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* Metabolism Unit, Center for Metabolism and Endocrinology, Department of Medicine and Molecular Nutrition Unit, Center for Nutrition and Toxicology, NOVUM, Karolinska Institutet at Karolinska University Hospital in Huddinge, S-141 86 Stockholm, Sweden
Department of Pathology, Wake Forest University School of Medicine, Winston-Salem, NC 27157
Published, JLR Papers in Press, June 16, 2005. DOI 10.1194/jlr.M400450-JLR200
1 To whom correspondence should be addressed. e-mail: paolo.parini{at}cnt.ki.se
in re-
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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and HNF1ß interact with this region in the human ACAT2 gene in vitro and in vivo.
These data indicate that a) the identified HNF1 binding site serves as a positive regulator sequence, b) the binding site is functionally active both in vivo and in vitro, and c) the transcription factors HNF1 and HNF1ß, which bind to this site, play an important part in the regulation of the human ACAT2 promoter.
Abbreviations: Cdx-2, caudal-related homeodomain protein-2; C/EBP, CCAAT/enhancer binding protein; ChIP, chromatin immunoprecipitation assay; EMSA, electrophoretic mobility shift assay; HNF1, hepatic nuclear factor 1; MODY, maturity-onset diabetes of the young; TESS, transcription element search software
Supplementary key words liver • cholesterol • transcription factor • gene regulation • metabolism • acyl-coenzyme A:cholesterol acyltransferase • hepatic nuclear factor 1
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Based on this new knowledge, ACAT2 may be a viable target for the treatment and prevention of diseases associated with cholesterol accumulation (e.g., atherosclerosis). Thus, it is of great importance to characterize the mechanisms involved in ACAT2 transcriptional regulation. In the present study, we identify an important liver-specific cis-acting element in the promoter region of ACAT2 that acts as a putative binding site for the hepatic nuclear factor 1 (HNF1). This element serves as a positive regulator of gene expression and is functionally active. Finally, by chromatin immunoprecipitation assay, we show that the transcription factors HNF1 and HNF1ß bind to the identified promoter region of ACAT2 in vivo in human liver.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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expression vector was a generous gift from Prof. Pal R. Njølstad and Dr. Lise Bjørkhaug Gundersen (Haukeland University Hospital, Norway). HNF1ß expression vector was a generous gift from Prof. G. U. Ryffel at the Institute for Cell Biology (Essen, Germany). A human liver sample was obtained from a donor under a protocol approved by the ethics committee at Huddinge University Hospital Karolinska. All cell culture reagents were purchased from GIBCO (Paisley, Scotland). All chemicals used for the electrophoretic mobility shift assay (EMSA) and chromatin immunoprecipitation assay (ChIP) buffers were purchased from Sigma (St. Louis, MO). Protein G-Sepharose 4 Fast Flow was purchased from Pharmacia Biotech (Uppsala, Sweden).
Cell culture
HepG2 and HEK293 cells were grown in DMEM supplemented with 10% FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 2 mM L-glutamine. HuH7 cells were grown in DMEM supplemented with 10% FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin. Cells were maintained in 75 cm2 cell culture flasks and passaged when they reached 
90% confluence.
Construction of ACAT2 promoter constructs
The 5' untranslated genomic sequence (4 kb) of the ACAT2 gene was amplified from human genomic DNA (forward primer, 5'-CTGCCCTCAGCCTATCTTGGT-3'; reverse primer, 5'-TGCGGTCTCCAGCGGGCAG-3'). To facilitate directional cloning into pGL3 vector and to isolate the promoter region, the product was digested with KpnI and PstI, because these restriction enzymes generated an 1.4 kb fragment from –1,305 to +86. 5' nested deletions of the human ACAT2 promoter construct were obtained by PCR using the 1.4 kb construct as template. The forward primers for the four truncations were as follows: –269, 5'-GACAAGCTTCTGAGAGG-3'; –782, 5'-CTAATATATGAGCTCATCC-3'; –1,044, 5'-TTAGTCCTTCCTGTGACAGC-3'; and –1,196, 5'-GAGTGTTGGTGTTGGCTGG-3'. All constructs were sequenced to confirm homology with the human ACAT2 promoter region.
Identification of possible transcription factor binding sites and mutagenesis
To search for putative transcription factor binding sites as potential regulators in the human ACAT2 promoter region, the sequence was analyzed using the transcription element search software (TESS) database (www.cbil.upenn.edu) at the University of Pennsylvania (Philadelphia, PA). Plasmids carrying specific point mutations for these cis-elements were generated using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) according to the manufacturer's recommendations. A pair of complementary primers was designed using Stratagene's World Wide Web-based primer design software program (http://labtools.stratagene.com). Mutant constructs of the p–1044 and the full-length promoter (p–1305) constructs were generated by deletion of the six bases indicated with lowercase letters. For the CCAAT/enhancer binding protein (C/EBP), the forward sequence was 5'-CTGCCCACCAGGAgttgggAGTGGGAAGGGGA-3' and the reverse sequence was 5'-TCCCCTTCCCACTcccaacTCCTGGTGGGCAG-3'. For HNF1, the forward sequence was 5'-GGAGGGGAAGGATTAATAgttaatCCCAGCAGGAACCC-3' and the reverse sequence was 5'-GGGTTCCTGCTGGGattaacTATTAATCCTTCCCCTCC-3'. These primers were used to amplify the entire plasmid DNA by PCR using Pfu Turbo DNA polymerase (Stratagene). Plasmids were sequenced (Cybergene, Stockholm, Sweden) to confirm that correct deletions were obtained.
EMSA and supershift assay
Nuclear extracts were prepared from HuH7 cells and cultured in 75 cm2 cell culture flasks, as described by Azzout-Marniche et al. (10). The forward sequence for the HNF1 binding site was 5'-AAGGATTAATAGTTAATCCCAGCAGGAACCC-3'. The underlined bases were deleted in the forward sequence for the mutated HNF1 binding site. Unlabeled double-stranded probes were generated by mixing 1 µg of forward and 1 µg of reverse oligonucleotide, 5 µl of 1 M NaCl, and double-distilled water up to 50 µl and annealed at 95°C for 10 min. Labeled probes were generated by mixing 5 µl of unlabeled probe, 2 µl of 10x PNK buffer (Promega, Madison, WI), 1 µl of 0.1 M DTT, 1.5 µl of T4 polynucleotide kinase (Promega), 2 µl of [
-32P]ATP, and double-distilled water up to 50 µl and incubated at 37°C for 1 h, before passing through a Sephadex G-50 column (Amersham, Uppsala, Sweden). The labeled probes (40,000 cpm) were combined with 10 or 20 µg of nuclear extract along with 2 µg of poly(dI-dC) (Amersham, Piscataway, NJ), 7.5 µl of DNA binding buffer [80 mM Hepes-NaOH, pH 7.6, 0.2 M NaCl, 40 mM DTT, 20% (v/v) glycerol, 2 mM EDTA, and 1.2 mg/ml bovine serum albumin], 1 µl of 0.1 M DTT, 4 µl of buffer C [10 mM Hepes-KOH, pH 7.9, 0.4 M NaCl, 0.1 mM EDTA, 5% (v/v) glycerol, 1 mM DTT, and protease inhibitors], and double-distilled water up to 30 µl. Binding reaction mixtures were incubated for 20 min at room temperature and resolved on a nondenatured (5%, w/v) acrylamide gel in 1x Tris borate EDTA at 4°C for 3 h. For competition assays, 100-fold molar excess of unlabeled probe was added to the binding reaction mixture. For supershift assays, 2 µg of HNF1 antibody (catalog number Sc-6547X; Santa Cruz Biotechnology, Santa Cruz, CA) or 2 µg of HNF1ß antibody (catalog number Sc-22840X; Santa Cruz Biotechnology) was added to the binding reaction mixture. After electrophoresis, gels were dried and exposed overnight to X-ray film at –80°C.
ChIP
ChIP was performed as described (11), with modifications. Approximately 200 mg of human liver was diced on ice, suspended in PBS, and fixed at room temperature in 1% (v/v) formaldehyde to cross-link proteins to DNA. Reactions were stopped by the addition of glycine (0.125 M) and pelleted two times by centrifugation, first at 3,000 rpm for 10 min and then at 7,500 rpm for 5 min. The final pellet was then suspended in ChIP buffer [50 mM Tris, pH 8.0, 1 mM EDTA, 0.5 mM EGTA, pH 8.0, 1% Triton X, and 0.14 M NaCl supplemented with a protease inhibitor mixture from Roche (Mannheim, Germany)] and incubated for 10 min at 4°C. Genomic DNA was sheared by sonication on ice using a Branson sonifier 250. Supernatant, collected by centrifugation for 10 min at 13,000 rpm, was preimmunocleared by the addition of protein G-Sepharose slurry for 2 h at 4°C. After centrifugation for 2 min at 5,000 rpm, the supernatant was divided into aliquots and immunoprecipitated overnight at 4°C in a solution containing 5 µg/µl sonicated salmon sperm DNA, 10 mg/ml BSA, and the specific antibodies for HNF1 or HNF1ß or an IgG antibody (catalog number Sc-2027; Santa Cruz Biotechnology) used as a baseline control. Immunocomplexes were precipitated by the addition of protein G-Sepharose slurry and centrifugation for 2 min at 5,000 rpm. Pellets were washed twice in TSE I (1% Triton X, 2 mM EDTA, 20 mM Tris, pH 8.0, and 0.15 M NaCl), once in LiCl buffer (20 mM Tris, pH 8.0, 1 mM EDTA, 0.25 M LiCl, and 1% NP40), and twice in TE buffer (10 mM Tris, pH 8.0, and 1 mM EDTA). The precipitated immunocomplexes were dissociated by incubation in TE buffer with 1% SDS at room temperature for 30 min, and the cross-linking was reversed by incubation for 6 h at 66°C. DNA fragments were purified using Wizard SV Gel and the PCR Clean-Up System kit (Promega). For real-time PCR, the ABI PRISM 7700 sequence detector and qPCR Mastermix Plus for SYBRGreen (Med Probe, Oslo, Norway) were used. The forward sequence for the human ACAT2 promoter was 5'-TTAATCCCAGCAGGAACCCAG-3' and the reverse sequence was 5'-AGTAACAGAAGGGTATGTGCTTTGAG-3'. Primers used to standardize for DNA loading in the PCR were designed using human exon 9 of ACAT2, with the forward sequence 5'-GCTATAGCCTTGGGCCACC-3' and the reverse sequence 5'-TCACAAGAATTCGACAGCCAGAT-3'. Cycle numbers for the human ACAT2 promoter minus the respective cycle numbers for human exon 9 of ACAT2 were used to calculate the amount of specific HNF1 cis-element sequence in the immunoprecipitated samples, according to the delta-cycle threshold (delta-CT) calculation. Delta-CT values were thereafter linearized and expressed as fold induction from baseline control (IgG). After 40 cycles, PCR products were separated on a 2% agarose gel to monitor that a single amplicon of the correct correspondent size was present.
Transfection and reporter activity assays
HepG2, HuH7, and HEK293 cells were plated out on six-well tissue culture plates so that they reached 70% confluence after 24 h. Transfections of HepG2 cells were performed with 2 µg of promoter construct and 2 µg of pSV-ß-galactosidase control vector (Promega) or, in the case of cotransfection assays, 2 µg of promoter construct, 2 µg of pSV-ß-galactosidase control vector, and 0.1 µg of expression vector using Fugene-6 reagent (Roche, Indianapolis, IN) at a ratio of 6:1 (Fugene-6/DNA). Transfections of HuH7 cells were performed like those for HepG2 cells, except that Lipofectin reagent (Invitrogen, Carlsbad, CA) at a ratio of 3:1 (Lipofectin/DNA) was used. Transfections of HEK293 cells were performed with 1 µg of promoter construct and 1 µg of pSV-ß-galactosidase control vector or, in the case of cotransfection assays, 1 µg of promoter construct, 1 µg of pSV-ß-galactosidase control vector, and 0.1 µg of expression vector using Lipofectamine 2000 reagent (Invitrogen) at a ratio of 0.25:1 (Lipofectamine 2000/DNA). The pSV-ß-galactosidase control vector was used to correct for variation in transfection efficiency. Forty-eight hours after transfection, cellular lysates were prepared in reporter lysis buffer (Promega). ß-Galactosidase and luciferase activities were determined using a ß-galactosidase assay kit and a luciferase assay kit, respectively, according to the manufacturer's instructions (Promega). All transfection data are expressed as luciferase activity corrected by ß-galactosidase activity. Experiments were performed in five replicates and were repeated at least twice. Data represent means ± SEM.
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RESULTS |
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1.4 kb promoter region was used as template to create four deletion constructs, termed p–1196 (–1,196 to +86), p–1044 (–1,044 to +86), p–782 (–782 to +86), and p–269 (–269 to +86). We then transiently transfected them into HepG2, HuH7, and HEK293 cells using pGL3 basic vector as a baseline control. As shown in Fig. 1, the p–1044 construct conferred maximum luciferase activity in both HepG2 and HuH7 cells, and the activity declined appreciably when comparing the p–1044 with the p–782 construct, suggesting the presence of potential positive regulatory elements in this region. Moreover, the promoter activity increased >4-fold with the p–1044 construct compared with the full-length (1.4 kb) promoter construct (p–1305) in both HepG2 and HuH7 cells, suggesting the presence of potential repressor elements. Further deletions from nucleotide positions –782 to –269 did not have a significant effect on the ACAT2 promoter. Although the HepG2 cells displayed higher basal activity than HuH7 cells, both cell lines followed similar patterns of activity.
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We screened the sequence using TESS to search for putative transcription factor binding sites as potential positive regulators in the promoter region between –1,044 and –782 bp upstream of the ATG start codon of the human ACAT2 gene. Several cis-elements were located in this core region, two of which displayed a 100% match. The first was a putative binding site for the transcription factors HNF1 and HNF1ß, and the second was a putative binding site for the transcription factors C/EBP and C/EBPß. Therefore, we performed specific mutation analyses to examine the relative importance of these elements.
Constructs carrying these specific deletions, termed p–HNF1 mutant (deletion of the putative cis-element for HNF1 and HNF1ß) and p–C/EBP mutant (deletion of the putative cis-element for C/EBP and C/EBPß), were transiently transfected into HepG2 and HuH7 cells (Fig. 2A). For mutation analyses, we used the p–1044 construct because of its high transcription activity. As shown in Fig. 2B, C, deletion of the six bases GTTGGG, which correspond to the putative cis-element for C/EBP and C/EBPß, did not have a significant effect on the luciferase activity in either HuH7 or HepG2 cells when comparing the p–1044 with the p–C/EBP mutant construct. However, deletion of the bases GTTAAT, which correspond to the putative cis-element for HNF1 and HNF1ß, decreased the activity by 5- to 6-fold in both HuH7 and HepG2 cells. Furthermore, deletion of the putative binding site for HNF1 decreased the luciferase activity to a level similar to that observed for the p–782 construct. Transfection with a construct carrying deletions for both the putative HNF1 and C/EBP binding sites (p–HNF1 and p–C/EBP mutant constructs) had the same effect on luciferase activity as deletion of the HNF1 cis-element alone. We also generated a deletion of the putative cis-element for HNF1 in the full-length promoter construct (termed p–HNF1 mutant p–1305) and transiently transfected it into HepG2 cells. Similar to what was observed for the p–1044 construct, deletion of the putative HNF1 binding site decreased the luciferase activity almost 9-fold compared with that in the full-length promoter construct, as shown in Fig. 2D.
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and HNF1ß with the human ACAT2 promoter in vitro and in vivo antibody, shifted the specific band even farther up (lane 5). The supershift was not observed when the HNF1ß antibody was used (lane 6). Furthermore, preincubation of 10 and 20 µg nuclear extracts with labeled mutated HNF1 probe did not result in the appearance of any specific band (lanes 7 and 8). Lane 9 represents labeled HNF1 probe incubated with BSA, which served as a negative control.
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and HNF1ß to bind to the HNF1 element in the human ACAT2 promoter in vitro does not necessarily confirm that these are bound to the HNF1 element in vivo. Access to DNA binding sites may be influenced by chromatin structure, occupancy of the binding sites by other DNA binding proteins, and intranuclear availability of the transcription factors (11). To determine whether HNF1 and/or HNF1ß interact with the human ACAT2 promoter in vivo, we performed ChIP using human liver (Fig. 4). After cross-linking, DNA fragments were immunoprecipitated with specific antibodies and amplified by PCR. An IgG antibody was used as a baseline control and to compare the levels of specific DNA fragments. Before the immunoprecipitations, a small aliquot of chromatin was saved and used as an input control. Immunoprecipitation with the HNF1 antibody led to an almost 15-fold enrichment of the ACAT2 promoter, whereas immunoprecipitation with the HNF1ß antibody led to a 10-fold enrichment of the promoter. These data strongly suggest that both HNF1 and HNF1ß are associated with the human ACAT2 promoter in vivo, despite the fact that we could only show a direct binding of HNF1 to this cis-element in vitro.
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and HNF1ß overexpression on the human ACAT2 promoter and/or HNF1ß could regulate expression through the identified binding site in the human ACAT2 gene. We transiently transfected HuH7 and HepG2 cells with the p–1044 or p–HNF1 mutant construct with or without expression vectors for HNF1 and HNF1ß. In HuH7 cells, transfection with the p–1044 construct along with HNF1 caused a 3- to 4-fold induction of luciferase activity (Fig. 5A). A >2-fold induction was observed when HNF1ß expression vector was used. Furthermore, no synergistic effects were observed when both HNF1 and HNF1ß expression vectors were cotransfected in HuH7 cells. In HepG2 cells, a 33% increase in luciferase activity was observed when the p–1044 construct and the HNF1 expression vector were cotransfected (data not shown). HNF1ß overexpression failed to increase p–1044 promoter activity in HepG2 cells (data not shown). We also transfected HEK293 cells with the p–1044 construct and the HNF1 or HNF1ß expression vector. Both expression vectors caused a 3-fold induction of luciferase activity (data not shown).
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and HNF1ß could regulate the human ACAT2 promoter through another cis-element, we also transfected HuH7 and HepG2 cells with the p–HNF1 mutant construct along with the HNF1 and HNF1ß expression vectors. A complete loss of HNF1-dependent stimulation by the expression vectors was seen in HuH7 cells when the p–HNF1 mutant construct was used, as shown in Fig. 5B, and in HepG2 cells (data not shown). This indicates that deletion of this HNF1 cis-element in the 5' region of the human ACAT2 gene prevents activation by HNF1 and HNF1ß in both hepatocellular cell lines.
We also examined whether the ability of HNF1 and HNF1ß to activate the promoter pertained to the 1.4 kb ACAT2 promoter construct. Therefore, we transfected HepG2, HuH7, and HEK293 cells with the full-length ACAT2 promoter construct (p–1305) along with the HNF1 or HNF1ß expression vector. The luciferase activities increased using the HNF1 expression vector and decreased using the HNF1ß expression vector for both HepG2 and HuH7 cells, as shown in Fig. 6A, B. However, the luciferase activity increase was more pronounced in HEK293 cells using the expression vectors for HNF1 and HNF1ß, as shown in Fig. 6C.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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and HNF1ß bind in human liver.
HNF1, also called HNF1, is a dimeric protein composed of an N-terminal dimerization domain, a DNA binding domain, and a C-terminal domain that is essential for the transactivation of target promoters. HNF1 and vHNF1, also called HNF1ß, share strong homologies in both the dimerization domain and the DNA binding domain and thus are able to form heterodimers and bind to the same DNA sequences (12). HNF1 and HNF1ß are expressed in various organs, including the liver, kidney, stomach, and pancreas (13–15). The expression of HNF1 and HNF1ß overlaps; however, in adult liver, HNF1ß is weakly expressed but HNF1 is much more abundant (13–15).
The HNF1 element identified at positions –871 to –866 bp is important for the hepatic-specific expression of the human ACAT2 gene and can bind to and be regulated by the transcription factors HNF1 and HNF1ß, as we now have shown in vitro and in vivo. An interesting recent paper by Odom et al. (16) listed genes occupied by the transcription factors HNF1, HNF4, and HNF6 in human liver and in pancreatic islets. However, in the list of genes bound by HNF1 in hepatocytes, ACAT2 (or Soat2) is absent. The reason for this absence is uncertain, but it may be explained by the different technical approaches used. In their genome-scale location analysis, Odom et al. (16) targeted the region spanning 700 bp upstream and 200 bp downstream of transcription start sites of putative genes, possibly leading to failure to recognize a control of ACAT2 gene expression by HNF1. Furthermore, the fact that until recently (7) ACAT1 was thought to be the primary cholesterol-esterifying enzyme expressed in human liver also could have played a role.
We were also able to demonstrate the regulation of the ACAT2 promoter region by HNF1ß overexpression. However, we failed to show any direct binding of HNF1ß in the hepatocyte-derived HuH7 cells using EMSA. This might be attributable to the fact that EMSA experiments commonly reveal the most abundant and/or highest affinity interacting protein (17), which in this case might be HNF1. However, the binding of HNF1ß to the HNF1 cis-element in the human ACAT2 gene in liver was shown using ChIP, indicating the involvement of HNF1ß in the control of ACAT2 gene expression.
Transfection of HepG2 cells with the HNF1 or HNF1ß expression vector did not result in an induction of the activity of the p–1044 construct, as mentioned above. This may be explained by the previously reported presence of intermediate to high endogenous levels of HNF1 in these cells (18, 19). On the other hand, when transfecting HEK293 cells, which do not basally express HNF1 and HNF1ß (20) or ACAT2 (6), strong induction of the activity of the ACAT2 promoter construct was observed after overexpression of HNF1 and HNF1ß. This again suggests a very strong dependence of ACAT2 gene expression upon HNF1. Nevertheless, other hepatocyte-specific factors interacting with HNF1 must also be involved in the regulation of human ACAT2 expression, because the ACAT2 gene is not expressed in organs (i.e., the kidney) with high levels of HNF1 and HNF1ß expression (15).
The expression of ACAT2 in humans is confined to enterocytes and hepatocytes, and previous studies by Song et al. (6) showed that caudal-related homeodomain protein-2 (Cdx-2) elements in the 5' region are responsible for the enterocyte-specific expression of the human ACAT2 gene. Interestingly, one of these Cdx-2 elements is located immediately upstream to the HNF1 binding site (Fig. 7). Thus, the cell type-specific expression of the human ACAT2 gene seems to be regulated by a relatively short region of the ACAT2 promoter. In support of this concept, alignment of the untranslated 5' sequences of the human and mouse genes shows high overall homology (55%) and complete identity of the region containing the HNF1 and Cdx-2 elements (Fig. 7). In the mouse, but not in the human, promoter region of the ACAT2 gene, the untranslated 5' sequence contains another conserved putative HNF1 binding site, located closer to the transcription start site (Fig. 7). The presence of this second HNF1 cis-element may partly explain the higher expression level of ACAT2 observed in livers of mice compared with humans.
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Mutation of the TCF1 gene, which encodes HNF1, causes maturity-onset diabetes of the young (MODY)-3, a disease characterized by decreased insulin secretion in juveniles (24). Although a detailed study of lipoprotein metabolism has not been performed in this condition, in particular regarding the hepatic production of cholesteryl ester, it is of interest that VLDL secretion may also be reduced in MODY3 patients (25). In mice, deletion of Tcf1 leads to disturbed bile acid transport and to an increased synthesis of bile acids and cholesterol (26). Buoyant HDLs accumulate in plasma as a result of disturbed hepatic lipase and LCAT expression (26). Nevertheless, the impact of Tcf1 deletion on ACAT2 expression and activity has not been evaluated, despite the lower VLDL cholesterol content that seems to be present in Tcf1 knockout mice (26). Moreover, in vitro studies have identified HNF1 as an essential regulator of bile acid and plasma cholesterol metabolism (26, 27). Thus, it should be of interest to investigate whether factors that regulate the expression of HNF1 may in turn affect the expression of the human ACAT2 gene.
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ACKNOWLEDGMENTS |
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expression vector. This work was supported by the Swedish Medical Research Council (Grants 71XD-14847 and 03X-7137), the National Institutes of Health (Grants HL-49373 and HL-24736), the Swedish Medical Association, and the Swedish Heart-Lung, Åke Wiberg, Fernström, and Throne Holst Foundations. Manuscript received November 12, 2004 and in revised form March 8, 2005 and in re-revised form April 21, 2005 and in re-re-revised form June 2, 2005.
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