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. 2003 Apr 4;36(2):75–86. doi: 10.1046/j.1365-2184.2003.00262.x

Stromal cell‐derived factor 1α (SDF‐1α) induces gene‐expression of early growth response‐1 (Egr‐1) and VEGF in human arterial endothelial cells and enhances VEGF induced cell proliferation

Thomas Neuhaus 1, Sebastian Stier 1, Gudrun Totzke 2, Elisabeth Gruenewald 1, Stefan Fronhoffs 1, Agapios Sachinidis 3, Hans Vetter 1, Yon D Ko 1,✉
PMCID: PMC6496392  PMID: 12680875

Abstract

Abstract. Stromal cell‐derived factor‐1 (SDF‐1), mainly known as a chemotactic factor for haematopoietic progenitor cells, also provides angiogenetic potency. Since the intracellular signalling of SDF‐1‐induced neovascularization remains unclear, we studied in human umbilical arterial endothelial cells (HUAEC) the influence of SDF‐1α on induction of the genes of early growth response‐1 (Egr‐1) and VEGF, as well as the activation of extracellular regulated kinases (ERK) 1/2, which are all known to be involved in endothelial cell proliferation. We found a time‐dependent induction of Egr‐1 and VEGF mRNA expression and phosphorylation of ERK1/2 by SDF‐1α. Furthermore, we demonstrated that Egr‐1 expression is dependent on ERK 1/2 activation. Finally, we tried to confirm the relevance of the induced gene expression by detecting the [3H]thymidine incorporation as a marker for cell proliferation in HUAEC after stimulation with SDF‐1α alone or together with VEGF. This particular test showed, that SDF‐1α alone has no effect, but is able to significantly enhance VEGF induced DNA synthesis. In summary, SDF‐1α is involved in different steps of endothelial cell proliferation, but, since Egr‐1 and VEGF offer different functions, it may also play a so far undefined role on other conditions of the endothelium.

INTRODUCTION

Stromal cell‐derived factor‐1 (SDF‐1) is a CXC‐chemokine and is a ligand for the receptor CXCR‐4 (Loetscher et al. 1994; Bleul et al. 1996). Two isoforms are described: SDF‐1α and SDF‐1β. Both are encoded by one gene and develop from different splicing (Shirozu et al. 1995). SDF‐1 was first characterized as a pre‐B cell growth‐stimulating factor in mouse bone marrow, but as a chemotactic factor for CD34+ human progenitor cells (Aiuti et al. 1997; Naiyer et al. 1999) it is also involved in the homing of haematopoietic stem cells (Peled et al. 2000).

While the interaction of SDF‐1 with different haematopoietic cells is well characterized, only very little is known about the influence of SDF‐1 on endothelial cells (EC), though they also express the CXCR‐4 receptor (Volin et al. 1998) and show the haematopoietic surface marker CD34. SDF‐1α leads to neovascularization in vivo (Salcedo et al. 1999; Mirshahi et al. 2000) and the CXCR‐4 receptor is essential for vascularization of the gastrointestinal tract (Tachibana et al. 1998); however, the intracellular pathways of SDF in EC, which are related to angiogenesis, are not well defined.

Thus, we focused in our study on genes or proteins, which are known to be coupled with endothelial cell proliferation. Regarding the signalling cascade, this is true for the extracellular signal‐regulated kinases (ERK)1/2 (Yu & Sato 1999) and, looking downstream, for the transcription‐factor early growth response 1 (Egr‐1) (Biesiada et al. 1996; Hofer et al. 1996). Egr‐1 expression is associated with endothelial injury (Khachigian et al. 1997; Silverman & Collins 1999) and can be activated by growth factors like basic fibroblast growth factor (bFGF) (Ko et al. 1995). In addition, Egr‐1 up‐regulates the expression of platelet‐derived growth factor (PDGF), the VEGF‐receptor Flt‐1 (Khachigian et al. 1996; Vidal et al. 2000) and it triggers the expression of VEGF itself (Yan et al. 2000).

Since VEGF is the key growth factor inducing neovascularization (Neufeld et al. 1999), we examined a possible linkage between SDF‐1α and VEGF. Recently a SDF‐1α induced VEGF expression and production was described in microvascular endothelial cells (Mirshahi et al. 2000), but this group failed to demonstrate a proliferative potency of SDF‐1α in those cells. We repeated these experiments in HUAEC and furthermore analysed the influence of SDF‐1α in combination with VEGF on cell proliferation, using the [3H]thymidine incorporation.

MATERIALS AND METHODS

Materials

SDF‐1α was obtained from R & D systems (Minneapolis, MN). Fetal calf serum (FCS) and Dispase II was obtained from Roche Diagnostics (Mannheim, Germany). Collagenase I and TRI‐reagent were obtained from Sigma Chemical (Deisenhofen, Germany). Moloney murine leukaemia virus (MMLV) reverse transcriptase was obtained from Life Technologies (Karlsruhe, Germany) and Taq polymerase, random primers and dNTP's were obtained from Perkin Elmer (Weiterstadt, Germany). RNA‐Guard and all chemicals for oligonucleotide synthesis were obtained from Pharmacia (Freiburg, Germany). ERK1/2 antibodies, p38, the secondary antibody and the p38 positive control were obtained from New England BioLabs (Beverly, MA). The ECL Western blotting detection system was obtained from Amersham (Little Chalfont, UK).

Culture and stimulation of human umbilical arterial endothelial cells (HUAEC)

Endothelial cells were isolated from human umbilical cord arteries, cultured on human fibronectin‐coated culture dishes in Medium 199 supplemented with 20% v/v fetal calf serum, 10 µg/mL heparin and 30 µg/mL crude endothelial cell growth factor, and characterized as described in (Ko et al. 1995). Only umbilical cords from subjects who had a normal pregnancy and birth were used. For gene‐expression studies, confluent HUAEC of the third or fourth passage were washed twice with PBS and then (for the purpose of starving) exposed for 4 h to Medium 199 without serum or growth factor substitution. For Western‐blotting, EC of the third passage were transferred on 6‐well‐plates, grown until confluence and then exposed to starvation‐medium for 4 h. The cells were then stimulated with SDF‐1α for various time intervals.

RNA‐Extraction

Total RNA was extracted from cells with TRI reagent according to the manufacturer's protocol. RNA quantification was performed by spectrophotometer. The integrity of RNA was analysed by electrophoresis of 4 µg of total RNA on a 1% (w/v) agarose gel stained with ethidium bromide.

Primers

In the case of Egr‐1, the following primers were used: 5′‐primer: 5′‐cagcagtcccatttactcag‐3′, 3′‐primer: 5′‐gactggtagctggtattg‐3′. The resulting PCR product had a length of 345 bp. For VEGF, the primers were: 5′‐primer: gcagaatcatcacgaagtgg‐3′, 3′‐primer: 5′‐gcaacgcgagtctgtgtttttg‐3′, resulting in a PCR product with a length of 414 bp. For both primers, Southern blot analysis as a control of specificity and the determination of the exponential phase of the reverse transcriptase/PCR (RT/PCR) were performed as previously described (Ko et al. 1995; Ko et al. 1999). For standardization, each RT/PCR experiment was followed by a PCR for glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) (5′‐primer, 5′‐gccaaaagggtcatcatctc‐3′; 3′‐primer, 5′‐gtagaggcagggatgatgttc‐3′), a constitutively expressed gene in HUAEC.

Reverse transcription/polymerase chain reaction (RT/PCR)

A 4‐µg portion of total RNA was incubated with moloney murine leukaemia virus (MMLV) reverse transcriptase (200 U/µL) for 5 min at 25 °C, 5 min at 30 °C, 90 min at 37 °C and 5 min at 95 °C in a total reaction volume of 40 µL containing 1× RT‐buffer (50 mm Tris‐HCl, pH 8.3, 75 mm KCl, 3 mm MgCl2), 10 mm dithiothreitol, 0.5 mm of each dNTP, 50 U of RNA‐guard and 100 pmol of random hexamer primer. A 4‐µL aliquot of the RT sample was used for PCR‐reaction, which contained (in a final volume of 50 µL) 1× PCR buffer (10 mm Tris‐HCl (pH 8.3), 50 mm KCl, 1.5 mm MgCl2 and 0.001% gelatin), 0.2 mm concentration of each dNTP, 20 pmol of each primer and 2.5 units of Taq polymerase. The samples were covered with 40 µL of mineral oil and PCR cycles were then performed in an automated DNA thermal cycler (model 480: Perkin Elmer, Weiterstadt, Germany) with the following temperature profile: denaturation at 94 °C for 1 min, primer annealing at 60 °C for 1 min, and primer extension at 72 °C for 1.5 min After an initial denaturation step (94 °C for 5 min), the cycle was repeated 33 times using the VEGF primer, 27 times using the Egr‐1 primer and 25 times for the GAPDH primer. The last cycle was followed by a final extension step of 20 min at 72 °C. Portions of 20 µL of each sample were electrophoresed on a 1.5% (w/v) agarose gel and stained with ethidium bromide. Each PCR experiment was performed with a negative control (lacking MMLV reverse transcriptase) to detect DNA contamination. In order to assess the product size, φX‐174‐RF DNA HaeIII digest was used as a marker.

Western‐Blotting

Cells were lysed with SDS sample buffer containing 20 mm Tris‐HCl, pH 7.4, 50 mm NaCl, 50 mm NaF, 10 mm EDTA, 20 mm napyrophosphate, 1 mm sodium orthovanadate, and 1% Triton X‐100. Aliquots were used for protein determinations using the Bio‐Rad protein assay as previously described (Bradford 1976). Then, 0.1% bromphenol blue (w/v) was added to the aliquots. Thirty µg of protein were analysed by SDS‐PAGE in a 10% acrylamide gel with a thickness of 0.75 mm using the Mini Gel Protein system (Bio‐Rad). Proteins were transferred overnight to a cellulose nitrate membrane (Schleicher and Schuell, Dassel, Germany) by 100 mA with a buffer containing 25 mm Tris‐base, 192 mm glycin and 20% methanol, pH 8.3. The protein transfer was checked using Ponseau S. The membrane was then washed three times with 50 mm Tris‐Cl and 150 mm NaCl, pH 7.5. Saturation was performed with 50 mm Tris‐Cl, 150 mm NaCl, pH 7.5, containing 2% BSA (w/w), and 0.1% Tween 20. The sheets were incubated overnight at 8 °C with the various antibodies. Following six rinses with washing buffer containing 50 mm Tris‐Cl, 150 mm NaCl, pH 7.5 and 0.1% Tween 20, sheets were incubated for 1 h at room temperature in saturation buffer containing the antirabbit secondary antibody. Again, the sheets were washed six times. For detection, the ECL Western blotting detection system by Amersham was used.

Determination of the DNA Synthesis in HUAEC

The effect of VEGF and/or SDF‐1α on [3H]thymidine incorporation into cell DNA was assessed as performed previously (Sachinidis et al. 1995). HUAEC were seeded in 24‐well culture plates and grown to 70% confluence. The medium was then replaced by serum‐free Medium 199. Cultures were then exposed to 50 ng/mL VEGF and/or SDF‐1α 1, 10, 50 and 100 ng/mL for 20 h before 3 µCi/mL [3H]thymidine were added to the serum‐free medium. Four hours later, experiments were terminated by aspirating the medium and subjecting the cultures to sequential washes with PBS containing 1 mm CaCl2, 1 mm MgCl2, 10% trichloroacetic acid, and ethanol:ether (2 : 1, v/v). Acid‐insoluble [3H]thymidine was extracted into 250 µL/dish 0.5 m NaOH, and 100 µL of this solution were mixed with 5 mL scintillant (Ultimagold; Packard, Meriden, CT) and quantified using a liquid scintillation counter, model Beckman LS 3801 (Düsseldorf, Germany). Fifty microliters of the residual solution were prepared for the determination of protein using the Bio‐Rad protein assay as described previously (Bradford 1976).

Densitometric analysis

Densitometric analysis was performed on a two‐dimensional scanning densitometer (BIOMETRA, Göttingen, Germany) using the ‘ScanPack’‐software version 14.1 A 27. The ethidium bromide stained agarose gels were photographed. The densitometric results of gene‐expression were standardized to that of GAPDH expression from the same reverse‐transcribed mRNA sample.

Statistical analysis

Data were presented as mean ± standard deviation. Statistical differences were determined using analysis of variance with repeated measures. P‐values less than 0.05 were considered statistically significant.

RESULTS

Induction of Egr‐1 and VEGF expression

After 4 h in starvation, EC were stimulated with 50 ng/mL of SDF‐1α for different time intervals. SDF‐1α induced a time‐dependent increase in Egr‐1 mRNA levels (Fig. 1). Egr‐1 mRNA expression was highest at 60 min and returned to basal levels 2 h after stimulation. In contrast to Egr‐1, VEGF mRNA was found to be expressed in unstimulated EC. However, stimulation of EC with SDF‐1α, 50 ng/mL, resulted in an increase of VEGF mRNA above baseline levels (Fig. 2). Interestingly, the time course of SDF‐1α induced VEGF mRNA expression was different from Egr‐1 mRNA expression. The VEGF signal intensity peaked at 4 h after stimulation and returned to control levels 8 h after cell stimulation.

Figure 1.

Figure 1

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(a) and (b): Effects of SDF‐1α (50 ng/mL) on the expression of Egr‐1 mRNA in endothelial cells. Cells were serum‐deprived for 4 h prior to stimulation with SDF‐1α for the indicated time‐intervals. Expression of Egr‐1 and GAPDH mRNA was analysed by RT‐PCR. Analysis of GAPDH mRNA expression was performed as a control for the same amount of RNA. One representative experiment out of n= 3 with similar results is shown (a). The densitometric results of these experiments are shown in parallel (b).

Figure 2.

Figure 2

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a and b: Effects of SDF‐1α (50 ng/mL) on the expression of VEGF mRNA in endothelial cells. The procedures were similar to Figure 1.

Phosphorylation of ERK1/2

After 6 h in starvation, EC were stimulated with 50 ng/mL of SDF‐1α for different time intervals. The primary antibodies used recognized p38mapk, ERK‐1 and ERK‐2 only when they were catalytically activated by phosphorylation. There was a significant base‐line phosphorylation of ERK‐1 and ERK‐2. However, stimulation with SDF‐1α, 50 ng/mL, resulted in a time‐dependent increase of ERK1/2 phosphorylation with a peak at 10 min. In contrast, a phosphorylation of p38mapk by SDF‐1α, 50 ng/mL, could not be detected (Fig. 3).

Figure 3.

Figure 3

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Effects of SDF‐1α (50 ng/mL) on the phosphorylation of ERK1/2 and MAP kinase p38 in endothelial cells. Cells were serum‐deprived for 4 h prior to stimulation with SDF‐1α for the indicated time‐intervals. Lysates were prepared and subjected to gel electrophoresis. Western blotting was performed with antibodies that just recognize phosphorylated ERK1/2 and p38. In an experiment regarding phosphorylation of p38, a positive control was added. One representative experiment out of n= 3 with similar results is shown.

Effects of ERK1/2 or p38mapk Inhibition on Egr‐1 mRNA expression

For inhibition experiments, EC were set on starvation medium for 3 h. They were then incubated with PD098059, which specifically inhibits phosphorylation of ERK1/2, or SB203580, which has been shown to inhibit phosphorylation p38mapk, for 1 h in different concentrations. This was followed by a stimulation with SDF‐1α, 50 ng/mL for 60 min for Egr‐1 mRNA expression.

As shown in Fig. 4, preincubation with PD098059 resulted in a concentration‐dependent decrease of Egr‐1 mRNA expression. In contrast, SB203580 failed to suppress Egr‐1 mRNA expression, demonstrating an ERK1/2 mediated expression of Egr‐1 by SDF‐1α in EC.

Figure 4.

Figure 4

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a and b: Effects of inhibition of ERK1/2 and MAP kinase p38 on mRNA expression of Egr‐1 1 h after stimulation with SDF‐1α (50 ng/mL). Cells were serum‐deprived for 4 h prior to stimulation and preincubated with PD098059 and SB203580 with the indicated concentrations. Expression of Egr‐1 and GAPDH mRNA, was analysed by RT‐PCR. Analysis of GAPDH mRNA expression was performed as a control for the same amount of RNA. One representative experiment out of n= 3 with similar results is shown (a). The densitometric results of these experiments are shown in parallel (b).

Determination of the DNA Synthesis in HUAEC

As demonstrated in Fig. 5, stimulation of HUAEC with 50 ng/mL VEGF caused an increase of [3H]thymidine incorporation from 51 ± 8 to 183 ± 45 cpm/µg protein. In opposite, SDF‐1α did not influence the [3H]thymidine incorporation in neither concentration, but it significantly enhanced the VEGF induced DNA synthesis, leading to a [3H]thymidine incorporation of 265 ± 55 cpm/µg protein in case of a combination of VEGF with SDF‐1α 100 ng/mL.

Figure 5.

Figure 5

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Endothelial cell proliferation in the presence of VEGF alone and with SDF‐1α in different concentrations. HUAEC were seeded in 24‐well culture plates and grown to 70% confluence. The medium was then replaced by serum‐free Medium 199. Cultures were then exposed to 50 ng/mL VEGF and/or SDF‐1α 1, 10, 50 and 100 ng/mL for 20 h before [3H]thymidine were added to the serum‐free medium. Four hours later, experiments were terminated and 100 µL of this solution were mixed with 5 mL scintillant and quantified using a liquid scintillation counter. Fifty microliters of the residual solution were prepared for the determination of protein using the Bio‐Rad protein assay. The results are shown as mean ± SD of 4 different experiments. *p < 0.05 compared with VEGF alone.

DISCUSSION

This study is the first to demonstrate a stimulation of Egr‐1 mRNA expression in human endothelial cells by SDF‐1α. The presented kinetics of induced Egr‐1 or VEGF mRNA expression are comparable with known data in human EC (Choi et al. 1994; Ko et al. 1995; Ko et al. 1999), which showed a maximal expression of Egr‐1 within 30–60 min after stimulation and of VEGF within 4 h and a return to basal levels after 2 h for Egr‐1 and 8 h for VEGF.

Furthermore, we demonstrated a time‐dependent phosphorylation of the extracellular signal‐regulated kinases (ERK)1/2. This is in accordance with previous studies that observed an ERK 1/2 phosphorylation after stimulation with SDF‐1α in haematopoietic cells (Ganju et al. 1998; Popik et al. 1998) and in EC (Molino et al. 2000). The kinetics described in these studies also show a peak expression of phosphorylated ERK1/2 at about 5–10 min after stimulation and a return to basal levels within 15–30 min. The comparatively strong signal of phosphorylated ERK1/2 we found at the base‐line was probably due to the long period EC were kept in starvation medium, which is a highly nonphysiological procedure.

In contrast to ERK1/2, we did not observe a phosphorylation of MAP kinase p38 in EC after stimulation with SDF‐1α. The data concerning a SDF‐induced phosphorylation of p38 are contradictory. While some studies described such a phosphorylation in T‐cells or neuronal cells (Kaul & Lipton 1999; Misse et al. 2001), others did not (Ganju et al. 1998; Bajetto et al. 2001).

It is widely accepted that phosphorylation of MAP kinases like ERK1/2, correlates with their activation (Seger & Krebs 1995; Bokemeyer et al. 1996). Thus, ERK1/2 should be involved in the intracellular signalling in EC after stimulation with SDF‐1α and mediate the expression of certain genes. In fact, we found a reduction of Egr‐1 mRNA expression dependent upon the concentration of the ERK1/2 inhibitor PD098059, while an inhibition of p38 MAP kinase with SB203580 had no effect on the amount of Egr‐1 mRNA expression. The dependency of Egr‐1 mRNA expression on activation of ERK1/2 was previously described in different kinds of cells like monocytes or EC (Schwachtgen et al. 1998; Guha et al. 2001; Morimoto et al. 2001), but a p38 mediated expression of Egr‐1 in Jurkat T‐cells was also reported (Rolli et al. 1999).

Since Egr‐1 and VEGF evoke a wide variety of responses, the results presented here indicate that SDF‐1α may participate in different physiological and pathophysiological processes of endothelial cells. For example, Egr‐1 mRNA expression can be enhanced by certain growth factors like bFGF or VEGF (Ko et al. 1995; Liu, Tsai & Aird 2000) and, in turn, Egr‐1 itself results in an expression of growth factors like VEGF, PDGF or transforming growth factors (Khachigian & Collins 1997; Houston et al. 2001), but also alters transcription of genes like adhesion molecules, cytokines and plasminogen activator (Silvermann & Collino 1999). Thus, Egr‐1 is considered to be involved in angiogenesis and inflammation or in the pathogenesis of vascular diseases like arteriosclerosis (McCaffrey, Fu & Du 2000). A participation in angiogenesis also seems to be true for ERK1/2 (Redlitz et al. 1999; Yu & Sato 1999).

VEGF mainly acts via its receptors KDR and Flt‐1. While KDR seems to be the receptor which is responsible for the angiogenetic potency of VEGF, Flt‐1, in addition to other functions, mediates VEGF‐induced migration in human monocytes and macrophages (Barleon et al. 1996; Hiratsuka et al. 1998; Neufeld et al. 1999; Hiratsuka et al. 2001). At that time the significance of the SDF induced expression of Egr‐1 and VEGF as well as its function in endothelial cells remains unclear, but further studies focusing on this point seem to be reasonable.

Because of the induction of VEGF mRNA expression by SDF‐1α we analysed the influence of this chemokine on proliferation of HUAEC. Using the method of [3H]thymidine incorporation as a marker of cell proliferation we found, that SDF‐1α alone did not affect the DNA synthesis rate, but it significantly enhanced the [3H]thymidine incorporation by VEGF.

SDF‐1α is known to influence the proliferation of certain cells, but while the proliferation in astrocytes is due to SDF alone (Bajetto et al. 2001), in other cells like granule precursor cells or haematopoetic progenitor cells it just, comparable to our data, enhances the proliferative effects of cytokines or Sonic hedgehog (Lataillade et al. 2000; Klein et al. 2001). As mentioned initially, the only study examining the influence of SDF‐1α on proliferation in endothelial cells by using a cell counter failed to demonstrate an effect (Mirshahi et al. 2000), but this group did not use a combination of SDF‐1α with a growth factor.

The mentioned enhancement of VEGF induced proliferation in EC by proteins with no or only less own effect was already described for, e.g. angiopoietins and insulin‐like growth factor (Huang et al. 1999; Castellon et al. 2002), for SDF‐1α this synergism has not been shown. It has to be elucidated if the proliferative potency of SDF is just caused by the induction of VEGF expression or if it is based on another pathway. However, since SDF‐1α fails to offer a proliferative effect in the absence of exogenous VEGF, the signification of the induced VEGF expression concerning the mitogenic potency of SDF‐1α seems to be rather low.

In conclusion, we were able to show, that SDF‐1α offers different effects in human endothelial cells and that it seems to be involved in certain physiological and pathophysiological conditions like, e.g. cell proliferation, thus playing a so far underestimated role in vascular biology.

ACKNOWLEDGEMENTS

We thank Professor Werner Stolpe and his staff from the Johanniter‐Krankenhaus, Bonn, Germany, for continuously providing umbilical cords. We thank Dr Artur‐Aron Weber from the Institut für Pharmakologie, Düsseldorf, for his review of the paper and helpful comments. We thank Dipl.‐Stat. C. Nicolay from the Institut für Medizinische Biometrie for her advice in statistics. This work was supported by a grant from BONFOR, Bonn, Germany (110/12).

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