Gene Transfer of NAD(P)H Oxidase Inhibitor to the Vascular Adventitia Attenuates Medial Smooth Muscle Hypertrophy
We previously showed that a systemic inhibitor of gp91phox (nox2)-based NAD(P)H oxidase abolishes angiotensin II (Ang II)–induced vascular hypertrophy. In the present study, we tested whether perivascular transfection with Ad-gp91ds-eGFP (an adenoviral bicistronic construct targeting NAD(P)H oxidase in fibroblasts) or controls Ad-CMV-eGFP and Ad-scrmb-eGFP would affect medial hypertrophy in response to Ang II. In C57BL/6J mice, we applied Ad-gp91ds-eGFP or controls to the left carotid adventitia, and 2 days later we implanted minipumps delivering vehicle or Ang II (750 μg/kg per day) for 7 days. None of the viral treatments affected Ang II–induced systolic blood pressure elevation. Immunohistochemical staining showed marker eGFP in adventitial fibroblasts and some macrophages, indicating expression of the gp91ds inhibitor. As expected, Ang II induced medial hypertrophy (medial cross-sectional area, 32.96±2.04 versus 20.57±1.00×103 μm2, Ang II versus control; P<0.001) that was significantly inhibited by Ad-gp91ds-eGFP (26.23±0.90×103 μm2; P<0.01) but not control viruses. Application of viruses alone did not change medial size under control conditions. Immunohistochemical staining and semiquantitative analysis showed a 70% increase in reactive oxygen species levels measured by the lipid peroxidation byproduct 4-hydroxynonenal (4-HNE) throughout the carotid wall in the Ang II group versus vehicle. After treatment with Ad-gp91ds-eGFP, 4-HNE generation was normalized. Thus NAD(P)H oxidases in adventitial fibroblasts and macrophages appear to modulate Ang II–induced medial hypertrophy.
Numerous reports demonstrate that hypertension leads to smooth muscle hypertrophy and increased vascular cross-sectional area, which arise from both pressure-dependent and pressure-independent mechanisms.1–5 Studies suggest that systemic or local angiotensin II (Ang II) may be an important direct mediator of vascular responsiveness and growth.5–7 In fact, early reports using cultured smooth muscle cells showed that Ang II directly induced medial smooth muscle hypertrophy via cell-surface receptors.4,8 More recently, NAD(P)H oxidase-derived reactive oxygen species (ROS) were shown to be involved in this activation,9 which in turn can stimulate a signaling cascade involving mitogen-activated protein kinases and transcription factors, leading to the growth response.10–13
We have postulated that vascular adventitial fibroblast and macrophage NAD(P)H oxidase-derived ROS partially mediate a paracrine signaling process leading to medial hypertrophy. Our laboratory and others have reported activation of adventitial fibroblast NAD(P)H oxidase by Ang II,14,15 atherosclerosis,16 and injury.17 Moreover, Ang II causes adventitial accumulation of macrophages enriched in NAD(P)H oxidase.18 A few key studies are consistent with a vascular paracrine effect on medial responsiveness initiated by ROS. These include our previous study showing that Ang II–induced impairment of endothelium-dependent relaxation of the isolated perfused mouse aorta was significantly improved by localized delivery of superoxide dismutase to the adventitia.19 Wang et al showed that Ang II stimulates NAD(P)H oxidase-derived ROS in the aortic adventitia and intima, concomitant with medial hypertrophy.20 This stimulation was significantly reduced in mice without gp91phox-containing NAD(P)H oxidase, suggesting a paracrine interaction between the media and the adjacent adventitia and intima, which contain gp91phox. Finally, studies have shown that oxidative stress causes cellular release of factors capable of activating growth-related kinases.21,22 Taken together, these findings suggest that adventitial NAD(P)H oxidase is capable of initiating a paracrine medial hypertrophic effect.
We showed that the NAD(P)H oxidase inhibitor gp91ds-tat (decoy peptide for cytosolic oxidase component p47phox) specifically blocks Ang II–induced aortic NAD(P)H oxidase activity in vitro and in vivo.23,24 Here we delivered the active gp91ds portion of the inhibitor to the adventitia by adenoviral techniques to examine whether NAD(P)H oxidase plays an important role in Ang II–induced vascular ROS production and medial hypertrophy. Delivery of gp91ds involved perivascular application of a replication-deficient adenovirus containing a fibroblast-active promoter that drives expression of the gp91ds inhibitory sequence. To increase the likelihood of localized adventitial expression, we used the platelet-derived growth factor β receptor (PDGFβrec) promoter, which is active in fibroblasts.25,26 Our data demonstrated localized adventitial expression of the oxidase inhibitor in fibroblasts and some macrophages. More importantly, we demonstrated partial reduction of Ang II–induced medial hypertrophy of the mouse carotid artery, suggesting that these cells are involved in the processes leading to medial hypertrophy.
Materials and Methods
Adenoviral Constructs: Ad-gp91ds-eGFP
We designed a bicistronic adenoviral expression vector, Ad-PDGFβrec-gp91ds-IRES-eGFP (Ad-gp91ds-eGFP), to co-express the gp91ds inhibitory sequence and enhanced green fluorescent protein (eGFP) under the control of the PDGFβrec promoter. pPDGFβrec–luciferase plasmid26 was generously provided by Dr. K. Funa (Göteborg University, Sweden). A 1468-bp portion of the pPDGFβrec–luciferase was removed by Sac1 digestion26 (PDGFβrec promoter sequence) and placed into a replication-deficient human adenovirus serotype 5 vector. This 1468-bp portion of the pPDGFβrec–luciferase vector was placed upstream from a Kozac sequence (GCC-ACC-ATG), followed by the nucleotide sequence encoding the gp91ds peptide (TGC-TCG-ACA-AGG-ATT-CGA-AGA-CAA-CTG), an internal ribosome entry site (IRES) sequence, a 719-bp sequence encoding eGFP, and a sequence for SV40 pRep8 poly A. Thus expression of eGFP would indicate expression of gp91ds.
We used 2 control viruses in these studies. The first, Ad-CMV-eGFP, expresses reporter protein eGFP under the control of the CMV promoter. In the second, Ad-CMV-scrmb-eGFP, the CMV promoter was placed upstream from a Kozac sequence (GCC-ACC-ATG), followed by the nucleotide sequence encoding the previously described scrambled gp91ds peptide sequence (TGC-CTG-AGG-ATT-ACA-AGG-CAA-TCG-AGG),23 an IRES sequence, a 719-bp sequence encoding eGFP, and a sequence for SV40 pRep8 poly A.
Animals and Viral Transfection
Twelve-week-old to 16-week-old male C57BL/6J mice (Jackson Laboratories, Bar Harbor, Me) were anesthetized with pentobarbital (50 mg/kg per day, intraperitoneally) and the neck was dissected to expose the left common carotid artery (CCA). Seven treatment groups were studied. In 3 groups (set 1), 15% pluronic gel (Poloxamer 407 NF; BASF) or gel with Ad-CMV-eGFP or Ad-CMV-scrmb-eGFP at a concentration of 3.5×108 pfu/mL was spread evenly around the outside of the left CCA using a spatula, and vehicle infusion was begun 2 days later. In 4 other groups of mice (set 2), gel alone or Ad-gp91ds-eGFP, Ad-CMV-eGFP, or Ad-CMV-scrmb-eGFP in gel was applied to the left CCA at a concentration of 3.5×108 pfu/mL, and Ang II infusion was begun 2 days later. After application of the adenovirus in gel or gel alone, the incision was closed and the animals were allowed to recover, with free access to water and chow. All protocols were approved by the Institutional Animal Care and Use Committee of Henry Ford Hospital and are consistent with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health.
Infusion of Vehicle or Ang II
Two days after viral transfection (day 0), the mice were re-anesthetized and osmotic minipumps (Alza 2ML1) were implanted subcutaneously for drug infusion. The mice in set 1 were infused with vehicle (saline with 0.01 N glacial acetic acid) at a rate of 10 μL/h for 1 week. The mice in set 2 were infused with Ang II (750 μg/kg per day; Bachem, Torrance, Calif) dissolved in vehicle.
Systolic Blood Pressure Measurement
Systolic blood pressure was measured on days −3 (basal) and 0 before pump implantation and 7 days after implantation using a standard tail cuff (IITC/Life Science Instruments).
Preparation of Tissue Samples
On day 7, mice were re-anesthetized and transcardially perfused with 10 mL phosphate-buffered saline (PBS) followed by 10 mL of 10% formaldehyde in PBS under pressure (120 mm Hg). The middle portions of left CCAs were harvested, taking care not to disrupt the adventitia. Each artery was processed, embedded in paraffin, and serially sectioned (6-μm sections).
Immunohistochemistry for Carotid Artery eGFP Expression, Macrophage Infiltration, and Lipid Peroxidation Byproduct 4-Hydroxy-2-Nonenal Generation
Monoclonal mouse anti-eGFP (Clontech) was used to detect eGFP expression; purified rat anti-mouse Mac-3 monoclonal antibody (BD Biosciences), a specific marker for mouse macrophages, was used to detect macrophage infiltration in the artery wall; monoclonal anti–4-hydroxy-2-nonenal (4-HNE) (Oxis) was used to detect lipid peroxidation as a result of ROS generation. The sections were heated at 58°C for 2 hours, deparaffinized and hydrated using standard techniques, and then boiled in 10 mmol/L citric acid buffer for 10 minutes for antigen retrieval. The sections were incubated in 0.3% hydrogen peroxide in 80% methanol for 30 minutes. For eGFP and 4-HNE, immunostaining was performed using a mouse IgG kit (Vector Laboratories). The sections were blocked with 5% normal horse serum for 30 minutes, then incubated overnight at 4°C with primary antibody (1:250 dilution for anti-eGFP and 10 μg/mL for anti–4-HNE in PBS containing 1% normal horse serum). Negative controls were incubated with the same concentration of matching IgG isotype (IgG2a for eGFP; IgG1 for 4-HNE). Sections were incubated with biotinylated secondary antibody for 30 minutes, followed by ABC reagent for 30 minutes (Vector Laboratories). The reaction was visualized using diaminobenzidine tetrahydrochloride (DAB) (Vector Laboratories). For Mac-3 immunostaining, the sections were blocked with 5% normal mouse serum for 30 minutes, then incubated overnight at 4°C with primary antibody (1:50 dilution in PBS containing 1% normal mouse serum). Negative controls were incubated with the same concentration of matching IgG isotype (purified rat IgG1). Sections were incubated with biotinylated mouse anti-rat IgG1/2a (BD Biosciences) as secondary antibody for 30 minutes, followed by ABC reagent for 30 minutes (Vector Laboratories). The reaction was visualized using DAB. Sections were counterstained with hematoxylin (Sigma) and the slides were dehydrated in alcohol, cleared in xylene, and mounted. Cell identification was performed by a staff pathologist.
Quantification of Immunostaining of 4-HNE
To assess changes in ROS levels, images of 4-HNE–stained carotid cross-sections were viewed at ×200 magnification and scored by 3 experienced observers in blind fashion. Area and intensity of staining were scored from 0 to 3 as follows: 0, no visible staining; 1, faint staining; 2, moderate staining; and 3, strong staining.
Histological Examination for Cross-Sections of Mouse Carotid Arteries and Measurement of Vascular Hypertrophy
Sections were stained with an Accustain Trichrome (Masson) kit (Sigma). Briefly, sections were deparaffinized and hydrated, then stained in preheated Bouin solution (Sigma) at 56°C for 15 minutes. The slides were cooled and washed in running water. Sections were stained in working Weigert iron hematoxylin solution for 5 minutes, rinsed in deionized water, then stained in Biebrich scarlet-acid fuchsin (Sigma) for 5 minutes and rinsed in deionized water. Slides were placed in working phosphotungstic/phosphomolybdic acid solution for 5 minutes and then in aniline blue solution (Sigma) for 5 minutes. They were washed in 1% acetic acid for 2 minutes, rinsed and dehydrated in alcohol, cleared in xylene, and mounted. Cross-sectional area and radial thickness of the media and circumference of the internal elastic lamina (CIEL) were measured using SigmaScan Pro 5.0 and Spot Version 3.4. The diameter of the lumen was calculated by dividing CIEL by π. Remodeling was determined among groups by comparing the ratio of medial thickness to lumen diameter.27
All values are expressed as mean±SEM; n indicates the number of experiments in vivo. ANOVA with repeated measures was used to analyze systolic blood pressure, thickness, 4-HNE, and body weight. ANOVA with contrast statements was used to analyze CIEL and CSA. Hochberg step-up method was used to adjust for multiple comparisons so that family-wise α levels of 0.05 were maintained.
Systolic Blood Pressure Measurements
Systolic blood pressure was significantly increased in mice receiving Ang II compared with vehicle, but there were no significant differences among Ang II, Ang II plus Ad-gp91ds-eGFP, or Ang II plus Ad-CMV-eGFP at any time point (Figure 1). Body weight did not increase during the treatment period in any of the groups, and there was no significant difference at any time point among vehicle-treated, Ang II–treated, and Ang II plus adenovirus-treated groups or within groups between days 0 and 7 (supplemental Table I, available in the online data supplement at http://circres.ahajournals.org).
Expression of gp91ds in the CCA
Application of Ad-gp91ds-eGFP to left CCAs resulted in eGFP expression in the adventitia, indicating effective delivery of gp91ds. Immunohistochemistry for eGFP in cross-sections of the left CCA 7 days after the start of Ang II infusion (9 days after transfection) showed staining in 5 of 6 arteries. In those showing eGFP expression, distribution was limited to adventitial cells. Transfected adventitial cells were identified as fibroblasts according to their fusiform morphology (Figure 2A, red arrows) and a subpopulation of macrophages (Figure 2A, blue arrows). We confirmed that the latter were macrophages using a monoclonal antibody against Mac-3 (a specific marker for mouse macrophages) (Figure 2C). Consistent with previous findings,18,28 macrophages were abundantly distributed in the adventitia of CCAs. Interestingly, however, some macrophage-like cells (Figure 2A, green arrows) were not transfected with eGFP. Thus, it appears that only a subpopulation of macrophages expressed eGFP. Importantly, we observed no eGFP expression in medial smooth muscle cells (Figure 2A).
In CCAs treated with Ad-CMV-eGFP, immunostaining was found throughout all 4 arteries. In contrast, eGFP was expressed in cells throughout the adventitia, with some staining in the media (Figure 2B), consistent with ubiquitous activation of the CMV promoter. Panel D shows a negative control from the Ad-gp91ds-eGFP group, in which control isotype IgG2a was used as the primary antibody in place of monoclonal anti-eGFP.
Ang II–Induced Vascular Hypertrophy
Trichrome staining revealed an Ang II–induced increase in medial CCA cross-sectional area compared with vehicle that was partially reduced by Ad-gp91ds-eGFP. Under basal conditions, transfection of CCAs with Ad-CMV-eGFP or Ad-gp91ds-eGFP alone did not alter medial CSA compared with control (Figure 3B and 3C versus 3A; Figure 4A). Figure 4A shows a significant increase in cumulative CSA between Ang II–treated and vehicle-treated mice. Moreover, transfection with Ad-CMV-eGFP in Ang II–treated mice did not attenuate medial CSA (Figure 3E versus 3D; Figure 4A). In contrast, transfection with Ad-gp91ds-eGFP in Ang II–treated mice partially reduced medial CSA (Figure 3F versus 3D and 3E). ANOVA showed a significant difference between Ang II plus Ad-gp91ds-eGFP versus Ang II or Ang II plus Ad-CMV-eGFP (Figure 4A). Measurements of medial thickness among these groups followed the same trend (Table). In a separate experiment, CCAs were also transfected with a recently available control virus, Ad-CMV-scrmb-eGFP, which had no effect on the Ang II–induced increase in medial CSA (Figure 4B).
The Table shows a trend toward outward remodeling of the CCA as measured by changes in the circumference of the internal elastic lamina (CIEL) in Ang II–treated groups. Normalization of the data comparing medial wall-to-lumen ratio in each group demonstrated significant outward remodeling in all of the Ang II–treated groups. However, none of the viruses had any effect on outward remodeling (Figure 5).
In Situ ROS Detection
Immunostaining showed a significant increase in the lipid peroxidation byproduct 4-HNE in all segments of the carotid wall in Ang II and Ang II plus Ad-CMV-eGFP groups compared with vehicle (Figure 6D and 6E versus 6A). After treatment with Ad-gp91ds-eGFP, 4-HNE generation was significantly decreased in all segments of the artery wall compared with Ang II and Ang II plus Ad-CMV-eGFP (Figure 6F). Applying the viruses under basal conditions did not alter 4-HNE levels compared with vehicle (Figure 6B and 6C versus 6A; Figure 7). Semiquantitative analysis of 4-HNE staining of the CCA showed an ≈80% increase in CCA 4-HNE staining in Ang II–treated versus vehicle-treated mice. Treatment with Ad-gp91ds-eGFP significantly attenuated 4-HNE to near-normal levels, whereas Ad-CMV-eGFP did not.
Our findings appear to suggest for the first time that vascular NAD(P)H oxidase is causally involved in medial hypertrophy in vivo and adventitial cells play a ROS-dependent, paracrine role in Ang II–induced medial hypertrophy of the mouse carotid artery. In the current study, we demonstrated that perivascular delivery of a novel adenoviral vector containing a PDGFβrec promoter driving expression of gp91ds, an NAD(P)H oxidase inhibitor, limited expression of the transgene to fibroblasts and some macrophages in the adventitia but not the vascular media. This treatment significantly attenuated vascular ROS detection across the vascular wall and partially attenuated Ang II–induced medial hypertrophy. A series of reports have suggested that vascular NAD(P)H oxidase activation is involved in medial smooth muscle cell hypertrophy,4,5,8 but this is the first report to our knowledge showing that vascular NAD(P)H oxidase is critically involved and plays a paracrine role in this response in vivo.
As expected, Masson trichrome staining revealed medial hypertrophy of the CCA in Ang II–treated mice. Interestingly, perivascular application of Ad-gp91ds-eGFP significantly reduced medial hypertrophy, but control virus Ad-CMV-eGFP and Ad-scrmb-eGFP did not. When the adenoviral constructs were applied under control conditions (vehicle infusion), medial size was not affected. We previously reported that NAD(P)H oxidase is involved in Ang II–induced vascular hypertrophy by demonstrating that systemic oxidase inhibition reduced medial CSA and thickness.29 Immunohistochemical data revealed that Ang II–enhanced ROS levels were significantly reduced by systemic infusion of the cell-permeant oxidase inhibitor gp91ds-tat, supporting the functional role of NAD(P)H oxidase in medial hypertrophy.29 Those data pointed to a significant role for a ubiquitous NAD(P)H oxidase in Ang II–induced vascular inflammation and hypertrophy, suggesting for the first time to our knowledge that these NAD(P)H oxidase-mediated processes may be independent of changes in blood pressure.29 However, because of the expected broad distribution of the cell-permeant peptide, it was unclear whether systemic or vascular inhibition of NAD(P)H oxidase contributed to this response. In the current study, delivery of the active portion of this inhibitor to the perivascular space caused significant inhibition of medial growth, suggesting that the vascular adventitia may play an important paracrine role in medial hypertrophy.
We wanted to determine the role of fibroblasts in this response by placing the expression of the active sequence of our gp91ds-tat inhibitor under the control of the PDGFβrec promoter. Thus we chose a promoter previously shown to be selective for proliferative cells (primarily fibroblasts).25,26 Because we deduced that fibroblasts are the most undifferentiated and proliferative cell type in the vessel wall under our experimental conditions, we postulated that Ad-gp91ds-eGFP would primarily direct expression to fibroblasts. In fact, we have shown that the promoter is active in cultured fibroblasts and not smooth muscle cells (unpublished observations, Dourron et al). Immunohistochemistry for eGFP in the current study suggested expression in fusiform cells that were identified as fibroblasts, but we also observed expression of eGFP in macrophages. The fact that only a subpopulation of macrophages was transfected is unclear but may suggest that these cells have a greater proliferative potential. Thus our initial goal of transfecting only fibroblasts was unsuccessful, and we cannot exclude a potentially important role of macrophage-derived ROS in this response. Despite this shortcoming, the current study appears to verify a paracrine role of fibroblasts and macrophages in medial hypertrophy, and it highlights the importance of the broader adventitial milieu in this response. Studies are ongoing to address the relative contribution of fibroblasts and macrophages, comparing the effect of our vector in animals lacking the ability to recruit macrophages into the vessel wall.
Previous studies suggested that Ang II exerts a direct hypertrophic effect on vascular smooth muscle.4,5 In cultured smooth muscle cell preparations, NAD(P)H oxidase-derived ROS have been implicated in the hypertrophic response,9,30 in turn activating signaling pathways and transcription factors involved in the growth response.10–13 Thus autocrine pathways are essential to smooth muscle growth in response to Ang II. However, data from this and other in vivo studies suggest a more complex scenario of paracrine ROS-mediated influences on medial responsiveness and hypertrophy. First, we and others have shown that Ang II causes a vascular inflammatory response resulting in recruitment of oxidase-enriched leukocytes into the vascular wall and primarily the adventitia, concomitant with a medial hypertrophic response.18,29 Moreover, deletion of the receptor for monocyte chemoattractant protein MCP-1 (CCR2) markedly reduces macrophage accumulation in the adventitia and attenuates the medial hypertrophic response to Ang II, suggesting that macrophages influence medial hypertrophy.18 Second, a provocative study by Wang et al20 suggested a paracrine interaction in the mouse aorta. The study showed that Ang II stimulates NAD(P)H oxidase-derived ROS in the aortic adventitia and intima, concomitant with medial hypertrophy, and that this stimulation was significantly reduced in mice without gp91phox-containing NAD(P)H oxidase, suggesting a paracrine interaction between the media (which appeared not to contain gp91phox) and the adjacent adventitia and/or intima (which did contain gp91phox).20 However, in that study, gp91phox deletion knocked out a small amount of gp91phox staining even in the media; therefore, it is possible that in the studies by Wang et al, gp91phox deletion accounted for at least part of its antihypertrophic effect via direct, autocrine medial smooth muscle oxidase inhibition. Because of this, the study, although important, did not examine the potential paracrine role for gp91phox in adventitial cells. Third, ROS in the adjacent adventitia seem to reduce medial responsiveness to endothelium-dependent agonists and participate in medial smooth muscle cell migration during injury.19,31 We do not believe that those findings or our current data in any way rule out the well-established role of smooth muscle oxidase in medial smooth muscle growth. In fact, we postulate that ROS in adjacent segments of the vessel wall play a permissive role in activation of smooth muscle oxidase, leading to medial smooth muscle hypertrophy.
Our current measurements of ROS in cross-sections of CCAs appear to concur with such an interaction. Immunohistochemical analysis of 4-HNE, a marker of ROS production,32 indicated a general increase in ROS across the vascular wall in response to Ang II, which was substantially reduced by Ad-gp91ds-eGFP. The ability of gp91ds to inhibit NAD(P)H oxidase and its specificity for this class of enzymes have been reported previously.23,24,33 Although our data provide evidence for local expression of a gp91ds inhibitor in the adventitia, perivascular application of Ad-gp91ds-eGFP caused generalized reduction of vascular ROS. These data seem to suggest cross-talk between the adventitia and other vascular sections with regard to ROS generation. Thus it is tempting to speculate on possible explanations for our findings. First, hydrogen peroxide or other peroxide derived secondarily from NAD(P)H oxidase-derived O2− would be capable of diffusing across tissue and stimulating medial cell hypertrophy.9,10 In fact, both O2− and H2O2 are capable of promoting peroxidation of cellular lipids, with potential propagation through the vessel wall.34 One further possibility is that small amounts of ROS from other adjacent cells may prime the smooth muscle oxidase and thus downstream signaling, leading to growth. Finally, it is possible that cross-talk occurs via the release of growth factors. ROS stimulate vascular smooth muscle cells to release heat shock protein 90α and cyclophilins,21 which may stimulate mitogen-activated kinases and mediate smooth muscle cell hypertrophy,35 but it is not known whether the same or similar mechanisms exist in fibroblasts or macrophages. Future studies will be necessary to address the direct or indirect paracrine effect of ROS in the vasculature in vivo.
Finally, our studies demonstrate that delivery of an NAD(P)H oxidase inhibitor to adventitial cells decreased widespread vascular ROS detection and caused a reduction in medial area. These data point to a paracrine influence of the adventitia on vascular medial growth and broader vessel wall responsiveness. Moreover, our data suggest that perivascular delivery of an NAD(P)H oxidase inhibitor can be an effective therapeutic means to attenuate the response leading to vascular medial smooth muscle hypertrophy. Studies targeting NAD(P)H oxidase in other vascular cell types will be necessary to clarify their relative contribution to this process.
This work was supported by National Institutes of Health grants HL55425 and HL28982 and by American Heart Association Grants 95011900 and 9808086W. We thank Dr Keiko Funa for generously supplying the PDGFβrec promoter.
Original received December 2, 2003; resubmission received July 20, 2004; revised resubmission received August 3, 2004; accepted August 4, 2004.
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