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(Investigative Ophthalmology and Visual Science. 2003;44:4994-5005.)
© 2003 by The Association for Research in Vision and Ophthalmology, Inc.
doi:10.1167/iovs.03-0291
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FGF-2 Antagonizes the TGF-ß1-Mediated Induction of Pericyte {alpha}-Smooth Muscle Actin Expression: A Role for Myf-5 and Smad-Mediated Signaling Pathways

Michael Papetti, Jaleel Shujath, Kathleen N. Riley, and Ira M. Herman

From the Program in Cellular and Molecular Physiology, Department of Physiology, Tufts University School of Medicine, Boston, Massachusetts.


   Abstract
Top
 Abstract
Materials and Methods
 Results
 Discussion
 References
 
PURPOSE. Although the FGF and TGF-ß families are known to play an important role in regulating vascular endothelial and smooth muscle cell behavior, the influence of these matrix-binding growth factors on microvascular pericyte morphogenesis is not well understood. The current study was undertaken to examine the molecular mechanisms that mediate the effects of the endothelium-produced growth regulators FGF-2 and TGF-ß1 on retinal pericyte proliferation and contractile phenotype.

METHODS. Using purified retinal pericytes, a series of assays were implemented, including RT-PCR, DNA binding, immunoprecipitation, electrophoretic mobility shift, and indirect immunofluorescence, in an attempt to elucidate the FGF/TGF-ß1 signaling cascades that mediate retinal microvascular cell growth and contractile phenotype.

RESULTS. Treatment of retinal pericytes with FGF-2 and heparin stimulated nearly a log order increase in proliferation, whereas removal of FGF-2 or addition of TGF-ß1 caused withdrawal from the growth cycle, inducing a smooth-muscle-like contractile phenotype, as indicated by upregulation of {alpha}-smooth muscle actin ({alpha}-SMA). This switch from a growth-potentiated to a growth-arrested state followed induction of the transcriptional regulator myf-5, as well as the nuclear translocation of myf-5 and Smad2.

CONCLUSIONS. Several critical features of the endothelial cell-extracellular matrix-pericyte molecular signaling axis were elucidated in the study that are likely to be responsible for regulating retinal microvascular morphogenesis during normal development, as well as the pathologic angiogenesis accompanying several ocular disorders, including diabetic retinopathy and age-related macular degeneration.

Biochemical analyses and gene cloning studies have clearly demonstrated the importance of the FGF family in fostering endothelial growth and angiogenesis.1 2 3 4 However, although it has been established that FGF-2 is a component of select subendothelial basement membranes in vivo (as well as a component of the endothelial extracellular matrix in vitro),5 its functional role in the matrix remains uncertain.6 Because matrix-bound FGF envelopes populations of quiescent endothelial cells, it has been suggested that it acts as a survival agent that is released after vascular injury.7 8 9 Nevertheless, if FGF-2 acts only in an autocrine or paracrine role to stimulate endothelial proliferation, then it is difficult to explain the exceedingly low rate of endothelial cell turnover in microvascular beds that possess FGF-2-enriched basement membranes.

Beyond playing a role in wound healing, it is possible that matrix-bound FGF-2 serves other important functions during vascular morphogenesis, such as signaling the recruitment, proliferation, and/or local differentiation of mural cells (e.g., vascular smooth muscle cells or pericytes). Indeed, little is known regarding the molecular and cellular cues that govern the nonrandom manner in which pericytes come to populate specific microvessels.10 Despite the demonstration that pericytes can inhibit endothelial cell growth through a cytokine-dependent pathway,11 12 there is no evidence to indicate that pericyte growth or contractile phenotype can be modulated by endothelium-produced or matrix-associated growth regulators such as FGF-2 or TGF-ß1.

Because of the pericyte’s unique position within the microvascular basement membrane13 and because it has been established that endothelial cells deposit FGF-2 in this matrix, we were eager to ascertain whether FGF-2 and other endothelium-produced growth regulators such as TGF-ß specifically influence pericyte proliferation and/or contractile phenotype. Furthermore, we attempted to identify the intracellular downstream effectors that signal to regulate pericyte growth and contractile phenotype.

Herein, we demonstrate that FGF-2 and TGF-ß1 serve antagonistic functions in pericyte growth and myogenic determination. Although FGF-2 markedly stimulates pericyte growth, its removal and/or the addition of TGF-ß1 causes the withdrawal of pericytes from the growth cycle and the induction of a contractile phenotype. This switch from a growth-potentiated to a growth-arrested state temporally follows a dramatic increase in the mRNA and protein levels of the basic helix-loop-helix (bHLH) transcriptional regulator of muscle differentiation myf-5, as well as the nuclear translocation of the TGF-ß phosphosignaling intermediate Smad2. These results help to fill an important gap in our understanding of the molecular mechanisms regulating smooth muscle commitment and myogenic determination in the pericyte lineage.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
Antibodies
For localization, growth neutralization, and immunoprecipitation studies, diethylaminoethyl (DEAE)-fractionated or antigen affinity-purified antibodies were used. Anti-FGF-2 IgG was prepared as described5 ; anti-isoactin IgG ({alpha}-smooth muscle and ß-nonmuscle) were prepared and used as described14 15 ; monoclonal anti-{alpha}-smooth muscle actin ({alpha}-SMA) was purchased from BioGenex (San Ramone, CA); and anti-myf-5 was prepared by injecting full-length glutathione S-transferase (GST)-myf-5 into New Zealand White rabbits.16 Preimmune and immune sera were isolated and characterized by Western blot analysis, immunoabsorption, and immunofluorescence, as previously described.17 For Smad antibodies, peptides generated from the proline-rich linker regions of Smad2, -3, -4, -6, and -7 were synthesized by the Tufts Protein and DNA Sequencing Core Facility (Boston, MA) and cross-linked with 0.1% glutaraldehyde. The cross-linked peptides at 5 mg/mL were dialyzed extensively against PBS and mixed 1:1 with gold adjuvant (Titermax; CytRx Corp., Norcross, GA). Female New Zealand White rabbits were initially injected intramuscularly and intradermally with 500 µg cross-linked peptide. Approximately 1 month after the first injection, the rabbits received a booster injection of 250 µg peptide, and 1 week later the first blood sample was obtained. Further booster injections of 125 µg were administered at approximate 2-week intervals, with respective blood samples collected approximately 1 week afterward. Blood was centrifuged to remove red blood cells, and sera and the resultant immune IgG fractions were tested for reactivity to Smad proteins by Western blot analysis and immunofluorescence. In all rabbits, six to nine boosts were performed, yielding immune sera for collection. The animal protocol adhered to the provisions of the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research.

Pericyte Cultures
Microvascular pericytes were isolated from bovine retinas, as previously described.18 Briefly, retinas were removed from bovine eyes under sterile conditions, and undesirable retinal pigmented epithelial cells were removed by aspiration under a dissecting microscope. Tissue was minced before collagenase digestion, and filtration (Nitex filter; Tetko, Depew, NY) was used to isolate microvessel fragments. Primary isolates were plated in Dulbecco’s modified Eagle’s medium supplemented with 10% calf serum and antibiotics, as previously published.14 18 19 20 Primary pericyte cell cultures were trypsinized and passaged once for all growth and localization studies. Pericyte cultures were characterized, as in the past, for the simultaneous expression of muscle and nonmuscle contractile proteins.14 18 19 20

Growth Studies
For testing the effects of positive growth modulators (FGF-2- or endothelial-cell-conditioned media), as well as negative regulators (TGF-ß1, anti-FGF-2, heparin), first- and second-passage pericytes were plated in triplicate into 24- or 48-well clusters. Initial seeding densities ranged from 5,000 to 14,000 cells per well. Cells growing in the presence or absence of the various test growth regulators received 5 to 12 ng/mL FGF-2 or 0 to 1 ng/mL TGF-ß1 every other day for up to 2-weeks. For heparin sensitivity studies, doses ranged from 1 ng to 100 µg/mL (Elkins-Sinn, Cherry Hill, NJ). Heparin or chondroitin 4-sulfate (Sigma-Aldrich Co., St. Louis, MO) were added alone or in combination with FGF-2 or neutralizing antibodies, which were used at 10 to 100 µg/mL. When effects of anti-FGF-2 on FGF-2-stimulated or endothelial-cell-conditioned media were analyzed, preimmune or absorbed IgG controls were added, either simultaneously or in sequence, with no demonstrable differences. At the end of a given experiment, which was performed at least six times for each condition, triplicate wells were washed, trypsinized, and directly counted (model ZF cell counter; Coulter Electronics, Inc., Hialeah, FL). Results were plotted on computer (KaleidaGraph; Synergy Software, Reading, PA), depicting mean values together with standard errors.

PCR Primers
For reverse transcription-polymerase chain reaction (RT-PCR), the following forward and reverse primers were synthesized based on published (BLAST2; www.ncbi.nlm.nih.gov/blast/ provided in the public domain by the National Center for Biotechnology Information, Bethesda, MD) cDNA sequences as follows: myf-5 (742-bp product), forward, 5'-ATGGACATGATGGACGGCTGCCAG-3', and reverse, 5'-GGTGATCCGATCCACTATGCTGGA-3'; forward (BP 690-1420), 5'-TCCAGCATAGTGGATCGGATCACC-3', and reverse, 5'-TTTATTTATGGTGGAGTTTCCAAAAAGCTC-3'; ß-actin (373-bp product), forward, 5'-TTCTACAATGAGCTGCGTGTGGCT-3', and reverse, 5'-GCTTCTCCTT-AATGTCACGCACGA-3'; {alpha}-SMA (636-bp product), forward, 5'-TGTGAAGAGGAAGACAGCACAGCT-3', and reverse, 5'-GATGGCTGGAAGAGGGTCTCCGGG-3'; myoD (394-bp product), forward, 5'-AGCTCCAACTGCTCCGACGGCATGAA-3', and reverse, 5'-AGGAGAGAATCC-AGTTGATGGAAACA-3'; myogenin (702-bp product), forward, 5'-GG-ACTGGGGACCCCTGAGCAT-3', and reverse, 5'-GTTAAATTCCCTTGCTGGGCT-3'; and herculin or mrf4 (349-bp product), forward, 5'-CCGGCTGGATCAGCAAGAGAA-3', and reverse, 5'-TAAGCTACTAGCCGTTCCGAT-3' and 5'-TTCTCCACCACCTCCTCCACG-3'.

Reverse Transcription-Polymerase Chain Reaction
Ten to 20 µg of total RNA21 that had been purified by CsCl density-gradient centrifugation was treated with 10 U/µL DNase I and RNase inhibitor before phenol-chloroform extraction of RNA. The aqueous phase was collected and precipitated before washing, resuspension in diethyl pyrocarbonate (DEPC) H2O, and reverse transcription. After denaturation at 70°C for 10 minutes, samples derived from pericytes grown under various test conditions were chilled on ice and brought up in an RT cocktail (10 µL cocktail/reaction = 4 µL 5x first-strand buffer, 2 µL 0.1 M dithiothreitol [DTT], 4 µL 2.5 mM dNTP). Samples were warmed at 37°C for 2 minutes and 1.0 µL (200 U) reverse transcriptase (SuperScript II; Invitrogen-Gibco, Gaithersburg, MD) was added before incubation for 60 minutes at 37°C and heat inactivation for 5 minutes at 95°C. For PCR, 4 µL RT reaction was added to 16 µL PCR cocktail and amplified with Taq polymerase for 38 cycles (94°C for 30 seconds, 62°C for 30 seconds, and 72°C for 1 minute, with a 10-minute 72°C extension) using the forward and reverse primers listed herein. The abundance of PCR products primed under these experimental conditions was quantified by quantitative analysis of ethidium bromide-stained bands electrophoresed in agarose gels (IS-1000 Digital Imaging System; Alpha Innotech, San Leandro, CA). All PCR products were sequenced in the Tufts University DNA/Protein Core Facility, to establish sequence identity and to confirm that the correct PCR products were obtained.

Generation of {alpha}-SMA p125 Promoter Fragments
{alpha}-SMA promoter fragments were generated by PCR with DNA polymerase (Pfu turbo; Stratagene, La Jolla, CA). Primers used are as follows: 3'-CCCAAGCTTGGGACTCTGGGTGGGTGGTGTCT and 5'-CGGGGTACCCCGAGGTCCCTATATGGTTGTGT. Base pairs shown in italic denote sites for HindIII (3' primer) and KpnI (5' primer; Invitrogen-Gibco) digestion for subcloning into the HindIII-KpnI site of the pA3 vector. Approximately 50 ng of {alpha}-SMA full-length promoter cloned into a vector (pCAT Basic; Promega, Madison, WI) was mixed with 1x buffer (Pfu; Stratagene), 0.2 mM dNTPs, 100 pM each primer, and 2.5 U DNA polymerase (Pfu turbo; Stratagene) in a total volume of 50 µL. Reactions were cycled for 30 rounds of the following conditions in a thermal cycler (PTC-100; MJ Research, Watertown, MA): 94°C for 1 minute, 60°C for 1 minute, and 72°C for 1 minute. Reaction products were run on 2% agarose/Tris-acetate-EDTA (TAE) gels, and for each construct, a parallel control lane was stained with ethidium bromide. The adjacent band corresponding to the correct size of each PCR fragment was then excised from the unstained lanes, electroeluted, phenol-chloroform extracted, and precipitated with ethanol. PCR products and approximately 500 ng pA3 vector, were then digested with both HindIII and KpnI at 37°C overnight. Digested products were mixed ({alpha}-SMA p125 or {alpha}-SMA p300+pA3), phenol-chloroform extracted, precipitated with ethanol, and resuspended in 15 µL H2O and 4 µL 5x T4 ligase buffer (Invitrogen-Gibco) with 1 U T4 ligase (Invitrogen-Gibco). Ligations were performed at 16°C overnight. Five microliters of ligation product was electroporated into DH5{alpha} Escherichia coli hosts (model 1652076 Gene Pulser; Bio-Rad, Richmond, CA), and positive transformants were selected by standard methods.22

Generation of Single-Stranded {alpha}-SMA 3' Untranslated Region Probe for Northern Analysis
The 1361 rat {alpha}-SMA actin cDNA23 cloned into a plasmid (pBluescript II KS; Stratagene) was digested with DdeI (Promega) and used as a template in an asymmetric PCR reaction that contained approximately 50 ng of DdeI-digested {alpha}-SMA/plasmid DNA; 1.5 mM MgCl2; 1x PCR buffer (Invitrogen-Gibco); 0.2 mM dATP, dTTP, and dGTP; 8 nM dCTP; 50 pM T3 primer; 1 µL 32P dCTP (3000 Ci/mM; ICN, Costa Mesa, CA); and 2.5 U Taq DNA polymerase (Invitrogen-Gibco) in a total volume of 20 µL. Reactions were cycled for 40 rounds in the following conditions: 94°C for 1 minute; 50°C for 2 minutes, and 72°C for 2 minutes. PCR products were purified through probe purification columns (NucTrap; Stratagene), as described by the manufacturer. The final product was a single-stranded ~191-bp fragment from the 3' end of the {alpha}-SMA clone.

Generation of Double-Stranded DNA Probes for EMSA and DNA-Binding Assays
For electrophoretic mobility shift assay (EMSA) and DNA-binding assay, double-stranded constructs were generated as follows. A probe comprising the first 125 bp of the rat {alpha}-SMA promoter was generated by PCR, as described earlier, using DNA polymerase (Pfu; Stratagene) and the primers listed herein, without the overhangs for the HindIII and KpnI sites. The 30-bp promoter fragments were synthesized as complementary single-stranded fragments by the Tufts Protein and DNA Sequencing Core Facility and were annealed by mixing equimolar amounts of each complementary strand, boiling for 5 minutes, and cooling slowly to room temperature.

Northern Blot Analysis
Northern blot analysis was performed as described previously,24 with minor modifications. Pericytes grown in 150-mm plastic dishes in the presence of 10% serum-DMEM were downshifted to 0.01% serum-DMEM for 48 hours. The cells were then treated with 1 ng/mL TGF-ß (Sigma-Aldrich) or 5 ng/mL FGF-2+1 µg/mL heparin for various periods. Total cellular RNA was extracted by density gradient centrifugation of guanidinium isothiocyanate lysates through 5.7 M CsCl cushions22 in a rotor (model SW41; Beckman Instruments, Fullerton, CA) at 24,000 rpm in an ultracentrifuge (model L8-60M; Beckman Instruments). RNA pellets were resuspended in 400 µL DEPC-treated H2O, extracted with phenol-chloroform and ethanol precipitated. For each condition, approximately 1 µg total RNA was run on a 1.2% agarose gel containing 3% formaldehyde. RNA was transferred to nylon membranes (Nytran; Schleicher and Schuell, Keene, NH) overnight in 10x SSC (1.5 M NaCl, 0.15 M Na citrate) by capillary phoresis. RNA was cross-linked to the membranes at optimal setting using a UV cross-linker (model FB-UVXL-1000; Fisher Scientific, Pittsburgh, PA), and the blot was probed for 24 to 48 hours with approximately 106 cpm/mL of a single-stranded 32P-labeled {alpha}-SMA 3' untranslated region (UTR) fragment generated by asymmetric PCR, as described for the rat {alpha}-SMA-specific cDNA. After incubation with probe, the blot was washed at 65°C as follows: two times for 5 minutes each in 2x SSC and 0.1% SDS and one time for 5 minutes in 0.1x SSC. The blot was then air dried at room temperature and exposed to film at -80°C. Cells untreated with cytokine were used as the negative control.

Isolation of Pericyte Nuclei
Nuclei were isolated from bovine retinal pericytes by a modification of the procedure of Workman.25 Nuclei were isolated intact by hypotonic lysis and NP-40 detergent extraction rather than by Dounce (Bellco Glass, Vineland, NJ) homogenization, and nuclear proteins were not extracted with high salt. Briefly, bovine retinal pericytes in 150-mm tissue culture plates (Corning Glass Co., Corning, NY) were cultured in 5% (for EMSA with a 125-bp fragment) or 10% (for EMSA with 30-bp fragments) bovine calf serum-DMEM for 48 hours and then downshifted to 1% serum, with or without 1 ng/mL TGF-ß, for various periods. The cells were iced and washed twice with PBS. Cells were scraped into 5 mL PBS per plate and were pelleted at 4°C and 1500 rpm for 10 minutes in a bench-top centrifuge (RT6000B; Sorvall, Newtown, CT). The cell pellet was gently vortexed 5 seconds, 2 mL per plate hypotonic buffer (10 mM Tris [pH 7.5], 10 mM NaCl, 3 mM MgCl2, 1 mM phenylmethylsulfonyl fluoride, 5 mM E-64 cysteine protease inhibitor, and 2.5 mM Na orthovanadate) was added drop-wise, and the cells were gently vortexed for 10 seconds. Resuspended cells were swollen for 5 minutes on ice, pelleted, and resuspended as before in hypotonic buffer with 1% NP-40. After 10 minutes on ice, the nuclei were pelleted and resuspended as before in hypotonic buffer with 1% NP-40 and were gently vortexed three times for 5 seconds each. Nuclei were pelleted and resuspended at approximately 106/mL in glycerol storage buffer (50 mM Tris [pH 8.4], 5 mM MgCl2, 0.1 mM EDTA, 20% glycerol, 1 mM phenylmethylsulfonyl fluoride, 5 mM E-64, 2.5 mM Na orthovanadate), snap frozen in liquid nitrogen, and stored at -80°C.

Electrophoretic Mobility Shift Assays
EMSAs were performed according to the procedure described by Hautmann et al.26 A double-stranded {alpha}-SMA promoter fragment probe (entire 125-bp fragment of the rat {alpha}-SMA gene promoter or segments spanning -1 to -29, -30 to -59, -60 to -89, -90 to -125, and -202 to -235 [E-box2], or -236 to -269 [E-box1]) was end labeled with 1 µL {gamma}[32P]ATP (3000 Ci/mM) and 1 µL T4 polynucleotide kinase (Invitrogen-Gibco) as described by the manufacturer. End-labeled 32P (~7 x 104 cpm) was incubated with approximately 5000 nuclei from bovine retinal pericytes that had been treated for various lengths of time with 1 ng/mL TGF-ß (Sigma-Aldrich) in EMSA binding buffer (20 mM HEPES [pH 7.9], 50 mM KCl, 0.2 mM EDTA, 0.5 mM DTT, and 2% NP-40) in a total volume of 20 µL 0.7 µg poly dA dT (Amersham Pharmacia Biotech, Piscataway, NJ) was added to eliminate nonspecific protein-DNA interactions. After 20 minutes at room temperature, complexes were separated on 4.5% nondenaturing polyacrylamide gel and visualized by autoradiography. For competition reactions, nuclei were incubated with an approximate 100x molar excess (over labeled DNA fragment) of an unlabeled promoter fragment for 10 minutes before incubation with the labeled one.

Localization Studies
In Vitro.
For localization of FGF-2, FGFR, myf-5, or the isoactins, first- or second-passage pericytes were plated onto glass coverslips in the bottom of 24-well plates or directly onto culture chamber slides (Tissue Tek; Sakura Finetek, Torrance, CA) in DMEM supplemented with serum, FGF-2, or TGF-ß1, as previously published.19 20 Duplicate or triplicate samples from several different retinal isolations (n > 6) were fixed after treatment with various growth factors for different lengths of time (0 minutes to several hours; doses and control ligands are identical with those evaluated for pericyte growth studies). Pericytes were fixed for 5 minutes at room temperature in a freshly prepared solution of 4% formaldehyde in DMEM, permeabilized in 0.1% Triton X-100 buffered with 50 mM HEPES (pH 7.1), 50 mM PIPES (piperazine-N-N'-bis(2-ethanesulfonic acid; pH 6.9), 0.1 mM EGTA, 75 mM KCl, and 1.0 mM MgCl2 for 90 seconds at room temperature before equilibration in PBS. For antibody localization studies, primary antibodies were used at 10 to 50 µg/mL in PBS and applied to cells for 1 hour at room temperature. Secondary antibodies were also applied for 1 hour at room temperature. Texas red, fluorescein, or rhodamine-labeled goat anti-rabbit or mouse IgG (100 µg/mL; Cooper Biomedical, Gaithersburg, MD) were used as previously described.19 20 Quantitative analyses of myf-5 and {alpha}-SMA positive pericyte cultures were performed in a blind fashion on triplicate cultures for each time point over the time course of the experiment. Each set of experiments was repeated at least five times. For myf-5 colocalization, positively stained nuclei were digitally counted and expressed as a percentage of the total nuclear pool, identified by DNA labeling with 4',6'-diamino-2-phenylindole (DAPI). For digital image analysis of immunofluorescence from cultured cells, nuclear fluorescence (for anti-myf-5 localization) and stress-fiber-associated fluorescence ({alpha}-actin IgG localization) were quantified directly on the stage of an inverted fluorescence microscope (Axiovert; Carl Zeiss Meditec, Thornwood, NY) equipped with high-resolution optics (Planapochromat, numeric aperture 1.4; Carl Zeiss Meditec) interfaced with a computer-based microscope imaging workstation (MetaMorph; Universal Imaging Co., Downingtown, PA) and an intensified target video camera (ISIT 66; Dage-MTI, Michigan City, IN). Digitized fluorescence from experimental and control groups was plotted along with standard error on computer (KaleidaGraph; Synergy Software).

Western Blot Analysis.
For Western blot analysis, cell lysates were derived from pericytes treated as described14 27 28 before electrophoresis and transfer to nitrocellulose. After blocking for 1 hour at room temperature in Tris-buffered saline-Tween 20 (TBST) containing 5% fat-free milk, membranes were incubated in antibodies, as previously described.28 Blots were washed extensively in TBST before incubation in secondary antibodies (1:3000 dilution, horseradish-peroxidase [HRP]-conjugated). After another hour of room temperature incubation, blots were washed and developed. Signals were detected using enhanced chemiluminescence, as described by the manufacturer (Super Signal; Pierce Chemical Co, Rockford, IL).


   Results
Top
 Abstract
 Materials and Methods
 Results
Discussion
 References
 
Endothelium-Derived Growth Regulators, Pericyte Growth, and Contractile Phenotype
Earlier work has demonstrated an abundance of pericytes within specific microvascular beds.29 Because there is a 1:1 correspondence between the presence of pericytes and the localization of FGF-2 within the cerebrovascular basement membrane,5 we hypothesized that endothelium-synthesized and matrix-bound FGF-2 plays a pivotal role in either signaling the recruitment of pericyte precursors or fostering pericyte proliferation. Because pericytes occupy the basal lamina along with the microvascular endothelium, presentation of either free FGF-2 or FGF-2-bound heparin sulfate proteoglycan would place the mitogen advantageously for signaling pericyte growth. When purified populations of retinal pericytes were plated in tissue culture medium in the presence of nanogram/milliliter doses of FGF-2, there was a rapid and marked induction of pericyte growth that was more than two times higher than in control cells (Fig. 1) . FGF-2 stimulation of pericyte growth was markedly enhanced when FGF-2 was added in combination with heparin (Fig. 1) , causing a four- to sixfold potentiation of pericyte growth over control cells. This potentiation of FGF-2-induced growth was not observed when other sulfated glycosaminoglycans were added simultaneously with FGF-2. For example, pericytes cultured with identical concentrations of chondroitin 4-sulfate together with FGF-2 showed no significant difference from cells grown with FGF-2 alone. In addition, FGF-2 growth stimulation was completely obliterated when a molar excess of anti-FGF-2 was added simultaneously to pericyte cultures, thus neutralizing the FGF-2-mediated growth response.5 Conversely, addition of nanogram/milliliter doses of TGF-ß1 caused pericyte growth arrest within several hours after its addition to serum-containing media, regardless of whether low (0.5%) or high (5%–10%) concentrations of serum were used (Fig. 1) .



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  		FIGURE 1. FGF-2 and TGF-ß1 modulated pericyte growth and contractile phenotype. Proliferation assays were performed in triplicate 24-well plates over a 4-day time course. FGF-2, FGF-2+heparin, TGF-ß1, or serum alone was added, and the number of cells was counted directly. The mean counts ± SE from triplicate wells and several experiments (n > 6) are shown.

 
Having demonstrated that soluble and matrix-bound growth regulators play an important role in stimulating pericyte proliferation, we recognized that removal of serum and/or FGF-2 from the culture medium might cause pericytes to withdraw from the cell cycle and differentiate, akin to what occurs in the cell culture conditions that have been established for launching skeletal myogenesis.30 31 If similar inductive events were regulating commitment and differentiation in pericytes, we would predict that vascular smooth muscle contractile protein gene expression would be switched on as the encoded contractile protein isoforms accumulated coincident with growth arrest. Indeed, isoelectric focusing and Western blot analysis, as well as immunofluorescence microscopy with isoactin-specific antibodies, revealed that whole-cell extracts and fixed pericyte cultures were devoid of {alpha}-smooth muscle contractile protein isoforms when pericytes were grown in the presence of FGF-2. In contrast, when cells were withdrawn from media containing FGF-2 (not shown) or when pericytes are cultured in the absence of FGF-2, but in the presence of TGF-ß1, a known inducer of {alpha}-SMA expression, smooth muscle contractile protein isoforms accumulated (Fig. 2) . Thus, when pericytes were forced to differentiate by withdrawal of FGF-2 or addition of TGF-ß1, they expressed {alpha}-SMA protein.



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  		FIGURE 2. Addition of TGF-ß1 induced pericyte expression of {alpha}-SMA. Shown are bovine retinal pericytes cultured as described and stained for {alpha}-SMA, after being grown in DMEM with 10% serum (A) or in low serum in the presence of 1 ng/mL TGF-ß (B), which caused an increase in the formation of {alpha}-SMA-containing stress fibers.

 
Effect of TGF-ß1 on the Level of {alpha}-SMA RNA in Retinal Pericytes
To determine the mode(s) of TGF-ß1 regulation of {alpha}-SMA under the aforementioned experimental conditions, we first performed Northern blot analysis on total cellular RNA isolated from bovine retinal pericytes treated with TGF-ß1. A radiolabeled DNA fragment specific for the 3' UTR of {alpha}-SMA32 was used as a probe. TGF-ß1 induced abundant {alpha}-SMA mRNA expression, whereas FGF-2/heparin stimulated little if any accumulation of {alpha}-SMA mRNA (Fig. 3A) . The TGF-ß1-stimulated increase in {alpha}-SMA mRNA plateaued after 4 hours to approximately 2.5-fold over the level observed in unstimulated cells and began to decline to baseline levels after 14 hours (Fig. 3B) .



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  		FIGURE 3. TGF-ß1 induced an increase in {alpha}-SMA mRNA in retinal pericytes. Bovine retinal pericytes were cultured in 10% bovine calf serum/DMEM and were downshifted to 0.01% serum/DMEM for 48 hours. (A) Downshifted pericytes were cultured in the presence of either 1 ng/mL TGF-ß1 or 5 ng/mL FGF-2+1 mg/mL heparin for 3 hours, and Northern blot analysis was performed on approximately 1 mg of RNA from each condition, using a 32P-labeled probe specific for {alpha}-SMA. (B) The cells were then cultured in the presence of 1 ng/mL TGF-ß1 for the indicated lengths of time, and Northern blot analysis was performed, as in (A).

 
Effect of TGF-ß1 on Protein Binding to the {alpha}-SMA Promoter
Because TGF-ß1 stimulated an increase in the steady state levels of {alpha}-SMA mRNA, we hypothesized that TGF-ß1 does this through transcriptional control mechanisms, perhaps by inducing transcription factors to bind to the {alpha}-SMA promoter. Because the first 125 bp of the promoter support TGF-ß1-driven transcription in rat smooth muscle cells,26 this construct was prepared and end labeled with 32P for use as a probe in an EMSA, using nuclei isolated from retinal pericytes grown under control conditions or after stimulation with TGF-ß1. Using this construct, a band shift was evident as early as 0.5 hour after stimulation with TGF-ß1 (Fig. 4A) . This shifted band increased in intensity by 1 hour, then decreased between 4.5 and 14 hours of TGF-ß1 treatment. The shifted complex was specific for the 125-bp promoter fragment, as an approximate 100-fold molar excess of unlabeled probe completely blocked binding (Fig. 4A) .



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  		FIGURE 4. Proteins from TGF-ß1-treated retinal pericytes bind the {alpha}-SMA promoter. (A) Bovine retinal pericytes were cultured in 5% serum+5 ng/mL FGF-2+1 µg/mL heparin for 48 hours. The cells were then washed and treated with 1 ng/mL TGF-ß1 for the times indicated. Nuclear lysates were incubated with the 32P-labeled {alpha}-SMA promoter fragment p125, and DNA complexes were separated on 4.5% polyacrylamide gels by native-PAGE. Where indicated, reactions included a 100x molar excess of unlabeled promoter fragment p125. Arrowhead: free probe; arrow: shifted complex. (B) EMSA was performed as in (A), but pericytes were first incubated in 10% serum+5 ng/mL FGF-2+1 µg/mL heparin, and nuclei from pericytes treated with 1 ng/mL TGF-ß1 for 30 minutes were incubated with the indicated 23P-labeled {alpha}-SMA promoter fragments. Where indicated, reactions included a 100x molar excess of unlabeled specific promoter fragment. Arrowhead: free probe. (C) EMSA was performed as in (B), but reactions included a 100x molar excess of the unlabeled nonspecific promoter fragment indicated. Arrow and arrowhead in (B) and (C) show band shifts created by probes 30 to 59 and 90 to 125, respectively, that were not completely competed by excess E-box1.

 
To further resolve where proteins are binding to the 125-bp {alpha}-SMA promoter fragment, EMSA was performed with 29- to 30-bp oligomers spanning the first 125 bp of the {alpha}-SMA promoter. Because a band shift was seen in an EMSA after 0.5 hour of TGF-ß1 stimulation with the 125-bp promoter construct (Fig. 4A) , end-labeled oligomers were incubated with nuclei from retinal pericytes treated with TGF-ß1 for 30 minutes after downshifting from 10% serum+FGF-2/heparin. Band shifts were evident with each probe used, and each was competitively blocked by an approximate 100-fold molar excess of unlabeled probe (Fig. 4B) . To further test the specificity of the band shifts, the same EMSA was performed, except that the approximate 100-fold molar excess of an unlabeled nonspecific probe was used as the competitor. Most of the band shifts generated by labeled probes spanning -1 to -29 and -60 to -89 were competitively blocked by unlabeled E-box1, indicating that they are probably representative of nonspecific DNA binding (Fig. 4C) . Furthermore, band shifts produced by labeled probes spanning each E-box were competitively blocked by the unlabeled segment spanning -1 to -29. Nevertheless, a few band shifts were not competitively blocked. These include the band shifts using the -90 to –125 probe (Figs. 4B 4C , arrowhead) and the -30 to -59 probe (Figs. 4B 4C , arrow). The -30 to -59 probe encompassed the TGF-ß control element that is necessary for TGF-ß-induced {alpha}-SMA promoter activation of transcription in rat smooth muscle.26 These results suggest that TGF-ß1 induces the binding of transcription factors to segments -30 to -59 and -90 to -125 of the retinal pericyte {alpha}-SMA promoter.

Effect of TGF-ß1 on the Level and Nuclear Translocation of Myf-5 RNA
TGF-ß1 induction of specific {alpha}-SMA promoter band shifts (Fig. 4) suggests that this cytokine stimulates nuclear accumulation of factors necessary for binding and transactivating the {alpha}-SMA promoter. In an effort to learn whether the myoD family of bHLH transcriptional regulators might be the molecular regulators controlling downstream smooth muscle contractile protein gene expression induced by TGF-ß1, we performed RT-PCR with primers sets specific for several of the well-studied skeletal muscle myogenic bHLH transcriptional regulators. RT mixes were prepared from mRNAs isolated from pericytes grown under a variety of conditions, including different concentrations of serum and serum containing FGF-2 (with and without heparin) or TGF-ß1. Results of these RT-PCR experiments revealed conclusively that myf-5 mRNA levels (as seen by RT-PCR) were increased nearly fourfold when pericytes were removed from media containing FGF-2 and cultured in 0.2% serum alone or with TGF-ß1 (Fig. 5) . Further, there was an 18-fold increase in the predicted myf-5 PCR product when pericytes were first stimulated to proliferate under the control of FGF-2 and heparin and then placed in low-serum medium (Fig. 5) . Similarly, TGF-ß1 alone induced myf-5 expression (described later). Neither myf-5 mRNA or protein nor any of the other mRNAs encoding other myoD family members was detected in pericytes growing in the presence of FGF-2 or FGF-2/heparin. Only myf-5, but none of the other myoD family members, was induced after the withdrawal of FGF-2 or the addition of TGF-ß1. Neither vascular endothelial cells nor smooth muscle cells expressed detectable myf-5 (or any myoD family member), regardless of whether mRNAs were isolated from growing or growth-arrested cells. Furthermore, ß-actin mRNA levels were not significantly altered in this set of experimental conditions (Fig. 5) .



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  		FIGURE 5. Myf-5 induction coincides with pericyte growth-arrest. Primers were designed, RNA purified and RT-PCR. Top: myf-5; bottom: ß-actin. Lane 1: 100 bp ladder; lane 2: 10% calf serum; lane 3: FGF-2; lane 4: FGF-2 treatment before growth factor and serum withdrawal (0.2%); lane 5: FGF-2+heparin; lane 6: FGF-2+heparin treatment before growth factor and serum withdrawal; lane 7: endothelial cells; lane 8: vascular smooth muscle cells. Note the induction of the myf-5 PCR product (732 bp) after the withdrawal of FGF-2 or FGF-2 and heparin. Neither endothelial cells nor smooth muscle cells expressed myf-5. ß-Actin control shows 373-bp PCR product. Ethidium bromide staining.

 
Using antibodies specific for myf-5 and {alpha}-SMA, we were interested in learning whether the accumulation of these respective proteins occurs concomitant with growth arrest. Indeed, immunofluorescence microscopy and quantitative image analysis of the anti-myf-5-treated cultures revealed that myf-5 accumulated in pericyte nuclei after the withdrawal of cells from heparin and FGF-2 or the addition of TGF-ß1 (Fig. 6) . Double-immunofluorescence microscopy using the myf-5- and {alpha}-SMA-specific antibodies revealed that the appearance of nuclear myf-5 preceded the emergence of {alpha}-SMA-containing stress fibers and persisted during their emergence (Fig. 7) .



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  		FIGURE 6. Quantitative analysis of myf-5 expression. Anti-myf-5 immunofluorescence was performed (Figs. 8 9) . After fixation and viewing, cells expressing nuclear myf-5 were digitized. Cells that possessed nuclear myf-5 were then expressed as a percentage of the total population counted. Results are from several experiments (n > 5) in which more than 100 cells were counted for each condition, plated in triplicate. Error bars denote SE. Note that FGF-2 and heparin treatment of pericytes blocked nuclear myf-5 accumulation, whereas TGF-ß1 treatment induced myf-5 nuclear localization.

 


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  		FIGURE 7. Myf-5 accumulated in the nucleus concomitant with growth arrest. Pericytes were grown in the presence of 10% serum or FGF-2/heparin. Cells were either maintained under these conditions or washed and changed to TGF-ß1-containing medium. Cells were then fixed and prepared for anti-myf-5 immunofluorescence. When pericytes were cultured in the presence of serum (A) or FGF-2/heparin (B), anti-myf-5 fluorescence was cytoplasmic. However, after the switch from 10% serum to TGF-ß1-containing medium (C), anti-myf-5 fluorescence was nuclear. Confocal imaging confirmed the nuclear localization of myf-5 in pericytes treated with TGF-ß1 (D), and stained simultaneously for myf-5 (green) and {alpha}-SMA (red).

 
Smad2 Translocation into the Nucleus
The TGF-ß receptor is a dimeric complex composed of type I and II serine/threonine kinase subunits. Receptor type II binds TGF-ß and phosphorylates receptor type I, and phosphorylated type I receptor in turn phosphorylates and activates the Smad proteins.33 The type III receptor may facilitate the binding of TGF-ß to the type II receptor.34 The Smad proteins are effectors of the activated TGF-ß receptor. On phosphorylation by activated type I receptor, Smads translocate into the nucleus and may directly bind to target gene promoters to activate gene transcription.35 Therefore, the Smads are likely candidates to bind and activate the {alpha}-SMA gene promoter in response to TGF-ß. To determine whether Smads are involved in TGF-ß1-mediated transactivation of the {alpha}-SMA promoter, we first generated polyclonal antibodies to Smad2, -3, and -4. Specific antibodies were obtained from serum of rabbits that were injected with synthetic polypeptides corresponding to the most nonconserved regions in the Smad hinge region (Fig. 8) . The specificity of these antibodies was assessed by using them to probe Western blot analysis of Smad-overexpressing COS cell lysates. Rabbit antisera to Smad2 recognized a specific band that corresponded to the correct molecular weight of this polypeptide (Fig. 9A) .



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  		FIGURE 8. Schematic of peptides used for rabbit injections to generate polyclonal Smad antibodies. Generalized domain structures of Smad2, -3, -4, -6, and -7 are shown. LVKKLKK represents the conserved nuclear localization signal present in Smad2 and -3, and SS(V/M)/S is the conserved C-terminal site of phosphorylation by TGF-ß receptor type I. MH1 and MH2 refer to the highly conserved N- and C-terminal domains of Smad2, -3, and -4. The divergent linker regions between MH1 and MH2 regions are also shown. Sequences of synthesized peptides used to inject rabbits for antibodies are indicated below each diagram.

 


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  		FIGURE 9. TGF-ß1 induced Smad2 nuclear accumulation in retinal pericytes. (A) Western blot, performed on whole-cell lysates from transfected COS cells, demonstrating the specificity of rabbit anti-Smad2 antibody. (B) Bovine retinal pericytes were seeded onto glass coverslips and continuously grown in the presence of 10% serum. Arrowheads: diffuse cytoplasmic localization of Smad2. (C) Cells were grown in 10% serum and downshifted to 0.01% serum for 48 hours. The downshifted cells were then treated with 1 ng/mL TGF-ß1 for the periods indicated. Arrowheads: punctate perinuclear staining of anti-Smad2; arrow: diffuse cytoplasmic Smad2 localization.

 
To determine the localization of Smad proteins in pericytes, we performed immunofluorescence on formaldehyde-fixed, permeabilized retinal pericytes, using the Smad-specific rabbit IgG. In retinal pericytes asynchronously growing in 10% serum, the antibody to Smad2 showed bright nuclear fluorescence (Fig. 9B) , as well as several cells that exhibited diffuse cytoplasmic staining without nuclear fluorescence (Fig. 9B , arrowheads). This heterogeneity in anti-Smad2 IgG localization suggests that in asynchronously growing cells in high-serum medium, some cells actively cycle, whereas others are committed to the smooth muscle lineage. To test this hypothesis, retinal pericytes were synchronized by downshifting to 0.01% serum for 48 hours, and Smad2 was localized after stimulation with TGF-ß1 for various times. After 5, 10, and 15 minutes of TGF-ß1 treatment, Smad2 appeared as puncta surrounding the nucleus (Fig. 9C , arrowhead) or was diffusely distributed in the cytoplasm (Fig. 9C , arrow, 15 min). After 30 minutes of TGF-ß1 treatment, however, there was an exclusive and robust nuclear fluorescence of Smad2. Therefore, we conclude that endogenous pericyte Smad2 translocates into the nucleus in response to TGF-ß1 treatment of retinal pericytes.


   Discussion
Top
 Abstract
 Materials and Methods
 Results
 Discussion
References
 
The results of these experiments revealed that pericyte growth and differentiation are differentially regulated by antagonistic matrix- and cell-associated signaling cascades. In vivo, these signals are likely to originate from the microvascular endothelium, its associated matrix, or bound growth regulators. The data described herein also help to establish the pivotal role that endothelial cell differentiation plays in modulating microvascular morphogenesis, especially at the level of perivascular cell recruitment and commitment to its smooth muscle phenotype.

Matrix-Bound Growth Regulators, Receptor Kinases, and Pericyte Proliferation
Earlier work has revealed that endothelium-derived extracellular matrix can alter pericyte growth and contractile phenotype.20 36 It is also well established that specific subpopulations of microvascular endothelial cells possess unique arrays of matrix-bound growth regulators (e.g., FGF-2).5 37 38 When these specific populations of vascular endothelial cells are grown in tissue culture, accumulation of matrix-bound FGF-2 occurs, though only after endothelial cell-cell contact, as the cells are withdrawing from the growth cycle.5 This suggests a role for FGF-2 beyond signaling the proliferation of endothelium. One possibility is that FGF-2 also influences growth and differentiation of the closely associated pericytes. These observations are in good agreement with the reported abundance of perivascular cells in the brain,17 19 retina,18 39 40 and kidney5 41 (e.g., in the kidney, mesangial cells associate with glomerular capillary basement membranes enriched in heparin sulfate-associated FGF-25 41 ). Together, these findings suggest that FGF-2 serves as a pericyte mitogen and/or motogen, signaling growth through its receptor tyrosine kinase (RTK). Clearly, FGF-2 and its receptor are colocalized and are seemingly essential for activation of cell cycle progression and pericyte growth. Furthermore, this and other pivotal vascular signaling cascades (e.g., PDGF-bb and TGF-ß1) may be cooperative or antagonistic in mediating pericyte growth and microvascular remodeling during development or in association with vasoproliferative disorders.

Collectively, our findings help to explain the manner in which pericytes are selectively recruited to specific microvascular beds and further identify the parallel growth-promoting RTK signaling pathways responsible for stimulating pericyte proliferation. In addition, because pericyte proliferation occurs when vascular smooth muscle contractile protein gene and protein expression are seemingly repressed,20 it calls into question the usefulness of smooth muscle-specific probes as markers for pericytes when the cells are actively proliferating.42

Myf-5 and Pericyte Myogenic Determination
During the past decade, considerable insights into the molecular mechanisms controlling skeletal myogenesis have been gained.43 44 45 46 Yet, despite this level of molecular understanding with regard to skeletal myogenesis, much less is known about the lineage, recruitment, and molecular switching events that regulate vascular smooth muscle commitment and differentiation. Our results point to a new role for myf-5, the earliest of the bHLH transcriptional regulators to be expressed, in helping to establish the smooth muscle phenotype in retinal pericytes. That myf-5 mRNA and protein levels are markedly induced in pericytes, but not in vascular smooth muscle cells, suggests that the myogenic precursor pools or the molecular signals responsible for seeding the vasculature with its mural components are distinct. This notion of site-specific myf-5 expression in specific perivascular precursor pools is consistent with recent observations regarding the temporal and spatial myf-5 expression patterns observed during murine embryogenesis, when visceral arch and somitic myf-5 expression patterns differ.47 Together with our results, these findings suggest that there are multiple and spatially segregated regulatory domains and pathways that may actively influence myf-5 expression in committed skeletal precursors and now, in pericyte precursors. Whereas a role for myf-5 in regulating perivascular cell commitment has not been considered, our results reflect a need for the reexamination of these transgenic animals, especially at the single-cell level and, more important, within the various embryonic and postnatal microvascular beds. This is especially important when considering the retinal microvasculature, which undergoes extensive postnatal remodeling, both normally and in association with pathologic angiogenesis. Clearly, these studies have helped to divulge important details in the molecular signaling pathways presumed to be responsible for regulating perivascular cell commitment and differentiation, but future work should yield new insights into the embryologic origins and developmental connections that presumably exist among pericyte, smooth muscle, and skeletal muscle precursors.

TGF-ß1 as an Inducer of Pericyte Commitment and Differentiation
TGF-ß1, a member of the TGF gene family, represents the prototype of this multimember family, coordinating a variety of activities, including the regulation of growth, differentiation, and extracellular matrix production.48 49 50 51 52 53 54 55 56 57 Its widespread expression during embryonic development points to its important role in orchestrating epithelium-mesenchyme interactions, angiogenesis, and chondro- and osteogenesis. Only recently, through the targeted disruption of TGF-ß1, have these essential role(s) been more clearly delineated.55 56 58 59 TGF-ß1 signals through two transmembrane serine/threonine kinases (STKI and STKII), which function together and share approximately 40% homology.60 61 62 Each STK possesses an extracellular cysteine-rich domain responsible for fostering ligand binding, a single transmembrane domain, and an intracellular serine/threonine kinase domain responsible for intracellular signaling through cytoplasmic phosphomediators (Smads). Our recent discovery that TGF-ß1 signaling may orchestrate the onset of pericyte differentiation not only indicates the putative importance of STK signaling during retinal microvascular development, but it seems evident that aberrations in TGF-ß1 signal transduction may also be critically involved in regulating pericyte dedifferentiation during the onset of disease, in consideration of the known alterations in the retinal vasculature that occur during diabetes.

We have implicated the Smads as mediators of TGF-ß1-driven {alpha}-SMA promoter transactivation. TGF-ß1 treatment of retinal pericytes drives Smad2 translocation into the nucleus after 30 minutes (Fig. 9) , after which {alpha}-SMA mRNA accumulates (Fig. 3) . Mounting evidence suggests that Smads can bind to DNA and activate transcription from target promoters. Drosophila Mad63 and mammalian Smad3 and Smad4 bind DNA and activate transcription in vitro,64 65 and a Smad3/4 complex binds the consensus sequence GTCTAGAC, which is found in promoters of many TGF-ß-responsive genes.35 66 Specificity in Smad activation of target genes, however, is due to the recruitment of accessory transcription factors by Smads to promoters. Smad2/4 and Smad3/4 form complexes with the winged-helix protein FAST-1 on promoters of Xenopus67 and human68 homeobox genes activated by TGF-ß and activin-like factors.69 70 Smads have also been shown to form complexes with activator protein (AP)-1 to activate transcription of artificial promoters in vitro.71 In addition, Smads can recruit coactivators such as p300 and CREB-binding protein,72 73 as well as corepressors,74 in response to TGF-ß. Therefore, the various effects of TGF-ß are regulated by the cellular context of Smad as it interacts with different proteins to form distinct transcriptional complexes. Perhaps Smads act in concert with myf-5 to induce pericyte {alpha}-SMA gene promoter transactivation and protein expression. In this sense, then, myf-5 would confer specificity in TGF-ß1 signaling to the myogenic lineage-derived pericyte.

The consensus Smad binding element AGAC, defined by Hua et al.,75 is found at -42 of the {alpha}-SMA promoter used in these experiments. Because region -30 to -59 was bound by a nuclear protein(s) from TGF-ß1-treated pericytes (Fig. 4B) and Smad2 translocated from nucleus to cytoplasm in response to TGF-ß1 treatment, it is possible that Smad 2 binds to this element and recruits the accessory proteins necessary for a complete transcriptional unit. We were unsuccessful in supershifting the specific band shift in the EMSA (Fig. 4) with Smad antisera. However, preliminary experiments show that mutation of 2 bp within a Smad consensus DNA binding element35 66 within the first 125 bp of the {alpha}-SMA promoter results in a drastic reduction in the {alpha}-SMA promoter-driven reporter gene expression driven by TGF-ß1 (data not shown). Flanking this element on the 5' end is the 10-bp "TGF-ß-control element" defined by Hautmann et al.26 Characterization of the proteins binding to these regions of the {alpha}-SMA promoter may elucidate the mechanism of TGF-ß1-stimulated {alpha}-SMA gene transcription.

Nevertheless, more work is needed to establish the exact signaling hierarchy between TGF-ß1 signaling and pericyte {alpha}-SMA expression. For example, it is possible that TGF-ß1 signaling intermediates activate both myf-5 and {alpha}-SMA gene transcription. Detailed molecular analysis of these intermediates is likely to reveal the manner in which TGF-ß1 acts as a molecular inducer of the pericyte smooth muscle contractile phenotype.

Antagonistic Signaling Cascades in Modulation of Pericyte Growth and Contractile Phenotype
Our observations suggest that pericyte growth and differentiation are inversely related. A simplistic explanation is that molecular control of either pericyte growth or differentiation is the sum of the positive and negative signals from the vascular basement membrane, with each signaling cascade targeting the regulators of cell cycle progression on the one hand or withdrawal from the cell cycle and differentiation on the other. In fact, there are reports that stimulation of growth by FGF-2 is mediated by phosphorylation of the retinoblastoma susceptibility gene product RB, which is thought to restrict cell cycle progression through late G1.76 77 78 79 Furthermore, based on our finding that myf-5 mediated pericyte {alpha}-smooth muscle contractile protein gene expression (Figs. 5 6 7) , our results suggest that commitment and differentiation occur simultaneously as pericytes withdraw from the growth cycle. Other work in skeletal myogenesis indicates that growth arrest, induction of the CDK inhibitor p21, and skeletal myogenesis ensue after expression of myoD and myf-5.80 This has been proposed as a protective mechanism guarding the differentiated skeletal myocyte against apoptosis. This phenomenon may be evolutionarily conserved, as it is seemingly in place during pericyte myogenesis.

Our findings demonstrate that FGF-2 and heparin promote pericyte growth (Fig. 1) and suppress {alpha}-smooth muscle contractile protein gene expression (Fig. 2) , whereas TGF-ß1 caused growth arrest (Fig. 1) and the induction of the contractile phenotype (Fig. 2) in vitro. These observations are consistent with the robust {alpha}-SMA mRNA and protein expression occurring "downstream" of Smad 2 accumulation in the nucleus (Fig. 9) . Taken together, these findings lend strong credence to the notion that positive and negative endothelium-derived growth regulators act to control the patterned placement of growing versus contractile pericytes within the retinal microcirculatory bed in vivo (Fig. 10) . Furthermore, in consideration of the role that TGF-ß1 signal transduction plays in fostering retinal pericyte differentiation (presumably through induction of smooth muscle gene transcription), it seems likely that other downstream targets are also affected. Because this matrix-driven and kinase-mediated phenotypic modulation could result in local pericyte dedifferentiation, pericytes would be neither contractile nor capable of restraining endothelial cells from reentering the cell cycle in a dedifferentiated state. As such, endothelial cell proliferation and angiogenesis could ensue. Our results predict that perturbations in these antagonistic, matrix-mediated signaling cascades, rather than pericyte death, per se, may serve as the early signals that elicit the recurrent rounds of endothelial proliferation observed during pathologic angiogenesis.



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  		FIGURE 10. Diagram depicting a suggested pathway of FGF-2 and TGF-ß1 induction of pericyte growth and differentiation during microvascular morphogenesis. Pericyte recruitment occurs after endothelial deposition of FGF-2 in the extracellular basement membrane. As pericytes begin contacting endothelial cells, TGF-ß1 becomes activated, therein fostering endothelial growth arrest as well as causing the induction of myf-5 and pericyte smooth muscle contractile protein gene expression. This smooth muscle contractile phenotype maintains the endothelium in its growth-arrested state until aberrations in the extracellular compartment prevent endothelial cell-pericyte contacts. In turn, pericyte and endothelial cell proliferation ensues, launching recurrent rounds of pathologic angiogenesis.

 


   Acknowledgements
 
The authors thank Nicholas Divaris, Justin Glotfelty, and Benson Chen for technical assistance; Amy Yee and Craig Dionne for the gift of antibody reagents, and Pat D’Amore and Anita Roberts for collaborative support and critical review of the manuscript.


   Footnotes
 
Supported in part by National Eye Institute Grant EY09033 (IMH).

Submitted for publication March 20, 2003; revised May 27 and June 27, 2003; accepted August 7, 2003.

Disclosure: M. Papetti, None; J. Shujath, None; K.N. Riley, None; I.M. Herman, None

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Corresponding author: Ira M. Herman, Program in Cellular and Molecular Physiology, Department of Physiology, Tufts University School of Medicine, 136 Harrison Avenue, Boston, MA 02111; ira.herman{at}tufts.edu.


   References
Top
 Abstract
 Materials and Methods
 Results
 Discussion
 References
 

  1. Moses, M, Klagsbrun, M, Shing, Y. (1995) The role of growth factors in vascular cell development and differentiation Int Rev Cytol 161,1-48[Medline][Order article via Infotrieve]
  2. Klagsbrun, M. (1995) Vascular cell growth factors and the arterial wall Haber, E eds. Molecular Cardiovascular Medicine ,63-68 Scientific American Publishing, Inc.
  3. Klagsbrun, M. (1990) The affinity of fibroblast growth factors (FGFs) for heparin: FGF-heparan sulfate interactions in cells and extracellular matrix Curr Opin Cell Biol 2,857-863[CrossRef][Medline][Order article via Infotrieve]
  4. Papetti, M, Herman, IM. (2002) Mechanisms of normal and tumor-derived angiogenesis Am J Physiol 282,C947-C970
  5. Healy, MA, Herman, IM. (1992) Density-dependent accumulation of basic fibroblast growth factor in the subendothelial matrix Eur J Cell Biol 59,56-67[Medline][Order article via Infotrieve]
  6. Nakagawa, S, Pawelek, P, Grinnell, F. (1989) Extracellular matrix organization modulates fibroblast growth and growth factor responsiveness Exp Cell Res 182,572-582[CrossRef][Medline][Order article via Infotrieve]
  7. D’Amore, PA. (1990) Modes of FGF release in vivo and in vitro Cancer Metastasis Rev 9,227-238[CrossRef][Medline][Order article via Infotrieve]
  8. Kostyk, SK, D’Amore, PA, Herman, IM, Wagner, JA. (1994) Optic nerve injury alters basic fibroblast growth factor localization in the retina and optic tract J Neurosci 14,1441-1449[Abstract]
  9. Ku, P, D’Amore, P. (1995) Regulation of basic fibroblast growth factor (bFGF) gene and protein expression following its release from sublethally injured cells J Cell Biochem 58,328-343[CrossRef][Medline][Order article via Infotrieve]
  10. Hirschi, K, Rohovsky, SA, D’Amore, PA. (1998) PDGF, TGF-ß and heterotypic cell-cell interactions mediate the recruitment and differentiation of 10T1/2 cells to a smooth muscle cell fate J Cell Biol 141,805-814[Abstract/Free Full Text]
  11. Antonelli-Orlidge, A, Saunders, KB, Smith, SR, D’Amore, PA. (1989) An activated form of transforming growth factor ß is produced by cocultures of endothelial cells and pericytes Proc Natl Acad Sci USA 86,4544-4548[Abstract/Free Full Text]
  12. Sato, Y, Rifkin, DB. (1989) Inhibition of endothelial cell movement by pericytes and smooth muscle cells: activation of a latent transforming growth factor-beta 1-like molecule by plasmin during co-culture J Cell Biol 109,309-315[Abstract/Free Full Text]
  13. Kuwabara, T, Cogan, DG. (1963) Mural cells of retinal capillaries Arch Ophthalmol 69,492-502
  14. DeNofrio, D, Hook, TC, Herman, I. (1989) Functional sorting of actin isoforms in microvascular pericytes J Cell Biol 109,191-202[Abstract/Free Full Text]
  15. Hoock, TC, Newcomb, PM, Herman, IM. (1991) Beta actin and its mRNA are localized at the plasma membrane and the regions of moving cytoplasm during cellular response to injury J Cell Biol 112,653-664[Abstract/Free Full Text]
  16. Shih, HH, Tevosian, SG, Yee, AS. (1998) Regulation of differentiation by HBP1, a target of the retinoblastoma protein Mol Cell Biol Aug 18,4732-4743
  17. Herman, IM, Jacobson, S. (1988) In situ analysis of microvascular pericytes in hypertensive rat brains Tissue Cell 20,1-12[CrossRef][Medline][Order article via Infotrieve]
  18. Herman, IM, D’Amore, PA. (1985) Microvascular pericytes contain muscle and nonmuscle actins J Cell Biol 101,43-52[Abstract/Free Full Text]
  19. Herman, IM, Newcomb, PM, Coughlin, JE, Jacobson, S. (1987) Characterization of microvascular cell cultures from normotensive and hypertensive rat brains: pericyte-endothelial cell interactions in vitro Tissue Cell 19,197-206[CrossRef][Medline][Order article via Infotrieve]
  20. Newcomb, P, Herman, IM. (1993) Pericyte growth and contractile phenotype: modulation by endothelial-synthesized matrix and comparison with aortic smooth muscle cells J Cell Physiol 155,385-393[CrossRef][Medline][Order article via Infotrieve]
  21. Chirgwin, JM, Przbyla, AE, MacDonald, RJ, Rutter, WJ. (1979) Isolation of biologically active ribonucleic acid from sources enriched for ribonuclease Biochemistry 18,5294-5299[CrossRef][Medline][Order article via Infotrieve]
  22. Sambrook, J, Fritsch, EF, Maniatis, T. (1989) Nolan, C eds. Molecular Cloning: A Laboratory Manual 2nd ed. Cold Spring Harbor Press Plainview, NY.
  23. McHugh, KM, Lessard, JL. (1988) The nucleotide sequence of a rat vascular smooth muscle alpha-actin cDNA Nucleic Acids Res 16,4167[Free Full Text]
  24. Hoock, TC, Newcomb, PM, Herman, IM. (1991) Beta actin and its mRNA are localized at the plasma membrane and the regions of moving cytoplasm during the cellular response to injury J Cell Biol 112,653-664
  25. Workman, JL. (1996) Preparation of nuclear and cytoplasmic extracts from mammalian cells Ausubel, FM Brent, R Kingston, REet al eds. Current Protocols in Molecular Biology ,12.11.11-12.11.19 John Wiley and Sons New York, NY.
  26. Hautmann, MB, Madsen, CS, Owens, GK. (1997) A transforming growth factor beta (TGFbeta) control element drives TGFbeta-induced stimulation of smooth muscle alpha-actin gene expression in concert with two CArG elements J Biol Chem 272,10948-10956[Abstract/Free Full Text]
  27. Shuster, CB, Herman, IM. (1995) Indirect association of ezrin with F-actin: isoform specificity and calcium sensitivity J Cell Biol 128,837-848[Abstract/Free Full Text]
  28. Shuster, CB, Lin, AY, Nayak, R, Herman, IM. (1996) Betacap73: a novel beta actin-specific binding protein Cell Motil Cytoskeleton 35,175-187[CrossRef][Medline][Order article via Infotrieve]
  29. Herman, IM, Jacobson, S. (1988) In situ analysis of microvascular pericytes in hypertensive rat brains Tissue Cell 20,1-12
  30. Dias, P, Dilling, M, Houghton, P. (1994) The molecular basis of skeletal muscle differentiation Semin Diagn Pathol 11,3-14[Medline][Order article via Infotrieve]
  31. Buckingham, M. (2003) How the community effect orchestrates muscle differentiation Bioessays 25,13-16[CrossRef][Medline][Order article via Infotrieve]
  32. Newcomb, PM, Herman, IM. (1993) Pericyte growth and contractile phenotype: modulation by endothelial-synthesized matrix and comparison with aortic smooth muscle J Cell Physiol 155,385-393
  33. Nakao, A, Imamura, T, Souchelnytskyi, S, et al (1997) TGF-beta receptor-mediated signalling through Smad2, Smad3 and Smad4 EMBO J 16,5353-5362[CrossRef][Medline][Order article via Infotrieve]
  34. Blobe, GC, Liu, X, Fang, SJ, How, T, Lodish, HF. (2001) A novel mechanism for regulating transforming growth factor beta (TGF-beta) signaling: functional modulation of type III TGF-beta receptor expression through interaction with the PDZ domain protein, GIPC J Biol Chem 276,39608-39617[Abstract/Free Full Text]
  35. Massague, J, Wotton, D. (2000) Transcriptional control by the TGF-beta/Smad signaling system EMBO J 19,1745-1754[CrossRef][Medline][Order article via Infotrieve]
  36. Herman, IM, Castellot, JJ. (1987) Regulation of vascular smooth muscle cell growth by endothelial-synthesized extracellular matrices Arteriosclerosis 7,463-469[Abstract/Free Full Text]
  37. Padua, RR, Kardami, E. (1993) Increased basic fibroblast growth factor (bFGF) accumulation and distinct patterns of localization in isoproterenol-induced cardiomyocyte injury Growth Factors 8,291-306[Medline][Order article via Infotrieve]
  38. Gao, H, Hollyfield, JG. (1992) Basic fibroblast growth factor (bFGF) immunolocalization in the rodent outer retina demonstrated with an anti-rodent bFGF antibody Brain Res 585,355-360[CrossRef][Medline][Order article via Infotrieve]
  39. Herman, IM. (1993) Controlling the expression of smooth muscle contractile protein isoforms: a role for the extracellular matrix? Am J Respir Cell Mol Biol 9,3-4
  40. Herman, IM. (1993) Microvascular pericytes in development and disease Pardridge, WM eds. The Blood-Brain Barrier ,127-135 Raven Press New York.
  41. Schlondorff, D. (1987) The glomerular mesangial cell: an expanding role for a specialized pericyte FASEB J 1,272-281[Abstract]
  42. Lindahl, P, Johansson, BR, Leveen, P, Betsholtz, C. (1997) Pericyte loss and microaneurysm formation in PDGF-B-deficient mice Science 277,242-245[Abstract/Free Full Text]
  43. Caruso, M, Martelli, F, Giordano, A, Felsani, A. (1993) Regulation of myoD gene transcription and protein function by the transforming domains of E1A oncoprotein Oncogene 8,267-278[Medline][Order article via Infotrieve]
  44. Wright, WE, Sassoon, DA, Lin, VK. (1989) Myogen, a factor regulating myogenesis, has a domain homologous to MyoD Cell 4,607-617
  45. Rudnicki, M, Braun, T, Hinuma, S, Jaenisch, R. (1993) MyoD or Myf 5 is required for formation of skeletal muscle Cell 75,1351-1359[CrossRef][Medline][Order article via Infotrieve]
  46. Weintraub, H. (1993) The myoD family and myogenesis: redundancy, networks and thresholds Cell 75,1241-1244[CrossRef][Medline][Order article via Infotrieve]
  47. Patapoutian, A, Miner, JH, Lyons, GE, Wold, B. (1993) Isolated sequences from the linked Myf-5 and MRF4 genes drive distinct patterns of muscle-specific expression in transgenic mice Development 118,61-69[Abstract]
  48. Verbeek, MM, Otte-Holler, I, Wesseling, P, Ruiter, DJ, de Waal, RM. (1994) Induction of alpha-smooth muscle actin expression in cultured human brain pericytes by transforming growth factor-beta 1 Am J Pathol 144,372-382[Abstract]
  49. Desmouliere, A, Geinoz, A, Gabbiani, F, Gabbiani, G. (1993) Transforming growth factor-beta 1 induces alpha-smooth muscle actin expression in granulation tissue myofibroblasts and in quiescent and growing cultured fibroblasts J Cell Biol 122,103-111[Abstract/Free Full Text]
  50. Kurosaka, H, Kurosaka, D, Kato, K, Mashima, Y, Tanaka, Y. (1998) Transforming growth factor-beta 1 promotes contraction of collagen gel by bovine corneal fibroblasts through differentiation of myofibroblasts Invest Ophthalmol Vis Sci 39,699-704[Abstract/Free Full Text]
  51. Sporn, MB, Roberts, AB. (1992) Transforming growth factor-ß:recent progress and new challenges J Cell Biol 119,1017-1021[Free Full Text]
  52. Roberts, A, Sporn, M. (1996) Transforming growth factor-ß Clark, RAF eds. Molecular and Cell Biology of Wound Repair ,275-308
  53. Wakefield, LM, Roberts, AB. (2002) TGF-beta signaling: positive and negative effects on tumorigenesis Curr Opin Genet Dev 12,22-29[CrossRef][Medline][Order article via Infotrieve]
  54. Bonyadi, M, Rusholme, SAB, Cousins, FM, et al (1997) Mapping of a major genetic modifier of embryonic lethality in TGFß1 knockout mice Nat Genet 15,207-211[CrossRef][Medline][Order article via Infotrieve]
  55. Bottinger, E, Letterio, J, Roberts, A. (1997) Biology of TGF-ß in knockout and transgenic mouse models Kidney Int 51,1355-1360[Medline][Order article via Infotrieve]
  56. Heine, UI, Munoz, EF, Flanders, KC, et al (1987) Role of transforming growth factor-ß in the development of the mouse embryo J Cell Biol 105,2861-2876[Abstract/Free Full Text]
  57. Markowitz, S, Roberts, A. (1996) Tumor suppressor activity of the TGF-ß pathway in human cancers Cytokine Growth Factor Rev 7,93-102[CrossRef][Medline][Order article via Infotrieve]
  58. Roberts, WG, Palade, GE. (1995) Increased microvascular permeability and endothelial fenestration induced by vascular endothelial growth factor J Cell Sci 108,2369-2379[Abstract]
  59. Kulkarini, AB, Huh, CG, Becker, D, et al (1993) Transforming growth factor-beta 1 null mutation in mice causes excessive inflammatory response and early death Proc Natl Acad Sci USA 90,770-774[Abstract/Free Full Text]
  60. Okadome, T, Yamashita, H, Fransen, P, Moren, A, Heldin, CH, Miyazono, K. (1994) Distinct roles of the intracellular domains of TGFb type I and type II receptors in signal transduction J Biol Chem 269,30753-30756[Abstract/Free Full Text]
  61. Wang, X-J, Greenhalgh, DA, Bickenbach, JR, et al (1997) Expression of a dominant-negative type II transforming growth factor ß (TGF-ß) receptor in the epidermis of transgenic mice blocks TGF-ß-mediated growth inhibition Proc Natl Acad Sci USA 94,2386-2391[Abstract/Free Full Text]
  62. Attisano, L, Carcamo, J, Ventura, F, Weis, F, Massague, J, Wrans, J. (1993) Identification of human activin and TGFß type I receptors that form heteromeric kinase complexes with type II receptors Cell 75,671-680[CrossRef][Medline][Order article via Infotrieve]
  63. Kim, S, Ip, HS, Lu, MM, Clendenin, C, Parmacek, MS. (1997) A serum response factor-dependent transcriptional regulatory program identified distinct smooth muscle cell sublineages Mol Cell Biol 17,2266-2278[Abstract/Free Full Text]
  64. Dennler, S, Itoh, S, Vivien, D, ten Dijke, P, Huet, S, Gauthier, JM. (1998) Direct binding of Smad3 and Smad4 to critical TGF beta-inducible elements in the promoter of human plasminogen activator inhibitor-type 1 gene EMBO J 17,3091-3100[CrossRef][Medline][Order article via Infotrieve]
  65. Nagarajan, RP, Zhang, J, Li, W, Chen, Y. (1999) Regulation of Smad7 promoter by direct association with Smad3 and Smad4 J Biol Chem 274,33412-33418[Abstract/Free Full Text]
  66. Zawel, L, Dai, JL, Buckhaults, P, et al (1998) Human Smad3 and Smad4 are sequence-specific transcription activators Mol Cell 1,611-617[CrossRef][Medline][Order article via Infotrieve]
  67. Chen, PL, Riley, DJ, Chen-Kiang, S, Lee, WH. (1996) Retinoblastoma protein directly interacts with and activates the transcription factor NFIL6 Proc Natl Acad Sci USA 93,465-469[Abstract/Free Full Text]
  68. Zhou, Z, Wang, J, Han, X, Zhou, J, Linder, S. (1998) Up-regulation of human secreted frizzled homolog in apoptosis and its down-regulation in breast tumors Int J Cancer 78,95-99[CrossRef][Medline][Order article via Infotrieve]
  69. Pardali, E, Xie, XQ, Tsapogas, P, et al (2000) Smad and AML proteins synergistically confer transforming growth factor beta1 responsiveness to human germ-line IgA genes J Biol Chem 275,3552-3560[Abstract/Free Full Text]
  70. Hua, X, Liu, X, Ansari, DO, Lodish, HF. (1998) Synergistic cooperation of TFE3 and smad proteins in TGF-beta-induced transcription of the plasminogen activator inhibitor-1 gene Genes Dev 12,3084-3095[Abstract/Free Full Text]
  71. Zhang, Y, Feng, XH, Derynck, R. (1998) Smad3 and Smad4 cooperate with c-Jun/c-Fos to mediate TGF-beta-induced transcription Nature 394,909-913[CrossRef][Medline][Order article via Infotrieve]
  72. Feng, XH, Zhang, Y, Wu, RY, Derynck, R. (1998) The tumor suppressor Smad4/DPC4 and transcriptional adaptor CBP/p300 are coactivators for smad3 in TGF-beta-induced transcriptional activation Genes Dev 12,2153-2163[Abstract/Free Full Text]
  73. Pouponnot, C, Jayaraman, L, Massague, J. (1998) Physical and functional interaction of SMADs and p300/CBP J Biol Chem 273,22865-22868[Abstract/Free Full Text]
  74. Wotton, D, Lo, RS, Lee, S, Massague, J. (1999) A Smad transcriptional corepressor Cell 97,29-39[CrossRef][Medline][Order article via Infotrieve]
  75. Hua, X, Miller, ZA, Wu, G, Shi, Y, Lodish, HF. (1999) Specificity in transforming growth factor beta-induced transcription of the plasminogen activator inhibitor-1 gene: interactions of promoter DNA, transcription factor muE3, and Smad proteins Proc Natl Acad Sci USA 96,13130-13135[Abstract/Free Full Text]
  76. Olwin, BB, Rapraeger, AC. (1992) Repression of myogenic differentiation by aFGF, bFGF, and k-FGF is dependent on cellular heparan sulfate J Cell Biol 118,631-639[Abstract/Free Full Text]
  77. Rapraeger, AC, Krufka, A, Olwin, BB. (1991) Requirement of heparan sulfate for bFGF-mediated fibroblast growth and myoblast differentiation Science 252,1705-1708[Abstract/Free Full Text]
  78. Novitch, BG, Mulligan, GJ, Jacks, T, Lassar, AB. (1996) Skeletal muscle cells lacking the retinoblastoma protein display defects in muscle gene expression and accumulate in S and G2 phases of the cell cycle J Cell Biol 135,441-456[Abstract/Free Full Text]
  79. Goodrich, D, Wang, NP, Qian, YW, Lee, EYHP, Lee, WH. (1991) The retinoblastoma gene product regulates progression through the G1 phase of the cell cycle Cell 67,293-302[CrossRef][Medline][Order article via Infotrieve]
  80. Halavey, O, Novitch, B, Spicer, DB, et al (1995) Correlation of terminal cell cycle arrest of skeletal muscle with induction of p21 by myoD Science 267,1018-1021[Abstract/Free Full Text]



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