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Cardiovascular Research 2005 68(1):155-164; doi:10.1016/j.cardiores.2005.04.028
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Copyright © 2005, European Society of Cardiology

Reduced endothelial secretion and plasma levels of transforming growth factor-β1 in patients with hereditary hemorrhagic telangiectasia type 1

Michelle Letartea,b,*, Merry-Lynn McDonalda,c, Chenggang Lid, Kirishanthy Kathirkamathambya, Sonia Veraa, Nadia Pece-Barbaraa,e and Shant Kumard

aCancer Research Program, Hospital for Sick Children, 555 University Avenue, Toronto, Canada, M5G 1X8.
bHeart and Stroke Lewar Center of Excellence, and Department of Immunology and Medical Biophysics, University of Toronto, Toronto, Canada
cInstitute for Agricultural, Rural and Environmental Health, University of Saskatchewan, Saskatoon, Canada
dDepartment of Pathology, Medical School, The University of Manchester, Manchester M13 9PT, UK
eProgram in Molecular Biology and Cancer, Samuel Lunenfeld Research Institute, Mount Sinai Hospital, Toronto, Canada

* Corresponding author. Cancer Research Program, Hospital for Sick Children, 555 University Avenue, Toronto, Canada, M5G 1X8. Tel.: +1 416 813 6258; fax: +1 416 813 6255. Email address: mablab{at}sickkids.ca

Received 31 January 2005; revised 5 April 2005; accepted 25 April 2005


   Abstract
Top
 Abstract
1. Introduction
 2. Methods
 3. Results
 4. Discussion
 References
 
Objective: To determine if patients with hereditary hemorrhagic telangiectasia (HHT) show alterations in transforming growth factor (TGF)-β and its pathways.

Methods: Blood samples were obtained from HHT patients and controls, while endothelial cells were derived from umbilical veins of newborns (HUVEC) from HHT families. TGF-β1 in plasma, or secreted by HUVEC, and plasma endoglin levels were measured by ELISA. Cellular levels of endoglin and receptor Smad proteins were tested by metabolic labeling and immunoprecipitation, mRNA levels for endoglin and TGF-β1 by real-time PCR, and receptor Smad phosphorylation by Western blotting.

Results: TGF-β1 and endoglin plasma levels analyzed in 197 individuals showed an inverse correlation with age. Circulating levels of TGF-β1 were reduced in HHT1 patients (with Endoglin mutations) compared to control, but not in HHT2 patients (with ALK1 mutations). Endoglin levels were unchanged in plasma but decreased in activated monocytes and HUVEC with an HHT1 genotype. These HUVEC also expressed reduced levels of endoglin and TGF-β1 mRNA, secreted less TGF-β1, and showed normal receptor Smad expression and phosphorylation.

Conclusions: Decreased plasma TGF-β1 levels in HHT1 patients correlate with reduced production by endothelial cells. The lower endoglin expression in these cells may alter the regulation of TGF-β1 via Smad-independent pathways.

KEYWORDS cytokines; endothelial factors; endothelial receptors; hemostasis; signal transduction


   1. Introduction
Top
 Abstract
 1. Introduction
2. Methods
 3. Results
 4. Discussion
 References
 
Hereditary hemorrhagic telangietasia (HHT) is an autosomal dominant disorder affecting 1 in 8000 individuals worldwide. The most common signs of disease are mucocutaneous telangiectases and frequent nosebleeds [1]. Serious complications include pulmonary, cerebral and hepatic arteriovenous malformations (AVMs), which can lead to severe hemorrhage, stroke, or brain abscess [2].

Two genes are causally related to HHT: Endoglin (ENG; MIM # 131195 [OMIM] ; XM_055188) on chromosome 9q33-34, mutated in HHT type 1 (HHT1) [3] and ALK1 (activin receptor-like kinase-1, ACVRL1; MIM # 601284 [OMIM] ; Z22533) on chromosome 12q, mutated in HHT2 [4]. Mutations of all types are distributed throughout both genes and most families have a unique one. Disease severity is not correlated with a particular mutation, in agreement with a model of haploinsufficiency, where mutant proteins are poorly expressed and non-functional [5].

Both endoglin and ALK1 proteins are expressed predominantly in endothelial cells and belong to the TGF-β receptor family. Endoglin is a co-receptor, binding TGF-β1 and TGF-β3 [6], only when associated with the type II receptor (TβRII). Once bound to ligand, TβRII recruits a type I receptor, which is generally ALK5, and induces its phosphorylation [7]. This in turn leads to phosphorylation of the receptor regulated Smad2 and Smad3 [8]. Endothelial cells also express ALK1, which signals through the receptor regulated Smad1, Smad5 and Smad8 [9]. However, ALK1 requires ALK5 for activity and the ALK1 receptor complex contains both type I receptors and TβRII [10,11]. One of these phosphorylated Smads then associates with the common Smad4 for translocation to the nucleus and regulation of gene transcription.

The mechanisms underlying the pathology of HHT remain unclear. Mice heterozygous for Endoglin or ALK1 genes show clinical signs of HHT [12,13] confirming that loss of a single allele causes disease. The Endoglin or ALK1 null mice die at mid-gestation due to severe cardiovascular abnormalities [9,12,14]. Their phenotype is reminiscent of mice deficient in TGF-β1, ALK5 or TβRII, indicating that both ALK5 and ALK1 pathways are essential for vascular development.

We previously reported reduced circulating amounts of TGF-β1 in Endoglin heterozygous mice relative to control littermates [15]. Another study confirmed lower levels of plasma TGF-β1 protein and reduced TGF-β1 mRNA levels in kidneys and lungs of these mice [16].

In this study, we analyzed plasma TGF-β1 levels in patients with HHT and a control group. We ascertained whether endothelial cells (HUVEC) derived from newborns with ENG mutations (HHT1) have altered levels of TGF-β1, endoglin and receptor Smads and show impaired Smad phosphorylation. Our results indicate that in HHT1, endothelial cells produce less TGF-β1, which might account for reduced circulating TGF-β1 levels. However, these effects were not due to changes in Smad phosphorylation, suggesting that endoglin may regulate TGF-β1 production in endothelial cells via Smad-independent pathways.


   2. Methods
Top
 Abstract
 1. Introduction
 2. Methods
3. Results
 4. Discussion
 References
 
2.1 Families with HHT and their clinical and molecular evaluation
Expecting parents from families with a clinical diagnosis of HHT provided informed consent for umbilical cord and placenta samples. Informed consent was also obtained from all blood donors. All procedures were reviewed and approved by the Research Ethics Board of the Research Institute at the Hospital for Sick Children. The investigation conforms to the principles outlined in the Declaration of Helsinki. All members of families with HHT are given a number and referred to with the prefix H (for HHT).

A definite clinical diagnosis for HHT was given by physicians and required three of the following criteria: recurrent epistaxis, telangiectases, evidence of autosomal inheritance, and visceral involvement [17]. The molecular diagnosis of HHT1 or HHT2 was based on identification of ENG or ALK1 mutations. Newborns with an ENG or ALK1 mutation are considered to have HHT, despite no clinical sign. Mutation analysis was performed by QMPCR and sequencing as described [18,19].

2.2 Cell culture
Human umbilical vein endothelial cells (HUVEC) were obtained from newborns of HHT families within 48 h of birth and prepared in parallel with those from normal local deliveries so that each HHT case was compared to a control HUVEC of equivalent passage. Cells were prepared, maintained and used at sub-confluence as previously described [18,20]. Acid–citrate–dextran-treated whole venous blood samples (10–20 mL) of clinically affected HHT patients and control volunteers (family members or unrelated age-matched) were fractionated by 1 x g sedimentation, through 4.5% dextran T-500/0.9% NaCl. The upper layer containing plasma and leukocytes was removed carefully and centrifuged at 1000 x g. Plasma aliquots were frozen at –70 °C and cells were fractionated by Ficoll–Paque density gradient centrifugation [20,21]. Mononuclear cells were recovered by adherence to a 100-mm plastic tissue culture treated dish for 1 h. Lymphocytes were washed and used for DNA preparation while adherent monocytes were incubated for 16–20 h in RPMI medium plus 10% fetal bovine serum (FBS) to induce optimal expression of endoglin [24].

2.3 Determination of plasma TGF-β1 and endoglin levels by ELISA
TGF-β1 was detected following previously reported procedures [22]. Plasma samples were activated with 0.15 M sodium phosphate buffer pH 2.0 and, after 20 min at 23 °C, neutralised with an equal volume of 0.15 M sodium phosphate buffer pH 12.0. The acid-activated samples contain the mature plus small latent TGF-β1. The samples were immediately transferred in duplicate and at a final two-fold dilution to plates coated with 100 µl mouse monoclonal antibody (mAb) against TGF-β1 (Genzyme, Cambridge, MA) at 1 µg/ml in PBS and blocked with 1% BSA in PBS-Tween 20 for 2 h at 23 °C. Following 4 °C overnight incubation, samples were incubated with chicken anti-TGF-β1 antibody (R and D Systems, Minneapolis, MN; 1000-fold dilution) in PBS-Tween 20 for 3 h at 4 °C, washed and incubated with HRP-conjugated rabbit anti-chicken IgG (Jackson ImmunoResearch, West Grove, PA; 2000-fold dilution) in PBS-Tween 20 plus 1% BSA for 30 min at 23 °C. The plates were washed and incubated with 100 µl/well of Amerlite signal reagent (Amersham, Aylesbury, UK) and light emission immediately measured at 420 nm in an Amerlite plate reader. A standard curve was generated from purified recombinant human TGF-β1 (R and D Systems).

Circulating endoglin was quantified by an indirect ELISA as described [22]. Briefly, 96-well white plates (Dynatech, Chantilly, VA) were coated with mAb E9 (100 µl/well, 1/1000). Test plasma samples diluted two-fold in PBS-Tween 20, were tested in duplicate. After overnight incubation at 4 °C, 100 µl/well of biotinylated mAb E9 (1/2000) was added, followed by HRP-conjugated streptavidin (DAKO, Glostrup, Denmark; 1/2000) in PBS-Tween 20 plus 1% BSA and developed with the Amerlite signal reagent. A standard sample with a pre-determined endoglin level of 100 ng/ml was titrated to generate a standard curve in each plate.

2.4 Secretion of TGF-β1 by HUVEC
Near confluent HUVEC monolayers were washed with PBS and incubated in M199 medium plus heparin and glutamine, but without FBS or growth factor. Each HUVEC sample was analyzed in a six-well plate giving 6 replicates. Cells were maintained in this serum-free medium for 7 h. The media was harvested and TGF-β1 was activated and neutralized as described above. Samples in triplicate were quantified using the Promega ImmunoAssay System (Promega, Madison, WI). The procedures and reagents are very similar to those described above and an internal TGF-β1 standard is provided. Both mature and small latent TGF-β1 are estimated; active TGF-β1 was found to be negligible.

2.5 Real-time PCR for TGF-β1 and endoglin mRNA levels
RNA was extracted by adding TRIzol (Gibco/Invitrogen, Burlington, ON, Canada) directly onto washed HUVEC monolayers and quality and concentration were assessed using the Agilent 2200 Bioanalyzer. RNA samples were reverse transcribed using oligo-dT primers (RT-PCR kit, Invitrogen) and cDNA samples were tested by real-time PCR. The expression of TGF-β1, endoglin and the housekeeping gene β2-microglobulin was measured using SYBR green (Qiagen, Mississauga, ON, Canada) with the ABI PRISM 7900 Real Time PCR System. All standard curves were generated with five-fold serial dilution (20–0.625 ng) of RNA equivalents from a standard control HUVEC. The primers used (0.5 µM) were TGF-β1 (F: GGACACCAACTATTGCTTCAGCT, R: CCTGGACACGCAGTACAGCAAG), endoglin (F: GTCTCACTTCATGCCTCCAGCT R: ACTGCCTCAACATGGACAGCCT), and β2-microglobulin (F:AGCGTACTCCAAAGATTCAGGTT, R:TACATGTCTCGATCCCACTTAACTAT). The PCR program consisted of initial activation at 95 °C for 15 min and 50 cycling steps of denaturation for 15 s at 94 °C, annealing at 58 °C for 30 s, and extension at 72 °C for 15 s. PCR data were analyzed using SDS 2.1 software (Applied Biosystems, Foster City, CA). The threshold cycle (Ct) values for TGF-β1 and endoglin curves were normalized to the β2-microglobulin Ct values. Results are expressed as a ratio representing the relative amount of TGF-β1 and endoglin mRNA levels expressed in the samples.

2.6 Metabolic labeling and immunoprecipitation
Equivalent numbers of patient and control HUVEC at sub-confluence (80–90%), or in separate experiments, of peripheral blood activated monocytes from patient and control were metabolically labeled for the optimal time of 3.5 h with [35S]-methionine, solubilized in 1% Triton X-100 plus inhibitors and immunoprecipitated with saturating amounts of mAb P3D1 and mAb P4A4 to endoglin and with Protein G and quantified as described [18,20]. Radioactivity in the band corresponding to fully processed endoglin (E) was quantified using a Phosphorimager and Image Quant Software. The patient to control pixel value ratios of E was calculated for each immunoprecipitate and the mean ± S.D. was determined from at least 4 values.

To measure Smad levels, HUVEC were metabolically labeled, lysed as above and immunoprecipitated with a mAb to Smad2/3 (Transduction Laboratories, Mississauga, ON, Canada) or a polyclonal antibody (pAb) to Smad1/5 (Upstate Biotechnology, Lake Placid, NY) and the respective bands quantified. To assess the effect of TGF-β1 on Smad2 phosphorylation, HUVEC from HHT1 and control groups were grown to sub-confluence, starved for 2 h and treated for 45 min with and without 300 pM TGF-β1 (unless otherwise indicated). Mink lung epithelial cells transfected with Smad2 served as a positive control. Cells were lysed and immunoprecipitated with the mAb to Smad2/3 and/or immunoblotted with rabbit anti-phospho-Smad2 (Upstate Biotechnology). To assess Smad1/5/8 phosphorylation, cells were treated for 15 min with 50 pM TGF-β1 or 50 pM BMP-6, conditions optimal for these Smad in murine embryonic endothelial cells. Lysates were probed with anti-phospho Smad1/5/8 (Cell Signaling Technology Inc., Beverley, MA) and with a pAb to Smad1/5. Anti-β actin (Sigma-Aldrich Canada, Oakville, ON) was used to re-probe the blots to determine equal loading.

2.7 Statistical analysis
Results of TGF-β and endoglin levels were analyzed using the statistical software package SPSS V.5. Initial bivariate analysis was performed to test for effect modification, irrespective of phenotype. As the data did not show a normal distribution, a nonparametric analysis was performed and data are reported as median plus the 25th and 75th percentile values. Wilcoxon rank statistics compared the protein levels between patients and controls. The Student t-test was employed to test the null hypothesis of no difference between patient and control real time PCR data.


   3. Results
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
4. Discussion
 References
 
3.1 Circulating levels of TGF-β1 are reduced in patients with HHT1
Levels of TGF-β1 and endoglin were measured in plasma samples collected from 197 individuals including 96 patients with a clinical diagnosis of HHT. From the bivariate analysis, it appears that circulating levels of TGF-β1 (Fig. 1A) and endoglin (Fig. 1B) were inversely correlated with age, being highest in children and decreasing progressively with age. There was no correlation with sex.


Figure 1
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  		Fig. 1 Circulating levels of TGF-β1 and endoglin are inversely correlated with age. Plasma levels of TGF-β1 (latent plus mature) were measured by ELISA in 197 individuals, with and without HHT, and plotted versus age at time of sample collection. A Spearman negative correlation was observed (r = –0.18, p<0.01) (Panel A). Levels of plasma endoglin were measured by ELISA in the same individuals and showed an inverse correlation with age. Spearman negative correlation (r = –0.22, p<0.002) (Panel B).

 
The groups were divided into patients with a defined ENG or ALK1 mutation corresponding to a molecular diagnosis of HHT1 and HHT2, respectively. Patients with a clinical diagnosis of HHT, irrespective of mutation status, were also analyzed. The HHT1 group expressed significantly lower levels of circulating TGF-β1 than the control group (25.5 versus 44.0 ng/mL) (Table 1) while the HHT2 group was unchanged. The clinical HHT group showed a trend towards reduced TGF-β1 levels, likely because of a predominance of HHT1 patients within the group (Table 1).


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  		Table 1 Summary of endoglin and latent TGF-β1 levels in HHT patients

 
The plasma endoglin levels were also measured. The HHT1 group did not show any changes in circulating levels of plasma endoglin nor did any other groups, suggesting that this parameter is not related to the HHT status, even in patients with ENG mutations (Table 1).

Endoglin levels were reduced in peripheral blood activated monocytes from the HHT1 group with a median of 53% relative to control monocytes included in each metabolic labeling experiment (Table 1). The HHT2 group had normal levels of endoglin with a median of 95.5%. The clinical HHT group showed significantly reduced levels of endoglin (62.5%) compared to control, supporting the notion of a high percentage of HHT1 patients within the clinically diagnosed group (Table 1).

3.2 HUVEC from newborns with an ENG mutation secrete reduced levels of TGF-β1 and express less TGF-β1 mRNA than control
To ascertain if the vascular endothelium contributes to plasma TGF-β1 production, levels of TGF-β1 were measured into HUVEC culture medium. The rate of TGF-β1 secretion by HUVEC from newborn H628, carrying a mutation in ENG exon 10 (see Table 2), was approximately half of the control HUVEC (Fig. 2A). The mean value for TGF-β1 secretion by 15 HUVEC with ENG mutations was 66 ± 12% of the control HUVEC analyzed in the same experiment (p<0.01) (Fig. 2B).


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  		Table 2 Characteristics of HUVEC samples with ENG mutations used in the current study

 

Figure 2
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  		Fig. 2 HUVEC with ENG mutations secrete less TGF-β1 than control cells. Sub-confluent monolayers of HHT1 and control samples, at equivalent densities and passages, were washed with PBS, incubated in medium without supplements, the culture medium harvested at different time points and TGF-β1 measured by ELISA. The kinetics of secretion of TGF-β1 was less in HUVEC from sample H628 than in a control HUVEC (Panel A). The level of TGF-β1 secreted (over 7 h) by 15 different HUVEC with HHT1 were significantly different (p<0.05) from the control HUVEC tested in the same experiment (Panel B).

 
The levels of TGF-β1 mRNA in HUVEC with HHT1 were compared to control by real-time PCR. Values were normalized with respect to β2-microglobulin mRNA levels. The relative TGF-β1 mRNA levels were significantly reduced (p<0.03) to 0.68 ± 0.08 in the HHT1 group (n = 11) compared to 0.98 ± 0.11 in the control group (n = 12) (Fig. 3).


Figure 3
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  		Fig. 3 Reduced levels of TGF-β1 and endoglin mRNA in HUVEC from newborns with ENG mutations. The levels of endoglin and TGF-β1 mRNA in HHT1 (n = 11) and control HUVEC (n = 12) were measured by Real Time PCR. The relative amounts of endoglin and TGF-β1 mRNA were obtained by normalization to β2-microglobulin levels. Both endoglin and TGF-β1 relative amounts were reduced in the HHT1 group compared to the control group (p<0.03).

 
3.3 Endoglin mRNA levels are reduced in HUVEC with ENG mutations
Mean endoglin mRNA levels, estimated by real-time PCR, were significantly reduced (p<0.03) in HUVEC with ENG mutations (0.24 ± 0.03) compared to control (0.56 ± 0.1) (Fig. 3). Table 2 lists the mutations present in the samples tested and the levels of endoglin mRNA estimated in these HUVEC. Most mutations analyzed were associated with reduced endoglin mRNA levels, further supporting previous data on reduced levels of endoglin in these samples [18,19,21,23–25].

3.4 Levels of Smad2 and Smad3 are normal in HUVEC with HHT1
We previously reported that binding of TGF-β1 was unchanged in HUVEC with ENG mutations [20], suggesting that downstream signaling events might not be altered. We therefore measured the levels of Smad2 and Smad3, known to mediate TGF-β signals via the ALK5 pathway. Fig. 4 illustrates that Smad2/3 levels are normal in HUVEC from the HHT1 group. Reduced levels of endoglin are shown for H739 HUVEC with an ENG exon 12 mutation (Table 2) while normal levels are seen in H767 HUVEC, without the familial ENG exon 10 indel mutation. Levels of Smad2 and Smad3 were unchanged in these samples, relative to the normal HUVEC ran in each experiment (Fig. 4A). These measurements were performed in 6 HUVEC with different ENG mutations (Fig. 4B). The endoglin levels were significantly different from control with an average of 39.9 ± 5.3% (mean ± S.D.), while the Smad2 (109.8 ± 23.7%) and Smad3 (118.3 ± 31.6%) levels were not. Fig. 4B shows that HUVEC from 3 newborns without the familial germline ENG mutation have normal levels of endoglin (94.3 ± 9.8%), Smad2 (105.0 ± 33.0%) and Smad3 (116.4 ± 42.5%). Thus levels of Smad2 and Smad3 are unaffected in HHT1 endothelial cells.


Figure 4
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  		Fig. 4 Levels of Smad2 and Smad3 are unchanged in HUVEC with ENG mutations. HUVEC at equivalent densities and passages and at sub-confluence were metabolically labeled with [35S]-methionine and extracts were immunoprecipitated with mAb P3D1 to endoglin or with a mAb to Smad2/3. Representative gels are shown for samples H739 with an ENG mutation and H767, without the familial ENG mutation (Panel A). E represents the mature form of endoglin while P represents the intracellular precursor. The intensity of band E in the patient HUVEC is expressed as percentage of control value and represents the mean of at least 4 determinations per sample (38 ± 2% for H739 and 83 ± 3% for H767). Levels of Smad2 and Smad3 were also quantified similarly and shown to be unchanged in H739 and H767 samples (Panel A). The mean ± S.D. obtained for endoglin (n = 6), Smad 2 (n = 6) and Smad3 (n = 5) levels in newborns with different ENG mutations are shown in black while those without the familial mutation are shown in gray (Panel B).

 
3.5 Phosphorylation of Smad2 is normal in HUVEC with ENG mutations
The induction by TGF-β1 of Smad2 and Smad3 phosphorylation in HUVEC from the HHT1 and control groups was measured. Smad2 phosphorylation was detected in four cases of HHT1 at levels similar to those observed in the control samples (Fig. 5, Panels A and B). No phospho-Smad3 was detected, likely due to the low cross-reactivity of the anti-phospho-Smad2 mAb. Levels of Smad2 phosphorylation were quantified and Fig. 5C and D show no difference between control and HHT1 samples.


Figure 5
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  		Fig. 5 TGF-β1 induced Smad2 phosphorylation is normal in HUVEC with ENG mutations. Mink lung epithelial cells (Smad2 control; S2C), control HUVEC (C) and HUVEC from the HHT1 group were grown to confluence, starved for 2 h and treated for 45 min with and without 300 pM TGF-β1 (unless otherwise indicated). Extracts were subjected to immunoprecipitation with mAb to Smad2/3, and immunoblotted with rabbit anti-phospho-Smad2 in H729 and H439 samples (Panel A). Cell extracts were analyzed directly by Western blot with rabbit anti-phospho-Smad2 for samples H713 and H455 (Panel B). Gels were scanned by densitometry and levels of phospho-Smad2 quantified for gel A (Panel C) and gel B (Panel D).

 
3.6 Phosphorylation of Smad1/5/8 is not induced by TGF-β1 in either normal or HHT1 HUVEC
Endoglin has been reported to promote the ALK1 signaling pathway [26,27]. Therefore we measured TGF-β1 effects on phosphorylation of the ALK1 regulated Smad1/5/8 in control and HHT1 groups. Preliminary experiments revealed that Smad1 and Smad5, measured by metabolic labeling and immunoprecipitation with a pAb recognizing both Smads, were expressed at similar levels in control and HHT1 HUVEC (data not shown). After treatment with TGF-β1, no significant induction of Smad1/5/8 phosphorylation was detected in HUVEC, whether they were normal or HHT1. Fig. 6 shows a representative experiment where sample H778, with a duplication of exons 2 to 4, is compared to control. Although the baseline level of Smad1/5/8 phosphorylation may be high in the HUVEC, BMP, a known inducer of this signaling pathway, gave some induction. Mouse embryonic endothelial cells stimulated in a similar manner were responsive to both TGF-β1 and BMP (data not shown). Therefore the ALK1 pathway is not inducible by TGF-β1 in HUVEC from either normal or HHT1 groups.


Figure 6
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  		Fig. 6 Smad1/5/8 phosphorylation is not induced by TGF-β1 in normal or ENG mutation bearing HUVEC. HUVEC from control (C) or from a newborn with an ENG mutation (H778) were grown to confluence, starved for 2 h and treated for 15 min with 50 pM TGF-β1or BMP-6. Extracts were fractionated by SDS-PAGE and immunoblotted with anti-phospho Smad1/5/8 or with anti-Smad1/5 to determine total Smad1/5 levels. The same gels were re-probed with anti-β actin for gel loading controls and showed no significant difference in levels of protein per lane.

 

   4. Discussion
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
References
 
Our studies demonstrate that patients with HHT1 have reduced levels of circulating TGF-β1 while HHT2 patients have normal levels. Endothelial cells from newborns with ENG mutations and therefore with a molecular diagnosis of HHT1 expressed lower levels of TGF-β1 mRNA and secreted less TGF-β1. We suggest that endothelial cells contribute significantly to the circulating pool of TGF-β1 and that endoglin level in these cells regulates the process. A significant reduction in endoglin mRNA and protein in HHT1 endothelial cells might therefore lead to reduced TGF-β1 secretion. However, the level of circulating endoglin was unchanged in HHT patients, suggesting that this parameter is not correlated with disease status.

In the overall population analyzed, an inverse correlation between age and levels of either plasma TGF-β1 or endoglin was observed, indicating that age is an effect modifier. However, our sample size was relatively small and further substantiation of this finding will require a larger population. Studies on the genetic epidemiology of HHT should consider age as a potential effect modifier and stratify patients according to age categories. HHT is a complex disease influenced by modifier genes and environmental factors acting on a vascular endothelium predisposed to lesions due to mutations in either ENG or ALK1 genes [2]. It is well documented that HHT gets increasingly worse with advancing age; for example, gastrointestinal bleeds and associated anemia are a problem of elderly patients. To what extent the lowering of TGF-β1 and endoglin plasma levels contributes to age-related disease progression remains to be evaluated. TGF-β is a key player in several complex diseases other than HHT, such as cancer, atherosclerosis and immune disorders.

TGF-β1 has multiple effects on most cell types and is generally considered a potent negative growth regulator. In endothelial cells, TGF-β1 stimulates extracellular matrix synthesis and deposition in the vessel wall and suppresses inflammatory processes. It also inhibits proliferation and migration of endothelial and smooth muscle cells and may play an important role in repairing vascular damage by reducing superoxide anions [28]. Therefore, lower TGF-β1 production by endothelial cells may predispose vessels of HHT patients to local inflammatory responses and focal damage.

In addition to paracrine effects such as between endothelial and smooth muscle cells in the vessel wall, TGF-β1 released in circulation has endocrine effects. Higher levels of plasma TGF-β1 contribute to the pathogenesis of chronic fibrosis and autoimmune diseases and may serve as a prognostic marker for carcinogenesis and atherosclerosis [29]. Current data indicate that lower levels of plasma TGF-β1 might be characteristic of HHT1, but not HHT2, a very related disorder. HHT1 often has an earlier age of onset, is associated with a higher incidence of pulmonary AVMs and is considered more severe than HHT2. The regulation of TGF-β1 observed in HHT1 and not HHT2, may be controlled by an endoglin-dependent but ALK1 independent mechanism.

TGF-β is secreted as a latent complex composed of the latency-associated peptide (LAP) non-covalently bound to the mature TGF-β and is found primarily in this latent form in vivo [30]. TGF-β can therefore circulate in this inactive form and reach its target cells prior to activation. The assays used in this study detect TGF-β1 released from the small latent complex by acid treatment, as well as any mature TGF-β1 present in the sample. A number of factors may influence plasma TGF-β1 levels, such as reduced secretion, sequestration by complex formation with other proteins including latent TGF-β1 binding proteins (LTBP) [30], which are undetectable by the assays employed in this study.

Reduced TGF-β1 synthesis and secretion by endothelial cells and decreased levels of plasma TGF-β1 are both associated with HHT1, suggesting that endoglin may affect TGF-β1 transcription in endothelial cells, which in turn would release less into circulation. The promoter of TGF-β1 is quite distinct from that of the -β2 and -β3 isoforms, as it does not contain a classic TATAA box but multiple regulatory sites that can be activated by immediate early genes (such as JNK) and various oncogenes and viral transactivating proteins. These unique features of the TGF-β1 promoter allow for the selective overexpression of this isoform in repair of injury, response to stress, viral-mediated diseases and carcinogenesis [31]. TGF-β1 can also regulate expression of its own mRNA in an actinomycin D sensitive way [32]. Two molecules have been implicated in the autoregulation of TGF-β1: Smad3 and the c-jun kinase (JNK), which acts at least in part by regulating AP-1 activity. The auto-induction of TGF-β1 was reduced in Smad3 null mice [31] while constitutive expression was seen in JNK-deficient cells [33]. In our study, Smad 3 levels were normal but TGF-β1-induced phosphorylation of Smad3 could not be detected with the available antibodies. However, both Smad2 and Smad3 translocation to the nucleus were normal (data not shown) and Smad2 phosphorylation was unchanged. We can therefore argue that the ALK5 pathway is functional and normal in HHT1 endothelial cells. JNK activity in HUVEC could not be stimulated by TGF-β1 (data not shown). Although there is cross-talk between Smad3 and JNK pathways in mediating TGF-β1 effects, it is unlikely that the reduced expression of TGF-β1 observed in endothelial cells of the HHT1 group is due to autocrine regulation via these pathways.

Two recent studies demonstrated that endoglin interacts with ALK1 and that its knockdown with siRNA resulted in loss of TGF-β-dependent Smad1/5/8 activation [26,27]. We found that HUVEC did not phosphorylate Smad1/5/8 when treated with TGF-β1 but only with BMP-6. HUVEC from the HHT1 group were no different from the control group. This lack of an ALK1 reactive pathway is likely due to the endothelial cell type since mouse embryonic endothelial cells tested under the same conditions did phosphorylate Smad1/5/8 in response to either TGF-β1 or BMP-6. Although we cannot ascertain the role of endoglin in the ALK1 pathway in our study, we can conclude that decreased TGF-β1 production in HHT1 HUVEC is not dependent on the ALK1 pathway. Therefore if endoglin affects TGF-β1 transcription, it is not via receptor-regulated Smads.

Our findings support a recent study of Eng null mice that proposed reduced availability of TGF-β1 at the endothelial–mesothelial interface in the yolk sac based on decreased immunostaining for phospho-Smad2 and reversal by addition of exogenous TGF-β1 [34]. We suggest that the reduced synthesis and secretion of TGF-β1 by endothelial cells can account for the decreased availability of TGF-β1 and the subsequent impairment in smooth muscle cell differentiation. We saw no intrinsic defect in Smad2 signaling in HHT1 endothelial cells and argue that Smad2 signaling mechanism is likely normal in smooth muscle cells.

In the current study, endoglin plasma levels were decreased with age but had no correlation with HHT. Previous studies have demonstrated that endoglin plasma levels are correlated with tumor burden and a heightened state of angiogenesis in tumors [22,35]. Previously we demonstrated that levels of endoglin were reduced to 50% in endothelial cells and activated peripheral blood monocytes of individuals with HHT1 but were normal in those with HHT2 [18–21,23,36–38]. We now show that levels of endoglin are reduced at the mRNA levels, indicating the instability of mutant RNAs and further supporting the haploinsufficiency model. It has been demonstrated that most mutations leading to frameshift and truncation result in non-sense mediated decay [39]. However, the missense mutations reported here were also associated with low mRNA transcripts and only precursor forms of endoglin protein [23], suggesting that such mutations also affect mRNA and protein stability. A recent paper has proposed that both HHT1 and HHT2 patients have reduced levels of endoglin on activated peripheral blood monocytes and that disease severity was correlated with a lowering of surface endoglin. The number of patients analyzed in that study was too small to permit such conclusions, which do not support our data with much larger number of patients [40].

Our studies reveal a selective decrease in plasma TGF-β1 level in HHT1 patients, correlating with reduced expression at mRNA and protein levels in endothelial cells. Thus reduced endoglin expression associated with HHT1 may affect the regulation of TGF-β1 in endothelial cells, but via Smad-independent pathways.


   Acknowledgements
 
This research was supported by grants from the Canadian Institute of Health Research, the Heart and Stroke Foundation of Canada and the March of Dimes to ML. We thank Dr Andrew D. Paterson and Ms Christina Goia for their invaluable assistance with statistical analysis and HHT Solutions for providing help with mutation analysis. CL is a Wellcome Trust fellow.


   Notes
 
Time for primary review 21 days


   References
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 References
 

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