sequential induction of angiogenic growth factors by tnf-α in choroidal endothelial cells

www.elsevier.com/locate/jneuroim

Journal of Neuroimmunolo

Sequential induction of angiogenic growth factors by TNF-a in

choroidal endothelial cellsB

Masanori Hangai a, Shikun He b, Stephan Hoffmann c, Jennifer I. Lim a,

Stephen J. Ryan a,c, David R. Hinton a,b,c,*

a Department of Ophthalmology, Doheny Eye Institute, Keck School of Medicine of the University of Southern California, Los Angeles, CA, United Statesb Department of Pathology, Doheny Eye Institute, Keck School of Medicine of the University of Southern California, Los Angeles, CA, United States

c Beckman Macular Research Center, Doheny Eye Institute, Keck School of Medicine of the University of Southern California, Los Angeles, CA, United States

Received 25 April 2005; accepted 19 September 2005

Abstract

Inflammatory mediators have been proposed to play a critical role in the pathogenesis of choroidal neovascularization, a blinding

complication of age-related macular degeneration. We evaluated the expression of TNF-a in human choroidal neovascular membranes and

found that it colocalized with cells expressing VEGF, angiopoietin (Ang)-1 and Ang2. In cultured choroidal endothelial cells we found that

TNF-a increased Ang2 mRNA (increased transcription) and protein levels prior to those of Ang1 and VEGF. The results raise the possibility

that during neovascularization, TNF-a may modulate endothelial plasticity and survival by sequential inactivation of Tie2 followed by

activation of Tie2 and VEGF receptors.

D 2005 Elsevier B.V. All rights reserved.

Keywords: Angiogenesis; Choriocapillaris; Cytokine; Growth factors; Gene expression

1. Introduction

Choroidal neovascularization (CNV) is an important

blinding complication of a wide range of ocular eye disorders,

most notably age-related macular degeneration (AMD)

(Macular Photocoagulation Study Group, 1991; Ambati et

al., 2003; D’Amato and Adamis, 1995; Ryan et al., 2001).

The mechanism of CNV formation has not been established,

but local upregulation of angiogenic growth factors such as

vascular endothelial growth factor (VEGF) and angiopoietins

0165-5728/$ - see front matter D 2005 Elsevier B.V. All rights reserved.

doi:10.1016/j.jneuroim.2005.09.018

i This work was supported by NIH grants EY01545, EY03040 and

grants from the Arnold and Mabel Beckman Foundation, and Research to

Prevent Blindness. Masanori Hangai was supported by Kyoto University

Foundation, Japan National Society for the Prevention of Blindness and

Nippon Eye Bank Association.

* Corresponding author. Department of Ophthalmology, Doheny Eye

Institute, Keck School of Medicine of the University of Southern

California, Los Angeles, CA, United States.

E-mail address: [email protected] (D.R. Hinton).

(Ang) has been reported and is thought to be critical for the

process (Frank et al., 1996; Ishibashi et al., 1997; Lopez et al.,

1996; Otani et al., 1999; Spilsbury et al., 2000).

Increasing evidence in human CNV specimens (Gehrs et

al., 1992; Lopez et al., 1991; Oh et al., 1999b; Saxe et al.,

1993) and in experimental CNV models (Kimura et al.,

1999; Pollack et al., 1986) has also implicated inflamma-

tion, and in particular macrophages (Sakurai et al., 2003), in

the pathogenesis of CNV and AMD; these studies support

the hypothesis that inflammatory cytokines such as tumor

necrosis factor-a (TNF-a) may play an important role in the

pathogenesis of this disorder.

TNF-a is a pleiotropic cytokine that mediates inflamma-

tory, proliferative, cytostatic, and cytotoxic effects in a

variety of cell types, including endothelial cells (Mantovani

et al., 1992). In addition, it is a potent inducer of

angiogenesis in vivo (Frater-Schroder et al., 1987; Leibo-

vich et al., 1987; Montrucchio et al., 1994). Despite its

potent angiogenic activity in vivo, TNF-a seems to inhibit

in vitro angiogenic activities such as endothelial prolifera-

gy 171 (2006) 45 – 56

M. Hangai et al. / Journal of Neuroimmunology 171 (2006) 45–5646

tion and tube formation (Frater-Schroder et al., 1987; Sato et

al., 1987), suggesting that TNF-a may induce angiogenesis

indirectly by activating other regulators of angiogenesis.

Therefore, it is critical to determine the mechanism by

which TNF-a regulates angiogenic factors that stimulate

angiogenesis by acting directly on endothelial cells.

Members of the VEGF and angiopoietin families are

important in the process of vasculogenesis and angiogenesis

(Ferrara et al., 1995, 1996; Gale and Yancopoulos, 1999,

Carmeliet et al., 1996; Fong et al., 1995; Sato et al., 1995).

VEGF is a well-characterized angiogenic factor that is

upregulated by hypoxia and inflammatory stimuli (Ben-Av

et al., 1995; Shweiki et al., 1992). In the eye, VEGF has

been shown to play a key role in neovascularization

involving the retinal and choroidal circulations (D’Amore,

1994, Adamis et al., 1994; Aiello et al., 1994; Cui et al.,

2000; Frank et al., 1996; Lopez et al., 1996; Miller et al.,

1994; Okamoto et al., 1997). Recent studies have demon-

strated the importance of cooperative interaction of VEGF

with the angiopoietins in angiogenesis and wound healing

(Asahara et al., 1998; Holash et al., 1999; Jones et al., 2001;

Kampfer et al., 2001; Maisonpierre et al., 1997).

The angiopoietin family consists of angiopoietin-1

(Ang1), angiopoietin-2 (Ang2) and their receptor Tie2

(tyrosine kinase with immunoglobulin-like loops and epi-

dermal growth factor homology domains). Ang1 is an

agonistic ligand for Tie2 and induces phosphorylation of

Tie2 (Davis et al., 1996). Contrary to this effect, Ang2 binds

to Tie2, but inhibits Ang1-mediated Tie2 phosphorylation

(Maisonpierre et al., 1997). While Ang1 is not mitogenic for

endothelial cells, it stimulates endothelial cell migration and

sprouting, and is synergistic with VEGF in the latter activity

(Koblizek et al., 1998; Witzenbichler et al., 1998). Ang1

likely plays a key role in vascular maturation and survival in

cooperation with VEGF in vivo (Asahara et al., 1998;

Papapetropoulos et al., 1999; Suri et al., 1996; Vikkula et

al., 1996). Furthermore, Ang1 overexpression results inmuch

enhanced angiogenesis, suggesting that Ang1 is capable of

inducing angiogenesis in vivo (Suri et al., 1998). Importantly,

Ang2 also appears to be angiogenic in vivo, because in the

presence of VEGF, Ang2 enhances VEGF-mediated angio-

genesis, probably by blocking the stabilizing or maturing

function of Ang1, thus allowing vessels to respond better to a

sprouting signal by VEGF (Asahara et al., 1998; Hackett et

al., 2000; Maisonpierre et al., 1997). In vitro studies using

retinal cells have shown that the retinal pigment epithelial cell

(RPE), a key source of VEGF in CNV (Lopez et al., 1996),

upregulates Ang1 in response to VEGF (Hangai et al., 2001).

Despite the potential importance of TNF-a and these

endothelial-specific angiogenic mediators in postnatal path-

ologic vascular remodeling, the mechanism by which they

interact is still not clear. Here, we show that TNF-a

colocalizes with Ang1, Ang2 and VEGF in human CNV

specimens, suggesting autocrine or paracrine interactions

between these factors. We then demonstrate that TNF-a

induces the sequential upregulation of Ang2 and then Ang1

and VEGF mRNA and protein expression in choroidal

microvascular endothelial cells in vitro. Thus, TNF-a may

exert its profound angiogenic action in vivo by stimulating

the appropriate sequence of endothelial-specific angiogenic

factors.

2. Materials and methods

2.1. Choroidal neovascular membranes (CNVM)

Surgical excision of AMD-related, subfoveal CNVMs

was performed in 12 eyes from 12 patients. All specimens

were obtained by one of the authors (JIL, 5 specimens)

during the course of patient treatment or were present in our

frozen archives (7 specimens). The tenets of the Declaration

of Helsinki, Finland, were followed — informed consent

was obtained, and approval by the institutional review board

(University of Southern California) was granted for this

study. Seven of the 12 specimens had been evaluated

previously for expression of other growth factors (Lopez et

al., 1996). Each of the fresh, surgically excised CNVMs was

placed in isotonic saline at 4 -C, then snap-frozen in

optimum cutting temperature compound (OCT, Ames/Miles

Inc., Elkhart, Ind.) within 1 h. Each specimen was serially

sectioned on a cryostat into 6-Am frozen sections on glass

slides. The sections were fixed in reagent grade acetone for

5 min at room temperature and stored at �80 -C.

2.2. Immunohistochemistry

Thawed sections were air-dried, fixed with reagent grade

acetone for 5 min, and washed with TRIS buffer (pH 7.4).

Sections were blocked for 15 min with 1% bovine serum

albumin (Sigma, St. Louis, MO) in TRIS buffer after

endogenous peroxide was blocked by 0.3% hydrogen

peroxide. The sections were incubated for 30 min with the

primary antibody, washed for 15 min with TRIS buffer, and

staining completed using the ABC immunoperoxidase kit or

alkaline phosphatase kit (Vector Laboratories, Burlingame

CA). The chromogen was either aminoethylcarbizole (AEC,

Vector) or the blue alkaline phosphatase substrate (kit II,

Vector). Most sections were counterstained with Mayer’s

modified hematoxylin. Negative controls included omission

of primary antibody or an irrelevant polyclonal or isotype-

matched monoclonal primary antibody; in all cases negative

controls showed only faint, insignificant staining. Human

Ang1, and Ang2 rabbit polyclonal antibodies (1 : 500

dilution) were provided by Regeneron Pharmaceuticals,

Inc. Specificity of the Ang1 and Ang2 antibodies was

confirmed by absorption test using an excess amount of the

peptides used to raise these antibodies (Ang1: N-terminal

peptide, NQRRSPENSGRRYNRIQHGQ; Ang2: N-termi-

nal peptide, NFRKSMDSIGKKQYQVQHGS). Polyclonal

rabbit antibody against human VEGF (1 :100 dilution) was

obtained from Santa Cruz Biotechnology (Santa Cruz, CA).

M. Hangai et al. / Journal of Neuroimmunology 171 (2006) 45–56 47

Monoclonal antibody against TNF-a (1 :100 dilution) was

obtained from R&D Systems (Minneapolis, MN). Markers

to specify cell types in the CNVM included monoclonal

antibody against cytokeratin 18 for RPE (1 :400 dilution,

Sigma), and CD31 for endothelial cells (1 :100 dilution,

DAKO Corporation, Carpinteria, CA). Double-label immu-

nohistochemistry consisted of performing immunohisto-

chemistry first with a polyclonal antibody and red

chromogen, washing profusely with TRIS buffer, followed

by monoclonal antibody immunohistochemistry using the

blue chromogen.

2.3. Cell culture

Bovine choroidal microvascular endothelial cells

(BCEC) were cultured as described previously with a

slight modification (Hoffmann et al., 1998). Briefly,

sensory retina was cut at the optic disk and the RPE

layer was removed by gentle scraping. The choroid was

mechanically dissected and microvessels were collected

with the aid of forceps under a dissecting microscope.

Large vessels were removed by sifting through 200 Amnylon mesh. The filtrate was washed three times with

Hank’s balanced salt solution (HBSS) containing 0.1%

bovine serum albumin (HBSS-BSA), incubated with 0.5%

trypsin for 20 min at room temperature, washed with

HBSS-BSA again, and further digested with HBSS

containing 0.1% collagenase (Boehringer Mannheim,

Indianapolis, Ind.), 0.15 mg/ml tosyl-lysine-chlor-methyl-

ketone (Sigma) and 20 U/ml type2 deoxyribonuclease I

(Sigma) at 37 -C for 30 min. The cell suspension was

then filtered through a 70 Am nylon mesh and the filtrate

was centrifuged to collect the digested microvascular

fragments, which were concentrated into 400 Al media

containing LEA (bovine specific lectin from Lycopersicon

esculentum; Sigma)-coated magnetic beads (Dynal, Oslo,

Norway). The bead–cell suspension was incubated at 4

-C for 1 h with agitation. To remove non-specifically

binding cells from the beads, the bead–cell complex was

washed 10 times using the manufacturer’s magnetic

device. The cells that were attached to the magnetic

beads were seeded on fibronectin-coated 6-well plates and

were grown in endothelial growth medium (EGM,

Clonetics, San Diego, CA) supplemented with 10%

FBS, bovine brain extract (12 Ag/ml), human epidermal

growth factor (10 ng/ml) and hydrocortisone (1 Ag/ml).

The identity of BCEC was confirmed by their cobblestone

morphology on phase–contrast microscopy and the purity

was determined by counting the number of the cells that

were immunoreactive for von Willebrand factor (Sigma)

and incorporated dil-acetylated low-density lipoprotein

(Biomedical Technologies, Stoughton, MA) (Hoffmann

et al., 1998). Only cultures that had more than 98% of

von Willebrand factor- and dil-acetylated low-density

lipoprotein-positive cells were used for analysis (results

not shown).

2.4. Northern blot

Total RNA was extracted from BCEC cells using the

Trizol reagent (Gibco BRL, Gaithersburg, MD) according to

the manufacturer’s instructions. Equal amounts of total

RNA (10–20 Ag/lane) were fractionated on 1% (wt/vol)

agarose/6.3% formaldehyde gels, and blotted on a nylon

membrane (Nylon Duralon-UV; Stratagene, La Jolla, CA)

using the traditional capillary system in 10� SSPE (1.5 M

NaCl, 100 mM sodium phosphate, and 10 mM Na2 EDTA).

Filters were then UVcross-linked (UV Stratalinker 1800;

Stratagene). Ang1, Ang2, and VEGF cDNAs were kindly

provided by Regeneron Pharmaceuticals, Inc. The cDNAs

were labeled (specific activity of 1�109 cpm/Ag) using a

random primer labeling kit (Rediprime; Amersham Phar-

macia Biotech, Piscataway, NJ) according to the manufac-

turer’s instructions. Membranes were hybridized at 68 -Cfor 2 h (ExpressHyb Hybridization Solution; Clontech

Laboratories, Palo Alto, CA) in a solution containing 0.1

mg/ml denatured salmon sperm DNAwith 2 to 3�106 cpm/

ml 32P-labeled Ang1 (a 0.57-kb SpeI–EcoRI fragment of

human Ang1 cDNA), Ang2 (a 0.64-kb EcoRI–HindIII

fragment of human Ang2 cDNA), or VEGF (a 0.6-kb

BamHI fragment of human VEGF cDNA). After hybridiza-

tion, filters were washed three times in 2� SSPE/0.1% SDS

for 15 min at room temperature and then in 0.1� SSPE/

0.1% SDS for 30 min at 60 -C. To correct for differences in

RNA loading, the filters were stripped and rehybridized

with a human S18 rRNA probe (Ambion, Austin, TX). The

filters were scanned, and radioactivity was quantified by

computer imaging (PhosphoImager with ImageQuant soft-

ware; Molecular Dynamics, Sunnyvale, CA).

2.5. mRNA stability analysis

BCECs were exposed to vehicle or TNF-a (10 ng/ml) for

1 h (Ang2) or 12 h (Ang1) and then incubated with

actinomycin D (5 Ag/ml) to stop RNA synthesis. Total RNA

was isolated at the indicated time points and used for

Northern hybridization as described above.

2.6. Nuclear run-on analysis

BCECs were treated with vehicle or TNF-a (10 ng/ml)

for 1 h (Ang2) or 12 h (Ang1). The cells were washed twice

with ice-cold phosphate-buffered saline (PBS), scraped off

the dish in ice-cold SSC (150 mM sodium chloride and 15

mM sodium citrate) and collected in a 15-ml tube by

centrifugation at 500 �g for 5 min at 4 -C. Subsequent stepswere performed at 4 -C. The cells were resuspended in 4 ml

of lysis buffer (10 mM Tris–HCl, pH 7.4, 10 mM NaCl, 3

mM MgCl2 and 0.5% Nonidet P-40) and then were

disrupted with Dounce homogenizers (10 strokes). Nuclei

were pelleted by centrifugation at 500 �g for 5 min and

were resuspended in 100 Al of glycerol storage buffer (10

mM Tris–HCl, pH 8.3, 40% (v/v) glycerol, 5 mM MgCl2,


0.1 mM EDTA) and frozen in liquid nitrogen. The nuclear

suspension was mixed with 0.1 ml of 2� reaction buffer

(100 mM Hepes, pH 8.0, 10 mM MgCl2, 300 mM KCl, 200

U of RNasin (Roshe Molecular Biochemicals, Indianapolis,

IN) per ml per 1 mM each ATP, GTP and CTP per 150 ACi(1 ACi=37 kBq) of [32P]UTP (3000 Ci/mmol; Amersham

Pharmacia Biotech)) and incubated for 30 min at 30 -C.Transcription was stopped by adding 20 Ag of DNase I,

followed by 80 Ag of proteinase K. The 32P-labeled RNA

was purified by extraction with phenol/chloroform and two

sequential precipitations with ammonium acetate. Equal

amounts of 32P-labeled RNA were hybridized in 50%

formamide, 5� SSC, 5� Denhardt’s solution, 1% SDS

(1� SSC=150 mM NaCl, 15 mM sodium citrate, pH 7.0) at

42 -C for 72 h. Filters contained 5–0.5 Ag each of linearizedplasmids immobilized on Zeta-Probe GT membranes (Bio

Rad Laboratories, Hercules, CA) after blotting in 12� SSPE

with a Bio-Dot SF microfiltration apparatus (Bio-Rad).

Filters were washed three times with 2� SSC, 0.1% SDS at

42 -C for 5 min, followed by two washes with 0.2� SSC,

0.1% SDS at 65 -C for 15 min, and then analyzed using a

PhosphoImager as described above. The Ang1 and Ang2

mRNA amount was standardized by comparison with the

amount of h-actin mRNA.

2.7. Western blot

The medium from established BCEC cell cultures was

replaced with serum-free defined medium and grown for 24

h, whereupon the cells were exposed to vehicle or TNF-a

(10 ng/ml). The conditioned medium was collected and cell

debris was removed by centrifugation at 14,000 �g. The

supernatant was used for Western blotting. A Bradford-

based assay kit (Bio-Rad) was used to measure protein

concentrations. Equal amounts of protein were fractionated

by 10% SDS-polyacrylamide gel electrophoresis and

transferred to polyvinylidene difluoride membranes (Immo-

bilon, Millipore Corp., Bedford, MA), and then probed with

polyclonal rabbit anti-human Ang1 (0.45 Ag/ml) and anti-

human Ang2 antibodies (0.22 Ag/ml) and a polyclonal

rabbit anti-human VEGF antibody (Santa Cruz Biotechnol-

ogy, Inc., CA). Membranes were washed and incubated with

a horseradish peroxidase (HRP)-conjugated goat anti-rabbit

IgG secondary antibody (Vector Laboratories, Burlingame,

CA) for 1 h at room temperature. Immunoreactive bands

were identified by adding ECL chemiluminescence detec-

tion solution (Amersham Pharmacia, Cleveland, OH). The

membranes were scanned and blue fluorescence intensity

was quantified on a Storm PhosphoImager running the

ImageQuant software (Molecular Dynamics).

2.8. Statistical analysis

Experiments were performed at least in triplicate. Values

were expressed as meanTSEM. Factorial ANOVA followed

by Fisher’s least significant test was performed and a P

value of 0.05 was considered significant. Dose–response

curves were analyzed by linear regression.

3. Results

3.1. Immunohistochemistry of human choroidal neovascular

membranes

The choroidal neovascular membranes (CNVMs) were

classified as vascular, fibrotic, or mixed by their appear-

ance on hematoxylin and eosin stains. The identity of

vascular channels was confirmed by CD31 staining. Of the

12 cases, 2 were predominantly vascularized, 2 were

predominantly fibrous, and 8 showed various amounts of

both vascular and fibrous tissue. The vascular tissue

contained many stromal cells; we have previously shown

that stromal cells are positive for cytokeratin 18 suggesting

that they are derived from the retinal pigment epithelium

(RPE) (Lopez et al., 1996). Most membranes had small

foci of residual RPE monolayer within the membranes.

Ang1, Ang2, and VEGF were positive to some extent in

all surgically excised CNVMs and were most prominent in

the vascular regions of the membranes. Ang2 (Fig. 1A)

was most prominent in large and small vascular channels

and was present to a lesser degree in stromal cells. In

contrast, Ang1 (Fig. 1B) staining was most prominent in

stromal cells of the CNVM and was focally present within

small capillary channels. VEGF (Fig. 1C) was present in

both large and small vascular channels and stromal cells.

TNF-a immunoreactivity was strongly localized selectively

to the vascular portions of the membranes and was

predominantly localized with the vascular endothelium

(Fig. 1E) and partially localized to stromal cells and

macrophages.

Human CNVMs have strong autofluorescence derived

from lipofuscin and pigments in RPE cells; this causes some

false positive staining when immunofluorescent methods

are used. Therefore, double-label immunohistochemistry

using two chromogens was chosen to demonstrate the

colocalization of the angiopoietins with vascular endothe-

lium and TNF-a. Cells positive for TNF-a and Ang1 were

found adjacent to one another; however, occasional cells

showed colocalization in small vascular channels (Fig. 1F

and inset). In contrast, Ang2 more frequently colocalized

with TNF-a in both large and small vascular channels in the

CNVMs (Fig. 1E and inset). VEGF also showed some

colocalization with TNF-a in the vascular channels (Fig. 1G

and inset).

3.2. Time- and dose-dependent regulation of mRNA

expression for angiopoietin-1, -2 and VEGF by TNF-a in

BCECs

Our histologic analysis suggested that TNF-a might

be an important autocrine and/or paracrine regulator of

Fig. 1. (A–D) Localization of growth factors in CNVM. Immunohistochemical staining of the vascular component of a representative highly vascularized

CNV membrane is shown. Localization of Ang2 (A), Ang1 (B), and VEGF (C) are shown in similar regions of a membrane. Immunoperoxidase staining uses

aminoethylcarbizole (A–D) as a red chromogen; nuclei are counterstained lightly with hematoxylin (A–D). Control for single immunoperoxidase stain in

which the primary antibody as adsorbed with excess antibody-specific peptide (e.g., Ang2 adsorbed antibody shown in (D)) with hematoxylin counterstain.

Arrows identify blood vessels, arrow heads identify stromal cells. Bar=100 Am.

(E–H) Localization of growth factors and TNF-a in CNVM. Immunohistochemical staining of the vascular component of a representative highly vascularized

CNV membrane is shown. Double staining for Ang2 (red) and TNF-a (blue) (E); Ang1 (red) and TNF-a (blue) (F); VEGF (red) and TNF-a (blue) (G) is

demonstrated. Immunoperoxidase staining uses aminoethylcarbizole (E–H) as a red chromogen while immunoalkaline phosphatase staining uses a blue

chromogen (E–H). There is no nuclear counterstain in the double-stained sections (E–H). Control for double immunoperoxidase stain in which both primary

antibodies are omitted but the remainder of the staining procedure is performed, (H). Arrows identify blood vessels, and arrow heads identify stromal cells.

Bar=100 Am. Insets in (E–G), magnified 2� further.


endothelial Ang1 and Ang2 in CNV. Northern blot

analysis revealed that both Ang1 and Ang2 mRNAs

were expressed at low levels in serum-starved bovine

choroidal endothelial cells (BCECs). In response to

stimulation with TNF-a (10 ng/ml), Ang2 mRNA levels

increased rapidly to a maximum of 2.1-fold (P <0.05)

over control at 1 h (Fig. 2A). After 8 h, the Ang2

mRNA level returned to base line. In contrast, Ang1

mRNA levels remained steady from 0 to 2 h after

TNF-a stimulation, but increased after 4 h to a

Fig. 2. Time course of mRNA induction of Ang2 and Ang1/VEGF by TNF-a in bovine choroidal microvascular endothelial cells. TNF-a induced sequential

upregulation of Ang2 mRNA (2.8 kB band) (A) and then Ang1 mRNA (4.4 kB band) and VEGF mRNA (4.0 kB band) (B). Total RNA was extracted from

BCECs at the indicated times after stimulation with TNF-a (10 ng/ml) and subjected to Northern analysis (15 Ag of total RNA/lane). Results were quantified ona PhosphoImager running the ImageQuant software. Differences in loading were normalized using the signal intensity of S18. The corrected density was

plotted as a percentage of the 0-h value. Results are meanTSEM from three independent cultures for each time point. Blots obtained from one representative

membrane are shown (the S18 control bands are shared by Ang1, Ang2 and VEGF).


maximum of 9.2-fold (P <0.02) over control at 24

h (Fig. 2B). The time course for TNF-a-regulation of

VEGF mRNA levels was similar to that of Ang1.

Fig. 3. Dose–response of mRNA induction of Ang2 and Ang1 by TNF-a in chor

and Ang1 (4.4 kB band) (B) mRNAs by TNF-a was dose-dependent. BCECs w

extracted at 1 h for Ang2 and at 24 h for Ang1 after the stimulation, and analyzed a

the control value. Results are meanTSEM from three independent cultures for ea

VEGF mRNA levels reached a maximum of 5.5-fold

(P <0.05) increase at 12 h (Fig. 2B). To confirm that

the effects of TNF-a on the induction of Ang1 and

oidal microvascular endothelial cells. Induction of Ang2 (2.8 kB band) (A)

ere stimulated with the indicated concentration of TNF-a. Total RNA was

s described for Fig. 2. The corrected intensity was plotted as a percentage of

ch concentration and the representative blots are shown.

Fig. 4. Western blot analysis comparing the time course of induction of Ang2 and Ang1/VEGF by TNF-a in choroidal microvascular endothelial cells. TNF-a

selectively upregulated Ang2 secretion (65 kDa band) after 6 h (A) and then upregulated Ang1 (70 kDa band) and VEGF (24 kDa band) secretion at 24 h (B).

BCECs were exposed to vehicle or TNF-a (10 ng/ml) and conditioned media were collected at the indicated times after the stimulation. Approximately 5 Ag ofprotein/lane was resolved by 10% polyacrylamide gel electrophoresis, transferred to a polyvinylidene difluoride membrane, and immunoblotted using

polyclonal antibodies to Ang1, Ang2 or VEGF. Bound antibodies were detected using a horseradish peroxidase-based chemoluminescence method and

quantified using a Storm PhosphoImager running the ImageQuant software. The intensity was plotted as a percentage of the vehicle value. Three independent

experiments were performed and each showed similar results; a representative blot with quantitation is shown.


Ang2 mRNA are dose-dependent, cells were stimulated

with various concentrations of TNF-a for 1 h (Ang2)

and 12 h (Ang1). The upregulation of Ang2 and Ang1

mRNA was dose-dependent with an ED50 of 0.24 ng/

ml and a maximal 3.3-fold (P <0.05) increase for

Ang2, and an ED50 of 2.2 ng/ml and a maximal 4.0-

fold (P <0.05) increase for Ang1 (Fig. 3). As a control

for the possibility that the endothelial cells were

conditioning the medium to increase growth factor

levels, we followed the expression of Ang1 and Ang2

for 24 h in culture without TNF-a stimulation; no

significant change in expression of either Ang1 or Ang2

was found (results not shown).

Fig. 5. Effect of TNF-a on Ang1 and Ang2 mRNA half-lives in choroidal micro

Ang1 (B) mRNA half-lives. BCECs were exposed to vehicle or TNF-a (10 ng/ml)

Total RNAwas extracted from the cells at the indicated times after actinomycin D

values of the corrected intensity were plotted as a percentage of the 0-h value in

3.3. Effects of TNF-a on Ang1, Ang2 and VEGF protein

levels

To determine whether the time-dependent increases in

Ang1, Ang2 and VEGF mRNA in BCECs were

associated with increases in Ang1, Ang2 and VEGF

protein levels, Western blot analysis was performed using

conditioned media (Fig. 4). For the untreated cells, Ang2

accumulation in the medium reached a maximum at 24

h. After 6 h of stimulation with TNF, there were

significantly increased levels of Ang2 in the medium

compared to controls at the same time point (2.2-fold,

P <0.05). By 24 h, the level of Ang2 in the medium

vascular endothelial cells. TNF-a did not significantly alter Ang2 (A) and

for 1 h (Ang2) or 12 h (Ang1) before addition of actinomycin D (5 Ag/ml).

treatment and Northern blot analysis was performed as in Fig. 2. The mean

logarithmic scale.


was equal in controls and TNF-treated samples suggest-

ing that the induction of Ang2 was early and transient.

Levels of both Ang1 and VEGF (Fig. 4B) proteins were

steady before 8 h, and increased only modestly in the

medium from untreated cells at 24 h, while in the

medium from TNF-treated cells there was increased

secretion of both Ang1 and VEGF above control values

at 24 h (1.6-fold for Ang1, P <0.01; 1.8-fold for VEGF,

P <0.05).

3.4. Transcriptional regulation of Ang1 and Ang2 by TNF-a

To elucidate the mechanism by which TNF-a

upregulates Ang1 and Ang2 mRNA, we measured

mRNA stability and transcription rate. The half-lives of

Ang1 and Ang2 mRNA at base line were approximately

3.7 and 1.7 h, respectively (Fig. 5), while the half lives

after exposure to TNF-a were 3.7 and 1.8 h, respec-

tively. Nuclear run-on experiments were performed to

examine the rate of transcription. Treatment with TNF-a

(10 ng/ml) increased transcription of the Ang2 gene 3.0-

fold (P <0.05), and the Ang1 gene 4.7-fold (P <0.05)

(Fig. 6A, B). These experiments demonstrate that the

increase in Ang1 and Ang2 in BCEC stimulated by

TNF-a was due to an increase in Ang1 and Ang2 gene

transcription.

Fig. 6. Effect of TNF-a on Ang1 and Ang2 transcription rate in choroidal microva

kB band) (A) and Ang1 (4.4 kB band) (B). BCECs were exposed to vehicle or TN

vitro transcription was allowed to resume in the presence of [a-32P] UTP. Equal

which each cDNAwas immobilized. Results were quantified on a PhosphoImager

using the signal intensity of h-actin. The corrected density was plotted as a perce

cultures and the representative blots are shown.

3.5. Induction of Ang1 by TNF-a in BCECs requires new

protein synthesis

To determine whether induction of Ang1 mRNA requires

de novo protein synthesis, we treated BCECs with the

protein synthesis inhibitor cycloheximide (2 Ag/ml) for 1

h before adding TNF-a (10 ng/ml) (Fig. 7). Interestingly, a

probable splice variant of larger size (4.8 kB), in addition to

the usual 4.4 kB transcript was detected following cyclo-

heximide treatment in non-TNF-treated BCECs. Effects of

TNF-a on the induction of Ang1 mRNA were abolished

(P <0.05) by cycloheximide, suggesting that induction of

Ang1 by TNF-a requires de novo protein synthesis.

4. Discussion

Ang2 mRNA expression in vivo is normally present at

the leading edge of invading vascular sprouts in the

developing fetus, and within zones of physiologic angio-

genesis in the adult (Maisonpierre et al., 1997). In studies of

pathologic angiogenesis including the analysis of human

and experimental brain tumors, Ang2 mRNA is also

induced in endothelial cells, although little is known about

mechanisms by which this is regulated (Holash et al., 1999;

Stratmann et al., 1998). In contast, Ang1 is constitutively

scular endothelial cells. TNF-a upregulated transcription rate for Ang2 (2.8

F-a (10 ng/ml) for 1 h (Ang2) or 12 h (Ang1). Nuclei were isolated and in

amount of 32P-labeled RNA probes was hybridized to nylon membrane on

running the ImageQuant software. Differences in loading were normalized

ntage of the vehicle value. Results are meanTSEM from three independent

Fig. 7. Effect of protein synthesis inhibition on regulation of Ang1 by TNF-

a in choroidal microvascular endothelial cells. BCECs were exposed to

vehicle, TNF-a (10 ng/ml), cycloheximide (2 Ag/ml), or a combination of

cycloheximide (2 Ag/ml) and TNF-a (10 ng/ml). Cycloheximide was added

1 h before TNF-a was applied. Total RNAwas extracted from the cells after

8 h of TNF-a stimulation, and Northern analysis was performed as

described for Fig. 2. An Ang1 transcript of 4.4 kB was found in each lane;

an additional transcript of 4.8 kB was found in non-TNF-treated BCEC

following cycloheximide treatment. Results are meanTSEM from three

independent cultures and the representative blots are shown.


expressed primarily by non-endothelial (mesenchymal,

smooth muscle and tumor) cells associated with blood

vessels during physiologic and pathologic angiogenesis

(Papapetropoulos et al., 1999; Thurston et al., 2000).

Unexpectedly, we have found that TNF-a upregulates all

of the key angiogenic growth factors including Ang2, Ang1

and VEGF in microvascular endothelial cells. This indicates

that TNF-a may be one of the important factors that induces

endothelial expression of the angiogenic growth factors. Our

histologic observations have shown that TNF-a colocalizes

with Ang2, Ang1 and VEGF in human choroidal neo-

vascular tissue, further supporting the likely important

interactions between TNF-a and the angiogenic growth

factors. More importantly, the sequential induction of Ang2

and then Ang1 and VEGF in endothelial cells raises the

possibility that endothelial cells may have autocrine loops

controlling the activation state of Tie2 in association with

activation of VEGF receptors along with previously

reported effects of TNF-aon the expression of the Tie2

and Ang1 (Kim et al., 2000; Scott et al., 2002; Willam et al.,

2000; Hashimoto et al., 2004). Further feedback may be

achieved by the anti-inflammatory properties of Ang1,

including its ability to inhibit TNF-a stimulated leukocyte

transmigration (Gamble et al., 2000).

Previous studies have shown that Ang2 mRNA is

induced by basic fibroblast growth factor, VEGF, angioten-

sin II and hypoxia in bovine microvascular endothelial cells

(Mandriota and Pepper, 1998; Oh et al., 1999a; Otani et al.,

2001). Recently, it was shown that TNF-a upregulates Ang2

in human umbilical vein endothelial cells (Kim et al., 2000)

and Ang1 in cultured human synoviocytes (Scott et al.,

2002). The Ang2 mRNA induction by TNF-a that we

observed here was more rapid and transient than that seen

for Ang2 mRNA induction by VEGF in retinal microvas-

cular endothelial cells and by TNF-a in human umbilical

vein endothelial cells. Recent evidence suggests that

angiogenesis requires initial inactivation of, or at least

weakening of, constitutive Tie2 signaling in endothelial

cells (Maisonpierre et al., 1997). It is postulated that Ang2

co-operates with VEGF at the leading edge of vascular

sprouts by blocking the stabilizing or maturing function of

Ang1, thus allowing vessels to revert to, and remain in, a

plastic state where they may be more responsive to a

sprouting signal by VEGF.

In normal eyes, VEGF is expressed by retina and RPE

(Kim et al., 1999). Polarized secretion of VEGF to the

choriocapillaris by RPE and polarized localization of VEGF

receptors on the inner choriocapillaris have been reported,

suggesting that VEGF plays a role in the maintenance of the

choriocapillaris under resting conditions (Blaauwgeers et

al., 1999). In established CNV, VEGF has been shown to be

expressed by various types of cells including vascular cells

and RPE cells (Frank et al., 1996; Ishibashi et al., 1997;

Lopez et al., 1996). Thus, VEGF is expressed and acts on

choroidal endothelial cells throughout the course of CNV. In

the subretinal microenvironment, endothelial cells exposed

to TNF-a may be rapidly shifted to a more plastic state

through autocrine weakening of their Tie2 activation, thus

allowing a better response to such constitutively expressed

or upregulated VEGF.

There is little evidence available about the factors that

upregulate Ang1 expression. Ang1 mRNA expression is

sustained in response to VEGF and hypoxia in retinal

microvascular endothelial cells (Oh et al., 1999a). At the

same time, the Ang1 receptor Tie2 is upregulated by

hypoxia and inflammatory cytokines including TNF-a

(Willam et al., 2000). In human fibroblasts, Ang1 mRNA

is down-regulated by growth factors, including platelet-

derived growth factor, epidermal growth factor and trans-

forming growth factor-h, and hypoxia (Enholm et al., 1997).

In contrast, our study has shown, for the first time, that in

endothelial cells, Ang1 is dramatically upregulated in

response to TNF-a. The induction of Ang1 mRNA was

due to an increase in transcription rate and required de novo

protein synthesis. Furthermore, this delayed induction of

Ang1 was temporally associated with induction of VEGF.

One important role of the upregulated Ang1 in associ-

ation with VEGF would be vessel-maturing activities, which

involve recruitment of vessel-supporting cells, strengthening

of intercellular junctions, and establishment of leakage-


resistant blood vessels (Gamble et al., 2000; Hanahan, 1997;

Thurston et al., 1999). It has been shown that a combination

of Ang1 and VEGF recruits smooth muscle actin-a-positive

cells to vascular walls (Asahara et al., 1998). Co-induction

of endothelial Ang1 and VEGF may exert such vessel-

maturating effects on the sprouting vessels in an autocrine

manner in the microenvironment of active sprouting. It has

been demonstrated that, while VEGF increases vascular

permeability, Ang1 protects against VEGF-induced vascular

leakage and strengthens junctions between endothelial cells

(Gamble et al., 2000; Thurston et al., 2000). Ang1 may also

play a role in protecting newly formed vessels from excess

leakage resulting from VEGF overexpression.

Another important reason for the induction of Ang1 and

VEGF would be to facilitate endothelial survival during

angiogenesis. It has been shown that Ang1 and VEGF can

prevent endothelial apoptotic cell death (Gerber et al., 1998;

Holash et al., 1999; Kwak et al., 1999). Ang1 can also

stabilize endothelial networks (Papapetropoulos et al.,

1999). Actively sprouting vessels may require an increase

in these anti-apoptotic and stabilizing effects. There is

increasing evidence that induction of endothelial apoptosis

is one critical mechanism by which the retina controls

subretinal CNV (Kaplan et al., 1999). The dramatic

upregulation of Ang1 and VEGF in choroidal endothelial

cells may participate in this mechanism as endothelial

surviving factors. Taken together, we speculate that sequen-

tial induction of Ang2 and then Ang1 and VEGF in

endothelial cells may provide precise and stage-appropriate

autocrine and/or paracrine angiogenic signals to endothelial

cells. The upregulation of these endothelial angiogenic

growth factors may account for the discrepancy that TNF-a

has strong angiogenic activities in vivo (Frater-Schroder et

al., 1987; Leibovich et al., 1987; Montrucchio et al., 1994)

even though it inhibits endothelial proliferation and tube

formation and can also induce endothelial apoptosis in vitro

(Frater-Schroder et al., 1987; Polunovsky et al., 1994;

Robaye et al., 1991; Sato et al., 1987).

In this study, we demonstrate for the first time, the

sequential upregulation of angiogenic growth factors in

microvascular endothelial cells by a single inflammatory

mediator. These data provide support for the importance of

TNF-a and Ang1/Ang2 and their autocrine regulatory loops

in neovascularization. Such pathways should be considered

as novel therapeutic targets in the development of treatments

for neovascular diseases.

The authors declare that they have no competing

interests.

Acknowledgements

We thank Dr. George D. Yancopoulos (Regeneron

Pharmaceuticals, Inc., Tarrytown, NY) for providing human

Ang1 and Ang2 cDNA and human Ang1 and Ang2 rabbit

polyclonal antibodies. Christine Spee is acknowledged for

providing choroidal endothelial cells, and Ernesto Barron

for assistance in preparation of the figures.

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