonies (distinguished by their
fibroblast morphology and tendency to form multilayers10) or
mixed choroidal/RPE cell colonies were regularly seen after
initial plating (Supplementary Fig. S1A; supplementary figures
are online at http://www.iovs.org/cgi/content/full/49/2/788/
DC1). When present, these contaminating colonies required
manual removal from culture flasks before subsequent passag-
ing. Therefore, in an effort to promote primary tissue adher-
ence and minimize early choroidal cell infiltration, a modifica-
tion of method 2 was used. In this third method, dissected
RPE–choroid explants were first grown as suspended sphe-
roids in SFRM-B27 until most of them adopted a uniform,
darkly pigmented appearance (Fig. 2D). Often, light micros-
copy could clearly identify a continuous surface layer of RPE
on these spheroids (Supplementary Figure S1B). The pig-
mented spheroids readily adhered to laminin-coated tissue cul-
ture plastic and gave rise to rapidly enlarging monolayers of
RPE cells with rare choroidal cell contamination (Figs. 2E, 2F).
RPE cells generated in this manner could be dissociated, ex-
panded and used to establish long-term monolayer cultures in
T-12.5, -25, and -75 flasks (Figs. 2G, 2H; Supplementary Figs.
S1C, S1D). Furthermore, the doubling time (1.59 ? 0.14 days;
n ? 3) and morphology of cell cultures established from
pigmented spheroids (method 3) were essentially indistin-
guishable from those originally obtained from isolated RPE
sheets (method 1) or RPE–choroid explants (method 2). Pig-
mented spheroids recovered after initial plating could be
stored frozen or replated at least three times, to establish
additional RPE cultures as needed. RPE cultures generated from
reused spheroids retained similar growth characteristics (Fig.
1) and morphologic features (Supplementary Figs. S1E, S1F)
exhibited by spheroids after initial plating. Based on these
collective findings, we elected to use RPE cultures established
by method 3 for subsequent experiments, except where oth-
Enhancement of RPE Proliferation by the
Addition of FGF2
Previous reports suggested that the inclusion of FGF2, a mito-
gen not present in B27 supplement, could significantly im-
prove cell proliferation in serum-containing RPE cultures.17,34
This finding prompted Tezel and Del Priore12to include FGF2
in their serum-free RPE culture medium preparation (Table 2).
To determine whether the addition of FGF2 to SFRM-B27 could
enhance RPE cell division compared with SFRM-B27 alone,
24-hour BrdU incorporation was measured in the presence or
absence of FGF2 beginning 2 days after culture passage (a
period corresponding to peak cell proliferation). Cultures (n ?
3) maintained in SFRM-B27 with 20 ng/mL FGF2 demonstrated
37.8% ? 2.8% BrdU incorporation, compared with 19.6% ?
2.6% in parallel cultures grown without FGF2 (P ? 0.003; Fig.
3). Thus, although SFRM-B27 is capable of supporting substan-
tial human RPE growth in vitro, its proliferative effects can be
augmented with the addition of one or more defined factors.
Expression of RPE-Selective Genes and Proteins
in Serum-Free Culture
RT-PCR was used to examine the gene-expression profile of
RPE monolayers (n ? 3) grown and maintained exclusively in
SFRM-B27. Complementary DNA was generated from long-
term RPE monolayer cultures, all of which expressed the RPE-
croscopic images of cultured RPE es-
tablished from isolated sheets of pri-
mary tissue (method 1) on days 0 (A),
1 (B), and 28 (C) after third passage
in SFRM-B27. (D) A typical pig-
mented spheroid (method 3) derived
from sectioned RPE–choroid sheets
after 25 days in suspension culture in
SFRM-B27. (E) Low- and (F) high-
power images of the same spheroid
shown in (D) 6 days after attachment
to laminin-coated tissue culture plas-
tic. (G) Low- and (H) high-power im-
ages of a T-12.5 flask containing
third-passage RPE (method 3) 6
weeks after seeding. Scale bars: (B,
C, H) 20 ?m; (A, F) 50 ?m; (D, E)
Phase-contrast light mi-
IOVS, February 2008, Vol. 49, No. 2
Serum-Free Human RPE Culture793
specific gene RPE65, as well as bestrophin, cellular retinalde-
hyde-binding protein (CRALBP), and receptor tyrosine kinase
Mer (MerTK), whereas von Willebrand factor, an endothelial
cell marker, was absent (Fig. 4A). PCR results obtained from
cultures established by using method 1 or 2 showed the same
pattern of gene expression (data not shown). Quantitative PCR
analysis further demonstrated ?2-fold variations in bestrophin
gene expression levels across multiple cultures (n ? 5) of
different methods of origin, age in culture, passage number,
and spheroid plating number (Supplementary Fig. S2).
Western blot analysis performed on cell lysates obtained
from RPE monolayer cultures (n ? 3) revealed protein expres-
sion of RPE65, bestrophin, and CRALBP in all samples (Fig. 4B).
Other RPE proteins prominently expressed in vivo were also
present in cell lysates, including ezrin and the tight junction
proteins claudin-10 and occludin (Fig. 4B). Of note, occludin
was often found in a low-molecular-weight form, although
higher-molecular-weight forms indicative of protein phosphor-
ylation35–37were also present on these blots.
Apical–Basal Polarity and Tight Junction Protein
Expression in RPE Cells in Serum-Free Culture
RPE cells are polarized in vivo, with their apical surface di-
rected toward the outer segments of photoreceptors and their
basal surface facing the choriocapillaris.1,6,8,38This orientation
is necessary for the RPE layer to perform many of its functions
properly in vivo, including photopigment recycling, outer seg-
ment phagocytosis, ion and fluid transport, and directed factor
secretion.1To assess the polarity of cultured RPE cells grown
and maintained in SFRM-B27, we performed immunocyto-
chemistry and confocal microscopy using antibodies directed
against RPE proteins expressed in the apical cell membrane
(ezrin), in the basolateral membrane (bestrophin), or within
the cytoplasm (CRALBP and RPE65). As shown in Figures
5A–D, these proteins were predominantly localized to their
expected regions within the cultured RPE cells.
Another important structural feature of native RPE mono-
layers is the existence of tight junctions between adjacent
cells, which are key components of the outer blood–retina
barrier.38We investigated the expression and localization of
the tight junction proteins zonula occludens (ZO)-1 and occlu-
din, both of which demonstrated localized intercellular stain-
ing (Figs. 5E, 5F). A third junctional complex protein, claudin-
10, was not consistently detected in the intercellular space by
immunocytochemistry (data not shown). Together, these re-
sults reveal that SFRM-B27 can support the development and
maintenance of proper RPE cell orientation, as well as the
expression and localization of some, but probably not all, of
the tight junction proteins found in native RPE.
Multiple Protein Factors Secreted by RPE Cells in
RPE cells secrete numerous growth factors and other proteins
that impact photoreceptor health and function.1,4,6,11The ca-
pacity of serum-free RPE monolayer cultures to produce and
secrete specific growth factors was investigated by RT-PCR,
Western blot analysis, and ELISA. First, we examined gene
expression of a host of growth factor genes known to be
expressed in the RPE, including pigment epithelium–derived
growth factor (PEDF), nerve growth factor (NGF), insulin-like
growth factor 1 (IGF1), vascular endothelial growth factor
(VEGF), epidermal growth factor (EGF), FGF2, transforming
growth factor ?1 (TGFB1), and brain-derived growth factor
(BDNF; Fig. 6A). These genes were expressed in our standard
serum-free conditions in all cultures tested (n ? 3). To deter-
mine whether selected growth factor gene transcripts are sub-
sequently translated and released, CM was collected from these
SFRM-B27 without (hatched bar) or with (solid bar) 20 ng/mL FGF2.
Results are expressed as the mean ? SEM of results of four experi-
ments on three separate cultures (**P ? 0.005).
Percentage of BrdU incorporation into RPE cultured in
in serum-free cultures as determined by RT-PCR and Western blot
analysis (n ? 3), respectively. VWF was used as a marker for endothe-
lial cell contamination. Results were consistent across all cultures
tested, regardless of the method of origin (methods 1–3), time spent in
culture (2–12 months), or number of passages (2–5).
Expression of characteristic RPE genes (A) and proteins (B)
794Gamm et al.
IOVS, February 2008, Vol. 49, No. 2
RPE monolayers, concentrated, and analyzed by Western blot
(n ? 3). All growth factors examined (PEDF, VEGF, and FGF2)
were present in RPE CM (Fig. 6B). Quantification of PEDF,
VEGF, and FGF2 in RPE CM by ELISA (n ? 3) revealed secretion
rates of 2453 ? 1099 ng/106cells/d, 0.94 ? 0.05 ng/106
cells/d, and 0.054 ? 0.005 ng/106cells/d, respectively
In addition to growth factors, RPE cells release a host of
regulatory proteins with diverse functions into their surround-
ing microenvironment.11The defined, serum-free medium
used for this study allowed straightforward identification of
major proteins present in the human RPE secretome. Concen-
trated RPE CM was separated by 2-D gel electrophoresis, and
isolated protein spots were subjected to tandem mass spec-
trometry. We identified the 12 most abundant proteins in RPE
CM by this method, including proteins involved in signaling,
metal binding, and immune regulation, in addition to one
carry-over from B27 supplement (bovine serum albumin; Table
3). Thus, cultured RPE retains a strong capacity to secrete a
variety of factors without the need for serum supplementation.
Numerous culture methods and media preparations have been
described that support the initial attachment, limited growth,
and long-term maintenance of primary human RPE cells in
vitro. Early media formulations were relatively simple, primar-
ily using a base medium supplemented with a high percentage
of serum.22,23,39–43A drawback of using serum supplementa-
tion in culture media is the ambiguous introduction of signal-
ing molecules, including hormones and cytokines, and other
known and unknown factors with the potential to influence
cell behavior.12,13,44With the development of more complex
and precise RPE media preparations, less reliance was placed
on undefined additives.6,8,18,45Some of these tailored medium
recipes called for the presence of a low percentage (1%–5%) of
animal serum and/or tissue extract throughout the culture
period,6,8whereas in others, high levels of serum (15%–20%)
were present at initial seeding.6,18By contrast, Tezel and Del
Priore12developed a chemically defined culture medium that
supported attachment, proliferation, and three serial passages
of adult human RPE in the total absence of serum. However,
characterization of the cultured RPE cells was limited to mor-
phology and cytokeratin expression, and preparation of the
customized serum-free medium required assembly of 12 sepa-
rate components (including 7 growth factors and hormones) in
addition to the base medium.
Comparison of the formulas used in successful low or no
serum medium preparations for human RPE cultures reveals
certain commonalities. All include a base medium (typically
MEM or DMEM supplemented with amino acids and inorganic
salts), transferrin and insulin, and most contain corticosteroid,
triiodothyronine, selenium, and putrescine.6,8,12,18Many of
these factors are necessary for the growth and maintenance of
mammalian cells in general. Others, such as triiodothyronine
and corticosteroid, are potentially important for RPE cell me-
tabolism, fluid flux, and tight junction formation.46–48Addi-
tional factors used in various RPE-specific media include EGF,
FGF2, follicle stimulating hormone, retinoic acid, linoleic acid,
ascorbic acid, progesterone, and taurine. These factors have
been shown to affect RPE cell adhesion, proliferation, survival,
and morphology, along with other critical cellular func-
tions.12,17,18,49–51Most of these substances are present in B27
supplement, a highly augmented form of N2 initially designed
to support the maintenance of hippocampal neurons and neu-
ronal cell lines.52Subsequently, B27 was found to promote
expansion of embryonic and adult neural stem cells in serum-
free neurosphere cultures.30,53Given its composition and
proven efficacy in serum-free cultures of other neuroectoder-
mal cell types, we predicted that B27 would also support RPE
growth and long-term survival. However, the fact that B27
alone could eliminate the need for serum to establish and
expand RPE cultures was unanticipated, since certain mitogens
absent from B27 are often used to promote cell growth in other
serum-free media preparations.12,18,32,33Even so, mitogens can
be used to enhance the growth potential of human RPE in
SFRM-B27, as demonstrated by the increased cell proliferation
observed after addition of FGF2. The effects of FGF2 supple-
mentation on other aspects of RPE cell culture (e.g., differen-
tiation) are currently being investigated by using our serum-
In the present study, a single medium was used for all
phases of RPE culture. Other culture systems use different
formulations for the adherence, expansion, differentiation
and/or maintenance of RPE cells in vitro.6,8Phenotypic matu-
ration of RPE monolayers in SFRM-B27, on the other hand,
ing and confocal imaging of RPE cul-
tures (passage 2 or 3) grown exclu-
sively in SFRM-B27. Top: an en face
view of an RPE monolayer presented
as a maximum-intensity projection
through the z-axis. Bottom: cross-sec-
tion through the z-plane of multiple
optical slices at the location indicated
by the white reference line in the cor-
responding top portion. Ezrin (A),
CRALBP (B), RPE65 (C), bestrophin
(D), occludin (E), and ZO-1 (F). Scale
bar, 10 ?m.
IOVS, February 2008, Vol. 49, No. 2
Serum-Free Human RPE Culture 795
occurred spontaneously over time after the cells reached con-
fluence, as evidenced by replicative senescence, adoption of a
compact, polygonal morphology, and repigmentation.
The profound difference in growth potential between se-
rum-free RPE cultures treated with B27 versus N2 supplement
may prove useful in the identification of candidate factors
important for RPE proliferation. There are 15 B27 components
lacking in N2 supplement, five of which (triiodothyronine,
vitamin A derivative, corticosteroid, selenium, and linoleic
acid) were also incorporated in the serum-free RPE medium
developed by Tezel and del Priore.12The availability of de-
fined, serum-free RPE culture protocols should facilitate the
future investigation of the roles of these and other factors in
human RPE growth.
Although SFRM-B27 supported RPE growth from isolated
sheets of human RPE (method 1) or RPE–choroid explants
(method 2), there were technical limitations to each of these
approaches. Primary attachment was inconsistent with isolated
RPE sheets, and early choroidal contamination was seen in
explant cultures by light microscopy. However, we noted that
chopped RPE–choroid explants that did not immediately plate
down would persist as suspended spheroids in SFRM-B27 and
become uniformly pigmented over time. Conversely, nonad-
herent RPE sheets degraded in culture, consistent with previ-
ous reports.54,55Over time, the explant-derived, pigmented
spheroids spontaneously attached to the culture plastic surface
and cells of a nearly exclusive RPE morphology emanated from
them. We postulate that the relative lack of choroidal contam-
ination in these cultures is due to the development of a layer of
RPE cells on the spheroid surface, which may sequester cho-
roidal fibroblasts from substrate after plating. Electron micro-
scopic studies are currently under way to investigate this pos-
sibility in more depth. RPE monolayers established from the
pigmented spheroid method (method 3) could be passaged
and expanded in a manner identical with those established
from isolated RPE sheets or RPE–choroid explants. These find-
ings were similar to those of Aronson,21who first noted that
rounded RPE–choroid fragments could generate RPE cultures
after plating. Later, Rezai et al.54and Gabrielian et al.56showed
that human fetal RPE could be grown on free-floating spheroi-
dal polymer scaffolds in medium containing 15% FBS. After
attachment of the RPE-coated polymer spheres to culture plas-
tic, RPE cells proliferated from the site of adhesion and formed
monolayers. Therefore, culturing primary RPE initially as sus-
pended spheroids on a natural (choroid) or synthetic substrate
promotes subsequent cell attachment and proliferation in the
absence or presence of serum, respectively.
Once grown to confluence in SFRM-B27, monolayer RPE
cultures expressed markers indicative of mature RPE cells.
Furthermore, immunocytochemical analysis revealed that the
normal apical–basal polarity of RPE cells was preserved, along
with the expression of characteristic junctional complex pro-
teins. An exception was claudin-10, a tight junction protein
shown to be expressed discontinuously by human RPE cul-
tured in the presence of serum.6In the present study, clau-
din-10 was detected in RPE lysates by Western blot but could
not be unequivocally localized to the junctional complex by
immunocytochemistry. Occludin expression also did not ap-
pear to be restricted to apical junctions, although it was clearly
present along the lateral RPE membranes. A possible explana-
tion for these observations is that SFRM-B27 lacks the elements
necessary to assemble the complete complex. However, tight
junction formation in cultured RPE cells can also be inhibited
by serum,57suggesting that multiple unknown factors, both
positive and negative, can influence this process.58The exis-
tence of serum-free RPE culture protocols should facilitate the
search for these and other factors involved in the development
and regulation of RPE structure and function in vitro.
A defined, serum-free environment is also conducive, al-
though not essential, to the study of cellular secretion in vitro.
Recently, An et al.11examined the secretome profile of adult
human RPE cultured in the presence of serum by labeling
newly synthesized proteins with isotope-tagged amino acids.
The authors detected a wide range of proteins in spent RPE
medium, most of which are known to be secreted. In the
present study, numerous discrete protein spots were visible
after 2-D gel electrophoresis of concentrated RPE CM from
serum-free prenatal cultures. Mass spectrometric analysis of the
major protein spots subsequently led to the identification of 12
individual proteins. Six of these were also found in the study by
An et al.11: PEDF, galectin-3 binding protein, prostaglandin D2
synthase, complement subcomponents 1s and 1r, and ?-actin.
Of the remaining six prominent proteins, four are known to be
produced by RPE (cystatin C, transthyretin, cathepsin D, and
ceruloplasmin), whereas the ocular expression of another (al-
cadein ?-1) has not been documented. The final identified
constituent in our study, bovine serum albumin, represents a
free RPE cultures as determined by RT-PCR. (B) Western blot analysis
of conditioned medium from serum-free RPE cultures reveal the pres-
ence of secreted growth factors. (C) Quantification of PEDF, VEGF, and
FGF2 levels in serum-free RPE conditioned medium. Results were
consistent across all cultures tested (n ? 3), regardless of method of
origin (methods 1–3), time spent in culture (2–12 months), or the
number of passages (2–5).
(A) Expression of selected growth factor genes in serum-
796Gamm et al.
IOVS, February 2008, Vol. 49, No. 2
carry-over contaminant from B27 supplement. This finding is
consistent with an earlier report in which serum contaminants
were present in RPE CM despite multiple washings and incu-
bation with serum-free medium before sample collection.11
Thus, short-term removal of serum from RPE culture medium is
not sufficient to eliminate serum components in subsequent
Most of the proteins identified by our mass spectrometric
analysis are secreted and have the potential to influence retinal
development, homeostasis, and/or disease. PEDF is an inhibitor
of retinal angiogenesis59,60with neuroprotective and neurotro-
phic properties60that is secreted by RPE before and after
birth.61,62Galectin-3 binding protein is apically secreted and
promotes integrin-mediated cell adhesion and binds numerous
extracellular proteins, including fibronectin.63,64Prostaglandin
D2 synthase, an enzyme postulated to function as a retinoid
transporter,65is released into the interphotoreceptor matrix
where it is taken up by photoreceptors. The roles of comple-
ment proteins C1s and C1r are less clear, although other
complement proteins have been implicated in the pathogene-
sis of age-related macular degeneration (AMD).66,67Cystatin C
is a secreted cysteine protease inhibitor thought to be involved
in neurodegeneration, neuroprotection, and central nervous
system (CNS) repair.68,69It is highly expressed in fetal and
adult RPE and is postulated to regulate photoreceptor degra-
emia.68,70–72Cystatin C may indirectly regulate the activity of
cathepsin D, a major lysosomal protease responsible for the
ments.68,73Although the proenzyme form of cathepsin D (52
kDa) is secreted by RPE,74its precise role after release into the
extracellular environment is not known. Transthyretin is a
homotetrameric serum transport protein for thyroxine and
triiodothyronine that also interacts with retinol binding pro-
tein.75In the eye, it is known to be secreted in abundance by
RPE,75predominantly from the apical surface,76,77but it is also
found in AMD drusen deposits.78Ceruloplasmin serves as an
antioxidant, converting ferrous iron to the less dangerous fer-
ric form.79,80It has a secreted form that is present within the
retina and vitreous,79–81and the absence of ceruloplasmin has
been linked to free radical injury and degenerative diseases
within the brain and retina.79,82Alcadeins indirectly associate
with amyloid ?-protein precursor (APP)83,84and are thought to
function physiologically in concert with APP, a molecule im-
plicated in the pathogenesis of Alzheimer disease and AMD.85
Like APP, alcadein ?-1 is a transmembrane protein whose
extracellular domain is subject to cleavage and release into the
extracellular space. However, the present analysis discovered
only the unprocessed form of alcadein ?-1 in RPE CM.
andafter trauma or isch-
photoreceptor outer seg-
Although PEDF was the only growth factor identified as a
major secreted protein by 2-D gel electrophoresis, RPE is
known to secrete a host of other growth factors as well.1,4,60It
is likely that these other molecules are more labile than PEDF
or are secreted at levels too low to detect on Coomassie- or
silver-stained 2-D gels, even after concentration of RPE CM.
However, the potential for serum-free RPE cultures to produce
and secrete multiple growth factors, including VEGF and FGF2,
was confirmed in this study by PCR, Western blot, and ELISA.
Thus, human RPE cultured exclusively in SFRM-B27 retains its
ability to secrete a host of growth factors implicated in the
maintenance of retinal health and function.
Altogether, the results confirm that human RPE cultures can
be established, expanded,and maintained in the complete ab-
sence of serum. By taking advantage of the commercially for-
mulated B27 supplement, we simplified the preparation of
serum-free medium without adversely affecting RPE cell mor-
phology, protein expression, or secretory capacity. Additions
to this minimal serum-free medium formulation or alterations
in the culture technique may optimize the method further.
However, the current protocol should facilitate examination of
RPE development, structure, function, and disease in a defined,
reproducible culture environment.
The authors thank Elizabeth Capowski and Jason Meyer for helpful
comments during the preparation of the manuscript and Jolien Conner
for assistance with confocal microscopy.
1. Strauss O. The retinal pigment epithelium in visual function.
Physiol Rev. 2005;85:845–881.
2. Pearson RA, Dale N, Llaudet E, et al. ATP released via gap junction
hemichannels from the pigment epithelium regulates neural reti-
nal progenitor proliferation. Neuron. 2005;46:731–744.
3. Raymond SM, Jackson IJ. The retinal pigmented epithelium is
required for development and maintenance of the mouse neural
retina. Curr Biol. 1995;5:1286–1295.
4. Ishida K, Yoshimura N, Yoshida M, et al. Expression of neurotro-
phic factors in cultured human retinal pigment epithelial cells.
Curr Eye Res. 1997;16:96–101.
5. Kuriyama S, Ohuchi T, Yoshimura N, et al. Growth factor-induced
cytosolic calcium ion transients in cultured human retinal pigment
epithelial cells. Invest Ophthalmol Vis Sci. 1991;32:2882–2890.
6. Maminishkis A, Chen S, Jalickee S, et al. Confluent monolayers of
cultured human fetal retinal pigment epithelium exhibit morphol-
ogy and physiology of native tissue. Invest Ophthalmol Vis Sci.
TABLE 3. Proteins Identified by Mass Spectrometry in the Conditioned Medium of Serum-Free RPE Cultures
Prostaglandin D2 synthase
Pigment epithelium-derived factor
Galectin-3 binding protein
Albumin [Bos taurus]
Complement component 1, s subcomponent
Complement component 1, r subcomponent
* Mascot scores greater than 78 are significant (P ? 0.05).
IOVS, February 2008, Vol. 49, No. 2
Serum-Free Human RPE Culture 797
7. Engelmann K, Valtink M. RPE cell cultivation. Graefes Arch Clin
Exp Ophthalmol. 2004;242:65–67.
8. Hu J, Bok D. A cell culture medium that supports the differentia-
tion of human retinal pigment epithelium into functionally polar-
ized monolayers. Mol Vis. 2001;7:14–19.
9. Marmorstein AD, Marmorstein LY, Rayborn M, et al. Bestrophin,
the product of the Best vitelliform macular dystrophy gene
(VMD2), localizes to the basolateral plasma membrane of the
retinal pigment epithelium. Proc Natl Acad Sci USA. 2000;97:
10. McKay BS, Burke JM. Separation of phenotypically distinct sub-
populations of cultured human retinal pigment epithelial cells. Exp
Cell Res. 1994;213:85–92.
11. An E, Lu X, Flippin J, et al. Secreted proteome profiling in human
RPE cell cultures derived from donors with age related macular
degeneration and age matched healthy donors. J Proteome Res.
12. Tezel TH, Del Priore LV. Serum-free media for culturing and serial-
passaging of adult human retinal pigment epithelium. Exp Eye Res.
13. Shah G. Why do we still use serum in the production of biophar-
maceuticals? Dev Biol Stand. 1999;99:17–22.
14. Griffiths JB. Serum and growth factors in cell culture media: an
introductory review. Dev Biol Stand. 1987;66:155–160.
15. Goldberg AM. Mechanisms of neurotoxicity as studied in tissue
culture systems. Toxicology. 1980;17:201–208.
16. Arrindell EL, McKay BS, Jaffe GJ, et al. Modulation of potassium
transport in cultured retinal pigment epithelium and retinal glial
cells by serum and epidermal growth factor. Exp Cell Res. 1992;
17. Song MK, Lui GM. Propagation of fetal human RPE cells: preser-
vation of original culture morphology after serial passage. J Cell
18. Oka MS, Landers RA, Bridges CD. A serum-free defined medium for
retinal pigment epithelial cells. Exp Cell Res. 1984;154:537–547.
19. Gaur VP, Liu Y, Turner JE. RPE conditioned medium stimulates
photoreceptor cell survival, neurite outgrowth and differentiation
in vitro. Exp Eye Res. 1992;54:645–659.
20. de Pomerai DI, Gali MA. Influence of serum factors on the preva-
lence of “normal” and “foreign” differentiation pathways in cul-
tures of chick embryo neuroretinal cells. J Embryol Exp Morphol.
21. Aronson JF. Human retinal pigment cell culture. In Vitro. 1983;
22. Ishida M, Lui GM, Yamani A, et al. Culture of human retinal
pigment epithelial cells from peripheral scleral flap biopsies. Curr
Eye Res. 1998;17:392–402.
23. Edwards RB. Culture of mammalian retinal pigment epithelium
and neural retina. Methods Enzymol. 1982;81:39–43.
24. Kanuga N, Winton HL, Beauchene L, et al. Characterization of
genetically modified human retinal pigment epithelial cells devel-
oped for in vitro and transplantation studies. Invest Ophthalmol
Vis Sci. 2002;43:546–555.
25. Davis AA, Bernstein PS, Bok D, et al. A human retinal pigment
epithelial cell line that retains epithelial characteristics after pro-
longed culture. Invest Ophthalmol Vis Sci. 1995;36:955–964.
26. Rambhatla L, Chiu CP, Glickman RD, et al. In vitro differentiation
capacity of telomerase immortalized human RPE cells. Invest Oph-
thalmol Vis Sci. 2002;43:1622–1630.
27. Dunn KC, Aotaki-Keen AE, Putkey FR, et al. ARPE-19, a human
retinal pigment epithelial cell line with differentiated properties.
Exp Eye Res. 1996;62:155–169.
28. Klimanskaya I, Hipp J, Rezai KA, et al. Derivation and comparative
assessment of retinal pigment epithelium from human embryonic
stem cells using transcriptomics. Cloning Stem Cells. 2004;6:217–
29. Lund RD, Wang S, Klimanskaya I, et al. Human embryonic stem
cell-derived cells rescue visual function in dystrophic RCS rats.
Cloning Stem Cells. 2006;8:189–199.
30. Wachs FP, Couillard-Despres S, Engelhardt M, et al. High efficacy
of clonal growth and expansion of adult neural stem cells. Lab
31. Bottenstein JE, Sato GH. Growth of a rat neuroblastoma cell line in
serum-free supplemented medium. Proc Natl Acad Sci USA. 1979;
32. Svendsen CN, ter Borg MG, Armstrong RJ, et al. A new method for
the rapid and long term growth of human neural precursor cells.
J Neurosci Methods. 1998;85:141–152.
33. Gamm DM, Nelson AD, Svendsen CN. Human retinal progenitor
cells grown as neurospheres demonstrate time-dependent changes
in neuronal and glial cell fate potential. Ann NY Acad Sci. 2005;
34. Schwegler JS, Knorz MC, Akkoyun I, et al. Basic, not acidic fibro-
blast growth factor stimulates proliferation of cultured human
retinal pigment epithelial cells. Mol Vis. 1997;3:10.
35. Peng S, Rahner C, Rizzolo LJ. Apical and basal regulation of the
permeability of the retinal pigment epithelium. Invest Ophthalmol
Vis Sci. 2003;44:808–817.
36. Hirase T, Staddon JM, Saitou M, et al. Occludin as a possible
determinant of tight junction permeability in endothelial cells.
J Cell Sci. 1997;110:1603–1613.
37. Williams CD, Rizzolo LJ. Remodeling of junctional complexes
during the development of the outer blood-retinal barrier. Anat
38. Rizzolo LJ. Development and role of tight junctions in the retinal
pigment epithelium. Int Rev Cytol. 2007;258:195–234.
39. Boulton ME, Marshall J, Mellerio J. Human retinal pigment epithe-
lial cells in tissue culture: a means of studying inherited retinal
diseases. Birth Defects Orig Artic Ser. 1982;18:101–118.
40. Wong HC, Boulton M, McLeod D, et al. Retinal pigment epithelial
cells in culture produce retinal vascular mitogens. Arch Ophthal-
41. Turksen K, Opas M, Kalnins VI. Cytoskeleton, adhesion, and ex-
tracellular matrix of fetal human retinal pigmented epithelial cells
in culture. Ophthalmic Res. 1989;21:56–66.
42. Zheng J, Guo Y, Jing X, et al. Modification of isolation and culture
of human retinal pigment epithelial cells (in Chinese). Yan Ke Xue
43. Topp KS, Bisla K, Saks ND, et al. Centripetal transport of herpes
simplex virus in human retinal pigment epithelial cells in vitro.
44. Tian J, Ishibashi K, Honda S, et al. The expression of native and
cultured human retinal pigment epithelial cells grown in different
culture conditions. Br J Ophthalmol. 2005;89:1510–1517.
45. Pfeffer BA, Clark VM, Flannery JG, et al. Membrane receptors for
retinol-binding protein in cultured human retinal pigment epithe-
lium. Invest Ophthalmol Vis Sci. 1986;27:1031–1040.
46. Lauber JK. Diurnal mitochondrial changes in avian retinal pigment
epithelium: a search for correlation with thyroid state. Curr Eye
Res. 1982;2:863–868, 1983.
47. Heth CA, Yankauckas MA, Adamian M, et al. Characterization of
retinal pigment epithelial cells cultured on microporous filters.
Curr Eye Res. 1987;6:1007–1019.
48. Arndt C, Sari A, Ferre M, et al. Electrophysiological effects of
corticosteroids on the retinal pigment epithelium. Invest Ophthal-
mol Vis Sci. 2001;42:472–475.
49. Campochiaro PA, Hackett SF, Conway BP. Retinoic acid promotes
density-dependent growth arrest in human retinal pigment epithe-
lial cells. Invest Ophthalmol Vis Sci. 1991;32:65–72.
50. van den Berg JJ, Cook NE, Tribble DL. Reinvestigation of the
antioxidant properties of conjugated linoleic acid. Lipids. 1995;
51. Kaven CW, Spraul CW, Zavazava NK, et al. Growth factor combi-
nations modulate human retinal pigment epithelial cell prolifera-
tion. Curr Eye Res. 2000;20:480–487.
52. Brewer GJ, Torricelli JR, Evege EK, et al. Optimized survival of
hippocampal neurons in B27-supplemented Neurobasal, a new
serum-free medium combination. J Neurosci Res. 1993;35:567–
53. Svendsen CN, Fawcett JW, Bentlage C, et al. Increased survival of
rat EGF-generated CNS precursor cells using B27 supplemented
medium. Exp Brain Res. 1995;102:407–414.
54. Rezai KA, Farrokh-Siar L, Botz ML, et al. Biodegradable polymer
film as a source for formation of human fetal retinal pigment
798Gamm et al.
IOVS, February 2008, Vol. 49, No. 2
epithelium spheroids. Invest Ophthalmol Vis Sci. 1999;40:1223–
55. Tezel TH, Del Priore LV. Reattachment to a substrate prevents
apoptosis of human retinal pigment epithelium. Graefes Arch Clin
Exp Ophthalmol. 1997;235:41–47.
56. Gabrielian K, Oganesian A, Farrokh-Siar L, et al. Growth of human
fetal retinal pigment epithelium as microspheres. Graefes Arch
Clin Exp Ophthalmol. 1999;237:241–248.
57. Chang CW, Ye L, Defoe DM, et al. Serum inhibits tight junction
formation in cultured pigment epithelial cells. Invest Ophthalmol
Vis Sci. 1997;38:1082–1093.
58. Rahner C, Fukuhara M, Peng S, et al. The apical and basal environ-
ments of the retinal pigment epithelium regulate the maturation of
tight junctions during development. J Cell Sci. 2004;117:3307–
59. Dawson DW, Volpert OV, Gillis P, et al. Pigment epithelium-
derived factor: a potent inhibitor of angiogenesis. Science. 1999;
60. Barnstable CJ, Tombran-Tink J. Neuroprotective and antiangio-
genic actions of PEDF in the eye: molecular targets and therapeutic
potential. Prog Retin Eye Res. 2004;23:561–577.
61. Tombran-Tink J, Shivaram SM, Chader GJ, et al. Expression, secre-
tion, and age-related downregulation of pigment epithelium-de-
rived factor, a serpin with neurotrophic activity. J Neurosci. 1995;
62. Behling KC, Surace EM, Bennett J. Pigment epithelium-derived
factor expression in the developing mouse eye. Mol Vis. 2002;8:
63. McFarlane S, Glenn JV, Lichanska AM, et al. Characterisation of the
advanced glycation endproduct receptor complex in the retinal
pigment epithelium. Br J Ophthalmol. 2005;89:107–112.
64. Hughes RC. Secretion of the galectin family of mammalian carbo-
hydrate-binding proteins. Biochim Biophys Acta. 1999;1473:172–
65. Beuckmann CT, Gordon WC, Kanaoka Y, et al. Lipocalin-type
prostaglandin D synthase (beta-trace) is located in pigment epithe-
lial cells of rat retina and accumulates within interphotoreceptor
matrix. J Neurosci. 1996;16:6119–6124.
66. Moshfeghi DM, Blumenkranz MS. Role of genetic factors and in-
flammation in age-related macular degeneration. Retina. 2007;27:
67. Sivaprasad S, Chong NV. The complement system and age-related
macular degeneration. Eye. 2006;20:867–872.
68. Wasselius J, Hakansson K, Johansson K, et al. Identification and
localization of retinal cystatin C. Invest Ophthalmol Vis Sci. 2001;
69. Wakasugi K, Nakano T, Morishima I. Association of human neuro-
globin with cystatin C, a cysteine proteinase inhibitor. Biochem-
70. Paraoan L, Grierson I, Maden BE. Fate of cystatin C lacking the
leader sequence in RPE cells. Exp Eye Res. 2003;76:753–756.
71. Zurdel J, Finckh U, Menzer G, et al. CST3 genotype associated with
exudative age related macular degeneration. Br J Ophthalmol.
72. Paraoan L, Grierson I, Maden BE. Analysis of expressed sequence
tags of retinal pigment epithelium: cystatin C is an abundant
transcript. Int J Biochem Cell Biol. 2000;32:417–426.
73. Rakoczy PE, Sarks SH, Daw N, et al. Distribution of cathepsin D in
human eyes with or without age-related maculopathy. Exp Eye
74. Hoppe G, O’Neil J, Hoff HF, et al. Products of lipid peroxidation
induce missorting of the principal lysosomal protease in retinal
pigment epithelium. Biochim Biophys Acta. 2004;1689:33–41.
75. Pfeffer BA, Becerra SP, Borst DE, et al. Expression of transthyretin
and retinol binding protein mRNAs and secretion of transthyretin
by cultured monkey retinal pigment epithelium. Mol Vis. 2004;10:
76. Ong DE, Davis JT, O’Day WT, et al. Synthesis and secretion of
retinol-binding protein and transthyretin by cultured retinal pig-
ment epithelium. Biochemistry. 1994;33:1835–1842.
77. Jaworowski A, Fang Z, Khong TF, et al. Protein synthesis and
secretion by cultured retinal pigment epithelia. Biochim Biophys
78. Mullins RF, Russell SR, Anderson DH, et al. Drusen associated with
aging and age-related macular degeneration contain proteins com-
mon to extracellular deposits associated with atherosclerosis, elas-
tosis, amyloidosis, and dense deposit disease. FASEB J. 2000;14:
79. Hahn P, Qian Y, Dentchev T, et al. Disruption of ceruloplasmin
and hephaestin in mice causes retinal iron overload and retinal
degeneration with features of age-related macular degeneration.
Proc Natl Acad Sci USA. 2004;101:13850–13855.
80. Hahn P, Dentchev T, Qian Y, et al. Immunolocalization and regu-
lation of iron handling proteins ferritin and ferroportin in the
retina. Mol Vis. 2004;10:598–607.
81. Chen L, Dentchev T, Wong R, et al. Increased expression of
ceruloplasmin in the retina following photic injury. Mol Vis. 2003;
82. Vassiliev V, Harris ZL, Zatta P. Ceruloplasmin in neurodegenerative
diseases. Brain Res Rev. 2005;49:633–640.
83. Araki Y, Miyagi N, Kato N, et al. Coordinated metabolism of
alcadein and amyloid beta-protein precursor regulates FE65-depen-
dent gene transactivation. J Biol Chem. 2004;279:24343–24354.
84. Araki Y, Tomita S, Yamaguchi H, et al. Novel cadherin-related
membrane proteins, alcadeins, enhance the X11-like protein-me-
diated stabilization of amyloid beta-protein precursor metabolism.
J Biol Chem. 2003;278:49448–49458.
85. Yoshida T, Ohno-Matsui K, Ichinose S, et al. The potential role of
amyloid beta in the pathogenesis of age-related macular degener-
ation. J Clin Invest. 2005;115:2793–2800.
IOVS, February 2008, Vol. 49, No. 2
Serum-Free Human RPE Culture 799