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1Division of Pulmonary Sciences and Critical Care Medicine, University of Colorado Health Sciences Center, Denver, Colorado; and 2Department of Respirology, Graduate School of Medicine, Chiba University, Chiba, Japan
Submitted 9 August 2006 ; accepted in final form 12 March 2007
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ABSTRACT |
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pulmonary circulation and disease; gene expression; genetically altered mice; smooth muscle proliferation and differentiation
Linking loss of BMPR2 to downstream molecular and hemodynamic events, however, is inherently difficult; BMPR2 may signal through several different pathways. In addition to canonical SMAD signaling (10), BMPR2 may signal through p38 and p42/44 MAPK (13, 20), LIMK (3), and Tctex (9). Furthermore, in cell culture, loss of BMPR2 results in increased signaling for its ligands through alternate pathways (22). Determining the meaningful molecular effects of loss of BMPR2 signaling in the absence of in vivo context is thus problematic at best.
The goal of this study was therefore to determine the broad molecular consequences of in vivo inhibition of BMPR2 signaling in smooth muscle in our SM22-rtTA x TetO7-BMPR2delx4+ mice. In this study we performed gene array analysis of whole lung from pooled RNA samples from sets of adult mice with transgene activated for either 1 or 8 wk compared with SM22-rtTA-only controls, also fed doxycycline. To examine sex differences, both male and female pools were used. To verify results, some follow-up experiments with Western blot and quantitative RT-PCR were performed on either sample tissue or cell culture with small interfering RNA (siRNA) to BMPR2.
The most striking finding resulting from this study was that loss of signaling through BMPR2, either from the dominant negative in vivo or through siRNA in vitro, resulted in a loss of markers of smooth muscle differentiation. In addition, we found sex-specific differences in response to loss of BMPR2 in inflammatory gene expression.
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METHODS |
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20% lower pressure than sea level). The final age of all mice was 12 wk. Twelve-week-old SM22-rtTA or SM22-rtTA x TetO7-BMPR2delx4+ mice (18) weighing 20–25 g were anesthetized with intraperitoneal injections of 200 mg/kg ketamine and 10 mg/kg xylazine. If further anesthesia was necessary, repeat doses of ketamine (100 mg/kg) and xylazine (5 mg/kg) were administered. Studies were conducted with mice positioned supine on a heated operating table while spontaneously breathing room air. RV pressure was directly measured with a 1.4 French Pressure Volume Conductance System SPR-839 (Millar Instruments, Houston, TX) inserted into the RV via the surgically exposed right jugular vein. Hemodynamics were continuously recorded with a Millar MPVS-300 unit coupled to a Powerlab 8-SP A/D converter, acquired at 1,000 Hz, and captured to a Macintosh G4 computer utilizing Chart 5.3 software. After lethal injection of pentobarbital, the heart and lungs were removed. Lungs were divided and processed for immunohistochemistry or molecular studies. All animal studies were preapproved by the University of Colorado Health Sciences Center Institutional Animal Care and Use Committee. Affymetrix arrays. All RNA for arrays consisted of pooled samples from three different animals. Samples were prepared for Affymetrix arrays using 2.5 µg of total RNA. First- and second-strand complementary DNA was synthesized using standard techniques. Biotin-labeled antisense complementary RNA was produced by an in vitro transcription reaction. Mouse Genome 430 2.0 microarrays (Affymetrix, Foster City, CA) were hybridized with 20 µg of cRNA. Target hybridization, washing, staining, and scanning probe arrays were done following an Affymetrix GeneChip Expression Analysis Manual. All array results have been submitted to the NCBI gene expression and hybridization array data repository (GEO, http://www.ncbi.nlm.nih.gov/geo/) as series GSE5255 [NCBI GEO] .
Western blot. Tissues were homogenized in 500 µl of RIPA buffer with 1% (vol/vol) protease and phosphatase inhibitor cocktail. Tissue and cell lysates were centrifuged at 4°C (15 min, 10,000 g), and protein concentration was determined using a Bradford microassay (Bio-Rad, Hercules, CA) on the supernatant. Equal amounts of protein extracts were denatured at 95°C in a denaturing sample buffer. Protein from each sample (20–50 µg) was separated by electrophoresis in an 8–16% Tris-glycine gel and then transferred to a PVDF membrane in 20% MeOH with 100 mM glycine and 10 mM Tris transfer buffer. The membrane was blocked for 1 h at room temperature with PBS containing 5% nonfat dry milk and 0.05% (vol/vol) Tween 20 and probed overnight at 4°C with mouse monoclonal Cnn1 primary antibody (1:50 dilution; Sigma, St. Louis, MO). The membrane was then incubated at 37°C for 45 min with horseradish peroxidase-labeled donkey anti-mouse immunoglobulin secondary antibody (1:1,000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA). Horseradish peroxidase was detected using the ECL+ Western blotting detection system (Amersham Biosciences, Piscataway, NJ). The amount of Cnn1 protein was quantified by densitometry.
Immunohistochemistry. Tissue blocks were made from the left lungs of BMPR2 or control mice. The embedded tissue was cut at 4 µm on an AO 820 microtome (American Optical, Southbridge, MA) and placed onto Superfrost Plus slides (Fisher Scientific, Pittsburgh, PA). Slides were baked for 1 h at 60°C and then deparaffinized through Citrisolv (Fisher Scientific) and a graded alcohol series. Antigen retrieval was then performed by heating the slides 4x for 3 min at 30% power in a 1,650-W microwave in a solution containing 16 mM sodium citrate-trisodium and 4 mM citric acid, pH 5.6. Slides were then rehydrated in PBS and incubated in a blocking solution of PBS/4% serum/0.1% Triton X-100 for 30 min and then in primary antibody overnight at 4°C in PBS/4% serum/0.1% Triton X-100. Polyclonal anti-smooth muscle actin was purchased from Abcam (Cambridge, MA) and used at a dilution of 1:500. Monoclonal mouse anti-titin was purchased from Chemicon International (Billerica, MA) and used at 1:200. Tissues processed for immunofluorescence were rinsed and placed in either Alexa Fluor 488 or 594 (Molecular Probes, Eugene, OR) secondary antibody at a 1:1,000 dilution for 1 h at room temperature in the dark. Slides were then rinsed with PBS and coverslipped with Vectashield with 4,6-diamidino-2-phenylindole (Vector Laboratories, Burlingame, CA). All staining was evaluated, and digital images were acquired using a Zeiss Axioskop 2 equipped with an AxioCam (Carl Zeiss Microscopy, Jena, Germany).
Quantitative RT-PCR. Primers were designed using PrimerExpress (Applied Biosystems, Foster City, CA) from sequences downloaded from GenBank, with primers tested for specificity by BLAST. Primers were tested for creation of a single, clean band on arrival. Primer sequences are listed in Table 1.
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Cell culture and siRNA.
Human pulmonary artery smooth muscle cells (PASMC; Clonetics, San Diego, CA) were grown in SmBM media supplemented with SmGM-2-SingleQuot (Clonetics) without antibiotics in a humidified incubator at 37°C with 5% CO2. Cells were plated on 10-cm2 plates and grown to 50% confluence at passage 4. Cells were mock transfected, transfected with BMPR2 siRNA (Ambion, Austin, TX) or negative control siRNA (Qiagen) with PLUS Reagent and Lipofectamine (Invitrogen, Carlsbad, CA) in Opti-MEM (GIBCO, Carlsbad, CA) for 48 h. The cells were harvested at the same time for mRNA and protein using a PARIS kit (Ambion).
Array analysis. Affymetrix Cel files were loaded into dChip 2005 array analysis software (7). The dChip algorithm is capable of detecting significant differences at signal strengths lower than those usable in Microarray Suite (7) (Affymetrix, Santa Clara, CA). Overall signal strength from arrays was normalized to the median array, and expression levels were determined using the perfect match/mismatch algorithm. Gene ontology was determined using the Classify Genes tool within dChip (23), with gene ontology files downloaded from the Gene Ontology Consortium (www.geneontology.org) (1). To avoid problems with either false negatives, or with determining an arbitrary fold-change cutoff, we set very loose definitions for changed genes (1.4x, with a minimum change of 200) and then determined statistically overrepresented gene ontology groups at a high stringency (P < 0.001) within the genes called as differentially regulated. By this method, the number of gene ontology groups produced by chance should be close to 0. Other specific details of analytic methods are included in RESULTS.
Statistics. Statistics for array analysis were handled by algorithms internal to dChip. For quantitative RT-PCR or for densitometry on Western blots, statistics were performed in StatView (SAS, Cary, NC), with ANOVA with post hoc tests or unpaired t-test as appropriate, and significance at P < 0.05.
Statistical significance for overrepresented gene ontology groups was determined by the one-sample z-test, incorporated into the dChip software. Whereas changes in individual genes cannot be statistically significant given single samples, changes in groups of genes can be statistically tested by this method (2, 15).
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RESULTS |
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300 beats/min. There were no significant differences in hemodynamic variables between male and female mice and no change in systemic pressures by tail cuff (not shown).
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A key problem in interpretation of gene array data is the presence of noise, or spurious results. The traditional strategy to overcome this is to establish an arbitrary fold-change cutoff, since there are fewer spurious results at higher fold changes. Our strategy to overcome this was to determine significance by looking for overrepresented gene ontology groups. That is, if certain gene ontology groups are significantly overrepresented in changed genes, then they are likely to represent meaningful changes. Gene ontology groups overrepresented at a statistically significant level (P < 0.001) were determined using the dChip gene classification tool. At this level, we would expect to see no gene ontology groups by chance. This strategy has the advantage of allowing the inclusion of differentially regulated genes at lower fold-changes as well as automatically sorting genes into functional groups.
Using this strategy, we found changes in expression of groups of genes at 1 and 8 wk. Both time points had large numbers of immune-related, developmental, matrix-related, cell cycle, signal transduction, and muscle development or contraction-related genes. At 8 wk, there were also cell motility, angiogenesis, and endocytosis-related groups. Examples selected from these groups are listed in Table 2.
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, 1q
, complement component 6 (C6), and the hemolytic complement (HC) all had increased expression in female mice by 1 wk in whole lung, whereas male mice were essentially unchanged (Fig. 2B). To confirm these findings, quantitative RT-PCR was performed for C6 and HC on RNA from each individual mouse included in the pools for the control and 1-wk groups. For both genes, every female had higher expression normalized to the average of all controls than every male at 1 wk, confirming the array results (Fig. 2C). Decreased smooth muscle differentiation genes in SM22-rtTA x TetO7-BMPR2delx4+ mice at 8 wk of activation. To more closely examine the nature of the changes in muscle structure and contractile genes, we compared expression levels as measured by the arrays to a set of canonical markers of smooth muscle differentiation (11). We found that every smooth muscle marker showed decreased expression at the 8-wk time point (Fig. 3A). Note that there is a feedback loop in these mice preventing excessive loss of smooth muscle markers; the promoter driving the transactivator responsible for our transgene expression is SM22, also known as transgelin, itself a smooth muscle marker.
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3-fold) at 8 wk (Fig. 3, B and C). There was no difference between male and female mice in these genes. To ensure that this also corresponded to a change in protein levels, we performed Western blot analysis with densitometry on Cnn1; we found an approximately twofold decrease in protein expression in the 8-wk mice (Fig. 3D).
We were interested in seeing whether these decreases would be visible by immunohistochemistry (IHC) in actin-positive cells. Titin was the muscle-related gene with the largest decrease by array (10x, Table 2) and was thus the most likely candidate for changes to be visible by IHC. We first confirmed this decrease by quantitative RT-PCR (Fig. 4A) before double staining tissue sections for titin and actin (Fig. 4B). We found that in control animals, titin stained the wall of actin-positive cells facing the vessel lumen; this was missing in 8-wk BMPR2 mice. Columnar epithelial cells showed nonspecific staining and were included to show that exposure was similar in control and 8-wk mice.
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80% decrease in BMPR2 protein by Western blot with densitometry (not shown) with siRNA to BMPR2 compared with scrambled. RNA from these cells was used for quantitative RT-PCR with primers for Cnn1 and Myh11. We found that both genes were reduced more than twofold (Fig. 5A). To confirm that this resulted in a change in protein, we performed Western blot for Cnn1 and found a strong decrease in Cnn1 protein expression as well (Fig. 5B). These suggest that the decrease in smooth muscle markers seen in transgenic mice was a direct result of altered BMP signaling rather than an indirect effect of developing pulmonary arterial hypertension (PAH).
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DISCUSSION |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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A decrease in angiogenesis genes is precisely the opposite of the effect found in hypoxia, in which angiogenesis is increased (4). A decrease in ability to produce new blood vessels, however, appears to be a prerequisite in some systems for the development of severe pulmonary hypertension (17), so this decrease may be relevant to disease progression.
Loss of smooth muscle markers was our most unexpected finding, involving every smooth muscle marker we checked. Our ability to replicate this through siRNA to BMPR2 in cell culture suggests that it is a relatively direct effect of loss of normal BMP signaling. This decrease apparently does not reflect a decrease in the number of cells; we previously published that the number of actin-positive cells appears to remain about the same or increase (18). We thus assume that this reflects a decrease in expression per cell; this was confirmed in the case of titin (Fig. 4). Since one function of the BMP pathway is to drive terminal differentiation, it seems possible that loss of normal BMP signaling results in loss of a fully differentiated phenotype in smooth muscle cells and thus a fundamentally defective vasculature.
We expected an increase in cytokines, based on our earlier work (19), but the apparent sex-specific difference was novel. The increase in the complement activation pathway is particularly interesting, in combination with the loss of smooth muscle differentiation pathways. To us, this lends credence to other theories about increased turnover of cells, with replacement by either circulating progenitors or fibroblasts (14, 16), especially in combination with the changes in endocytosis genes seen.
The combination of these results, with our earlier work and other published reports, leads us to the following broad hypothesis of pathogenesis in these transgenic mice as well as possibly in human IPAH. Loss of BMPR2 has two primary effects: loss of proper smooth muscle differentiation and loss of proper control over inflammation.
This loss of smooth muscle differentiation may be true in asymptomatic human BMPR2 mutation carriers, as suggested by their abnormal exercise tests (5, 12). This may also be entirely the effect we see in our SM22-rtTA x TetO7-BMPR2delx4+ transgenic mice; the increased pressures may be the result either of differences in mouse physiology or the result of a rather worse mutation than is generally seen in human patients. Our previous publication on Kv1.5 suggests that abnormal control of tone is at the heart of the pressures seen in our model (21).
Second, and perhaps more importantly for disease progression, these mice, and particularly female mice, have higher levels of a variety of immune-related genes. This does not result in a higher RV pressure in female mice. We suspect that this is because the presence of dysregulated cytokines is irrelevant in the absence of a second hit. In the presence of some other factor that results in increased inflammation, these increased cytokines, combined with the less fully developed vasculature, results in a runaway inflammatory process and clinical PAH.
Although this hypothesis is consistent with these results, we by no means claim it proven. It is, however, an interesting starting point for future studies that address the specific effects of smooth muscle cell differentiation and inflammation in pulmonary hypertension.
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GRANTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
* Y. Tada and S. Majka contributed equally to this work. 
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