Role Played by Prx1-Dependent Extracellular Matrix Properties in Vascular Smooth Muscle Development in Embryonic Lungs

Animals

Prx1 gene knockout mice were generated as described elsewhere. ³⁰ Polymerase chain reaction (PCR) was used to identify the three different expected genotypes: Prx1^WT (wild type), Prx1 heterozygous, and Prx1^null (homozygous-null). Genomic DNA from mouse tail was used in PCRs with the combination of primer pairs for the Prx1^WT allele and the Prx1^lacZ allele. All animal experiments were conducted in accordance with current National Institutes of Health (Bethesda, MD) guidelines.

Cell culture

C3H10T1/2 (10T1/2) cells (ATCC) were used as mesenchymal SMC precursor cells. ¹⁷ Except in the case of TGF-β stimulation experiments, cells were maintained in 10% fetal bovine serum containing Dulbecco's modified Eagle medium. Cells were cultured at 37^°C in a humidified atmosphere with 5% CO₂ until used for cellular experiments.

Transient transfection

Cytomegalovirus plasmid containing Prx1 construct ³⁰ and control vector were used to examine the downstream genes regulated by Prx1.

Decellularization of lungs

Intact fetal mouse lungs from E15.5 to D0 from Prx1^WT and Prx1^null mice were treated with hypotonic Tris buffer (PH 8) followed by hypertonic Tris buffer containing 1% Triton X-100 (pH 8). ³¹ After decellularization, lung scaffolds were processed for characterization or left unfixed for biomechanical testing. Intact lungs were used as controls for decellularization experiments.

Polyacrylamide (PAA) gels

PAA gels of variable stiffness were prepared on glass coverslips using an established method. ³² After being treated with sulfo-SANPAH (Thermo Fisher Scientific, Waltham, MA), collagen I (0.2 mg/mL; Advanced Biomatrix, Carlsbad, CA) was crosslinked to the gels before cells were plated.

Synthetic 3-D scaffolds (electrospun hyaluronic acid scaffolds)

Methacrylated hyaluronic acid (MeHA) was synthesized as described elsewhere.^33,34 Briefly, MeHA was cospun with the carrier polymer PEO (molecular weight, 900 kDa; Sigma-Aldrich, St. Louis, MO) as described elsewhere,^34,35 and fiber size and view morphology were quantified by scanning electron microscopy (SEM; 7500F HRSEM; Jeol, Glen Ellyn, IL). After polymerization was accomplished by exposing ~10 mW/cm ² 365-nm light (Omnicure S1000 UV Spot Cure System; Exfo, Quebec City, QC), scaffolds were immersed in deionized H₂O and washed to reach equilibrium swelling. The mechanical properties of MeHA gels were controlled by percent methacrylation or concentration of polymer used, and their stiffness was determined by atomic force microscopy (AFM).

Immunohistochemistry

The following primary antibodies were used: anti-Fbn1 and anti-Fbn2 antibodies (gift from R. Mecham, Washington University, St. Louis, MO), anti-tropoelastin antibody (Abcam, Cambridge, UK), anti-α-smooth muscle actin (SMA) antibody (Sigma-Aldrich), anti-γ-SMA antibody (Santa Cruz Biotechnology, Dallas, TX), anti-SM22α antibody (Abcam), anti-smooth muscle myosin heavy chain (SMMHC) antibody (Abcam), anti-TGF-β1 antibody (Promega, Madison, WI), and anti-TGF-β2 antibody (R&D Systems, Minneapolis, MN). For staining controls, equivalent concentrations of isotype-matched immunoglobulin (Ig) G were used. Sections were blocked for endogenous peroxidase activity, treated for antigen retrieval, and blocked for nonspecific antibody binding before being incubated with primary antibodies. The following dilutions were used for primary antibodies: Fbn1 and Fbn2 (1:100), tropoelastin (1:150), α-SMA (1:400), γ-SMA (1:50), SM22α (1:100), and SMMHC (1:100). Antigen detection was performed using species-specific secondary antibodies in conjunction with a Vectastain Elite ABC Kit (Vector Laboratories, Burlingame, CA), followed by the 3,3′-diaminobenzidine reaction (Vector Laboratories), counterstaining with hematoxyline (Vector Laboratories), and processing for bright-field imaging (90i microscope; Nikon, Tokyo).

Immunofluorescence staining

Cells or scaffolds were fixed in 4% paraformaldehyde and then permeabilized with 0.2% Triton X-100–containing phosphate-buffered saline. Next, materials were treated with blocking buffer and incubated with the following primary antibodies. To examine SMC differentiation, antibodies specific for α-SMA, γ-SMA, SM22α, and SMMHC were used. For detection of ECM components, antibodies specific for Fbn1, Fbn2, tropoelastin, and TGF-β1 were used. Species-specific fluorescent-conjugated antibodies (Molecular Probes, Thermo Fisher Scientific) were used for antigen detection. Stained materials were examined using a 90i fluorescence microscope (Nikon) or a LSM 710 two-photon confocal microscope (Zeiss, Jena; CDB Imaging Core at the University of Pennsylvania, Philadelphia, PA).

Electron microscopy

Western immunoblotting

Protein was extracted from lung tissue and concentrations were determined using the Pierce Protein Assay Reagent (Thermo Fisher Scientific). Ten micrograms of protein was separated on an 8%–16% Tris-HCl sodium dodecyl sulfate–PAA gel and then transferred to polyvinylidene fluoride membranes. After being incubated with blocking buffer, membranes were incubated with primary antibodies using the following: Fbn1 and Fbn2 (gift of R. Mecham), tropoelastin (Abcam), TGF-β1 (Promega), phosphorylated Smad2 (Cell Signaling Technology, Danvers, MA), and total Smad2/3 (Cell Signaling Technology). After incubation with species-specific secondary antibodies conjugated with horseradish peroxidase, protein was visualized using an enhanced chemiluminescence kit and examined using a Fuji imager (Fujifilm, Tokyo/GE Healthcare, Chalfont St Giles, UK). β-Actin (Sigma-Aldrich) was used as a loading control.

Real-time quantitative PCR

Total RNA was extracted from lung tissue or cells, and complementary DNA (cDNA) was then synthesized and processed for SYBR green–based real-time quantitative PCR analysis using the LightCycler 480 instrument (Roche, Basel). All target gene expressions were normalized to a housekeeping gene.

Stiffness measurements

TGF-β stimulation experiments

Cells were incubated with serum-free medium overnight before being treated with recombinant TGF-β1 (Cell Signaling Technology). ¹⁷ Either cells were fixed for immunostaining or total RNA was harvested for real-time PCR.

TGF-β function-blocking experiments

Statistics

All experiments were performed in triplicate unless otherwise stated. Statistical analysis was performed using Prism software (GraphPad, San Diego, CA) to compare the groups. One-way analysis of variance with the Newman-Keuls posttest was used to compare more than two groups, and the unpaired Student t test was used to compare two groups. A P value of less than 0.05 was considered significant. All values are shown as means ± SD unless otherwise specified.

Detailed methods are available in the supplement.

Prx1 regulates expression of elastic ECM components during lung development

We first examined expression of elastic ECM components in Prx1^WT and Prx1^null lungs to determine whether Prx1 regulates expression of the major elastic ECM components within developing lungs. We focused on E15.5 to newborn (D0) stages since the expression level of lung ECM components is higher during the late gestational period. The expression of tropoelastin, Fbn1, and Fbn2 was significantly lower in Prx1^null lungs than in Prx1^WT lungs at both the protein (Figs. 1A, 1B, S1; Figs. S1–S3 are available online) and the messenger RNA (mRNA; Fig. 1C) level. Immunohistochemical studies also suggested that tropoelastin protein expression was significantly reduced around the developing vessels and in the adjacent mesenchyme (Fig. 1A, bottom) as well as around airways. Collectively, these data suggest that Prx1 controls the composition of the elastic ECM in developing lung vessels. To investigate further, we examined the ultrastructure of vessels in E17.5 embryo lungs using TEM. Strikingly, the results showed that, whereas Prx1^WT lungs possessed a dense, solid elastic lamina, Prx1^null lungs displayed a thinner, patchy, discontinuous elastic lamina (Fig. 1D). To examine whether Prx1 directly affects the expression of these elastic ECM components, a full-length Prx1 cDNA-containing plasmid ³⁰ was transiently transfected to 10T1/2 cells, a mesenchymal precursor cell line, and mRNA expression of these elastic ECM components was examined (Fig. S1B–S1D). Overexpression of the Prx1 gene (Fig. S1B) was followed by increased mRNA expression of tropoelastin (Fig. S1C; 2.968e6 ± 1.175e7 vs. 0.9973e7 ± 0.9e9) and Fbn1 (Fig. S1D; 4.961e4 ± 1.566e4 vs. 3.234e4 ± 0.150e4). In support of the above results, because the promoter regions of tropoelastin, Fbn1, and Fbn2 contain conservative Prx1-binding sequences, it is plausible that Prx1 directly binds to and regulates expression of these elastic ECM components.

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Figure 1.

Prx1 controls expression of tropoelastin (TE), fibrillin (Fbn) 1, and Fbn2 during lung vascular development. A, Immunohistochemical staining for TE, Fbn1, and Fbn2 in lung sections derived from E15.5 Prx1^WT (left) and Prx1^null (right) mice. Arrows indicate developing vessels, in which TE expression is observed in Prx1^WT mice but not in Prx1^null mice. Brown color indicates positive staining; hematoxylin staining in blue is nuclear counterstaining. B, Western immunoblotting for Fbn1, Fbn2, and TE protein in Prx1^WT and Prx1^null lungs isolated at E17.5. β-Actin was used as a loading control. C, Real-time polymerase chain reaction for TE and Fbn2 messenger RNA expression in Prx1^WT and Prx1^null lungs isolated at E17.5. Target gene expression was normalized to GAPDH expression. One asterisk indicates P < 0.05, and two asterisks indicate P < 0.01. D, Semithin section images (top) and transmission electron microscope images (bottom) of pulmonary arteries of E17.5 Prx1^WT (left) and Prx1^null (right) mouse lungs. Arrows indicate the internal elastic lamina. Scale bars = 5 μm. Western immunoblotting was performed using at least three different litters. WT: wild type; EC: endothelial cell; SMC: smooth muscle cell.

Prx1null lungs have defective vascular SMC differentiation

Since elastic ECM properties are essential for vascular SMC differentiation,^13,14 we examined vascular SMC differentiation in Prx1^WT and Prx1^null lungs. Immunostaining of E15.5 and D0 lungs for α-SMA (Fig. 2A, left; Fig. 2B), γ-SMA (Fig. 2A, middle), and SM22α (Fig. 2A, right) showed that SMCs within developing pulmonary vessels in Prx1^WT mice strongly expressed SMC markers (i.e., α-SMA, γ-SMA, and SM22α) and possessed a spindle-shaped morphology (Fig. 2B). In contrast, Prx1^null lungs expressed reduced levels of the SMC markers examined, and vascular wall cells appeared to possess a rounded, more primitive morphology (Fig. 2B). Furthermore, whole-mount immunostaining of E17.5 lungs with α-SMA clearly showed that 3-D development and differentiation of vascular SMCs was deficient in Prx1^null lungs compared with Prx1^WT lungs (Fig. 2C). Consistent with the staining results, mRNA expression of α-SMA (Fig. 2D), SM22α (Fig. 2E), and SMMHC (Fig. 2F) in Prx1^null lungs at this stage was significantly reduced compared with that in Prx1^WT lungs.

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Figure 2.

Prx1 promotes vascular smooth muscle cell differentiation. A, Immunohistochemistry for α–smooth muscle actin (SMA; left), γ-SMA (middle), and SM22α (right) in lungs of E15.5 (top) and D0 (bottom) Prx1^WT and Prx1^null mice. Positive staining by 3,3′-diaminobenzidine (DAB) reaction is brown; nuclear counterstaining is blue. Arrows point to vessel walls. Scale bars = 20 μm. B, Higher magnification of α-SMA immunohistochemistry on D0 Prx1^WT and Prx1^null mice lung sections. Positive staining by DAB reaction is brown; nuclear counterstaining is blue. Arrows point to the smooth muscle layer of vessels. Scale bar = 20 μm. C, 3-D projection images of whole-mount lung immunofluorescence staining for α-SMA in E17.5 lungs from Prx1^WT (left) and Prx1^null (right) mice. Scale bar = 20 μm. D–F, Real-time polymerase chain reaction for α-SMA (D), SM22α (E), and smooth muscle myosin heavy chain (SMMHC; F) messenger RNA expression in E17.5 lungs isolated from Prx1^WT and Prx1^null mice. Target gene expression was normalized to GAPDH expression. Three asterisks indicate P < 0.001. WT: wild type.

TGF-β signaling was downregulated in Prx1null lungs

It was recently reported that ECM properties regulate the activity of TGF-β1 signaling,^19,38 a potent inducer of vascular SMC differentiation. ⁸ Since Prx1-deficient mice demonstrated compromised ECM composition and deficient SMC differentiation, we examined TGF-β expression and signaling in Prx1^WT and Prx1^null lungs. Immunohistochemistry on Prx1^WT and Prx1^null lung sections showed that both TGF-β1 and TGF-β2 strongly localize in and around the developing vessels in Prx1^WT lungs (Fig. 3A), while in Prx1^null lungs both TGF-β1 and TGF-β2 are less expressed in the vicinity of vessels and appear frequently in parenchyma (Fig. 3A). Interestingly, Western blotting and real-time PCR showed that total TGF-β1 expression in Prx1^WT and Prx1^null lungs was not significantly different at the protein and mRNA levels (Fig. 3B, 3C); however, phosphorylation of the TGF-β downstream signaling mediator Smad2 (Fig. 3D) and expression of the TGF-β-regulated protein plasminogen activator inhibitor type 1 (PAI-1; Fig. 3E) were reduced in Prx1^null lungs compared with Prx1^WT lungs. These results indicate that Prx1 regulates TGF-β signaling, potentially because of its effects on lung matrix.

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Figure 3.

Prx1 regulates transforming growth factor (TGF)–β signaling in embryonic lungs. A, Immunohistochemical staining for TGF-β1 and TGF-β2 in lung sections derived from E15.5 (top) and D0 (bottom) Prx1^WT (left) and Prx1^null (right) mice. Brown indicates positive staining of antibodies. While TGF-β localization is restricted to the developing vessel area (arrows) in Prx1^WT lungs, in Prx1^null lungs TGF-β expression is observed in areas other than the vessel wall vicinity (arrowheads). Hematoxylin staining (blue) is used for nuclear counterstaining. Scale bar = 40 μm. B, Western blotting for TGF-β1 protein expression in E17.5 lungs from Prx1^WT and Prx1^null mice. C, Real-time polymerase chain reaction (PCR) for TGF-β1 messenger RNA (mRNA) expression in E17.5 lungs from Prx1^WT and Prx1^null mice. D, Western blotting for phosphorylated Smad2 (P-Smad2; top) and total Smad 2/3 (middle) in E17.5 lungs from Prx1^WT and Prx1^null mice. E, Real-time PCR for plasminogen activator inhibitor type 1 (PAI-1) mRNA expression in E17.5 lungs from Prx1^WT and Prx1^null mice. For Western immunoblotting (B, D), β-actin was used as a loading control. For mRNA expression by real-time PCR (C, E), target gene expression was normalized to GAPDH. Two asterisks indicate P < 0.01. WT: wild type.

Prx1 controls the biomechanical properties of lung ECM

Since our data suggested a strong link between Prx1-dependent formation of elastic ECM and development of the lung vasculature, we further examined the ECM in Prx1^WT and Prx1^null lungs. To characterize the ECM derived from Prx1^WT and Prx1^null lungs and to elucidate the role played by Prx1-dependent ECM properties in vascular SMC differentiation, we developed a system to decellularize fetal mouse lungs in which proper removal of cells from the tissue leaves behind a complex ECM and associated proteins, including growth factors. ³⁹ Decellularized scaffolds have proven to be useful tools for studying cell signaling in an appropriate 3-D ECM environment as well as for tissue engineering studies designed for clinical applications. ⁴⁰ Therefore, we used the decellularized scaffolds to examine the architectural, biomechanical, and biochemical properties of the ECM in Prx1^null lungs.

First, we used hematoxylin nuclear staining on intact and decellularized frozen lung sections and confirmed that cells were removed successfully by the decellularization process (Fig. 4A). Next, we immunostained intact and decellularized Prx1^WT lungs to determine whether decellularized scaffolds retain ECM components. The results showed that the decellularized scaffolds clearly retained Fbn2, tropoelastin, and TGF-β1 (Fig. 4B) as well as basement component laminin (Fig. S2). We also examined the stiffness of intact and decellularized Prx1^WT lung scaffolds (Fig. 4C). As other groups have observed, ⁴¹ decellularization leads to reduced stiffness (1,486 ± 1,029 Pa) compared with that in intact lungs (2,577 ± 670 Pa), likely due to loss of cellular tension. These results confirmed that cells were successfully removed while ECM components were retained.

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Figure 4.

Fetal lung extracellular matrix (ECM) is well preserved after decellularization. A, Hematoxylin staining for cell nuclei on frozen sections derived from intact (left) and decellularized (right) E17.5 Prx1^WT lungs to confirm the removal of cells from tissue. Scale bar = 20 μm. B, Immunofluorescence staining for fibrillin (Fbn) 2 (top), tropoelastin (TE; middle), and transforming growth factor (TGF)–β (bottom) within intact (left) and decellularized (right) Prx1^WT lungs to examine the preservation of elastic ECM components in decellularized ECM scaffolds. Scale bar = 20 μm. C, Stiffness (Young's modulus) of intact and decellularized E17.5 Prx1^WT lungs, determined by indentation testing. WT: wild type.

We used these lung scaffolds to characterize Prx1-dependent properties of the lung ECM. Consistent with the staining of intact tissue (Fig. 1), immunostaining showed that Fbn2 and tropoelastin were decreased in Prx1^null scaffolds compared with Prx1^WT lung scaffolds (Fig. 5A). Furthermore, the localization of TGF-β1 in the ECM was diffuse in Prx1^null scaffolds, whereas it was strongly deposited in the vicinity of the airway and vessels in Prx1^WT lung scaffolds (Fig. 5A). We also examined the stiffness of E17.5 lung scaffolds by indentation testing with a 550-μm-diameter probe (Fig. 5B). Prx1^null lung scaffolds (398 ± 209 Pa) were significantly softer than Prx1^WT lung scaffolds (1486 ± 1029 Pa; Fig. 5B). We then used SEM and TEM studies to examine the ultrastructural architecture of the ECM. SEM images showed that Prx1^WT scaffolds possessed a well-assembled ECM, bearing a variety of sizes of interlinked fibrillar networks (Fig. 5C, left). In contrast, Prx1^null lung scaffolds possessed a less fibrillar and more disorganized structure (Fig. 5C, right). TEM images were consistent with SEM level analysis, showing that E17.5 Prx1^null lung scaffolds possessed a less developed fibrillar network (Fig. 5D, right) compared with that of the Prx1^WT scaffolds (Fig. 5D, left). Fiber sizes constructing lung ECM scaffolds were measured using XL20 ESEM imaging software (Philips) and ImageJ. We found that fiber sizes of Prx1^null ECM were significantly different from those of Prx1^WT ECM. Prx1^WT ECM fibrils (42.7 ± 30.5 nm) were on average of smaller diameter than Prx1^null ECM fibrils (69.8 ± 21.5 nm) but had a wider range of sizes (Fig. S3), which is consistent with the ultrastructural observations (Fig. 5C, 5D). These results clearly indicate that Prx1 controls ECM architecture and physical properties by regulating ECM composition.

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Figure 5.

Prx1 controls the biophysical properties of fetal lung extracellular matrix (ECM). A, Immunofluorescence staining for fibrillin (Fbn) 2 (top), tropoelastin (TE; middle), and transforming growth factor (TGF)–β (bottom) in Prx1^WT (left) and Prx1^null (right) lung scaffolds at E17.5. Arrows indicate decellularized vessels. Scale bar = 20 μm. B, Stiffness (Young's modulus) of decellularized E17.5 lung scaffolds from Prx1^WT and Prx1^null mice, determined by indentation testing. Two asterisks indicate P < 0.01. C, Scanning electron photomicrographs of decellularized lung scaffolds from Prx1^WT and Prx1^null mice demonstrating the ultrastructure and fine architecture of lung ECM. D, Transmission electron micrographs of decellularized lung scaffolds from Prx1^WT and Prx1^null mice demonstrating the 2-D ultrastructure and ECM fiber formation of lung ECM. WT: wild type.

ECM stiffness and TGF-β signaling tightly cooperate in vascular SMC differentiation in 2-D and 3-D synthetic substrates

To determine whether Prx1-dependent ECM stiffness regulates vascular SMC differentiation and whether TGF-β signaling is involved in this process, we first confirmed that 10T1/2 mesenchymal precursor cells differentiate into SMC-like cells in response to TGF-β1.^17,42 An immunocytochemical study showed that 10T1/2 cells respond to the TGF-β1 stimulation and induce SMC differentiation markers, such as α-SMA, γ-SMA, SM22α, and SMMHC (Fig. 6A), while cells not stimulated or cotreated with TGF-β and LY364947, an inhibitor of the TGF-β type I receptor, exhibited low-level expression (Fig. 6A). Activation of TGF-β signaling was confirmed by demonstrating increased expression of PAI-1, a well-recognized target of the canonical TGF-β pathway (Fig. 6B, bottom). Conversely, cotreatment of 10T1/2 cells with TGF-β1 and LY364947 resulted in PAI-1 expression at basal levels (Fig. 6B, bottom). In parallel, mRNA expression levels of α-SMA and γ-SMA were significantly increased in response to TGF-β stimulation but remained at basal levels with inclusion of the inhibitor (Fig. 6B, top, middle). Western immunoblotting (Fig. 6C) for α- and γ-SMA together with phosphorylated Smad2 showed the same response at the protein level as at the mRNA level. These studies confirm that 10T1/2 cells have the capacity to adopt a SMC-like phenotype in response to TGF-β signaling via the canonical pathway.

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Figure 6.

10T1/2 cell differentiation to a smooth muscle phenotype is transforming growth factor (TGF)-β dependent. A, Immunostaining for α-smooth muscle actin (SMA), γ-SMA, SM22a, and smooth muscle myosin heavy chain (SMMHC; all positive reactions are green) in 10T1/2 cells, with or without TGF-β1 treatment, in the presence or absence of a TGF-β type I receptor (TGF-βRI) inhibitor (LY364947). Scale bar = 10 μm. B, α- and γ-SMA messenger RNA expression in 10T1/2 cells after stimulation with TGF-β1, with or without LY364947. Plasminogen activator inhibitor type 1 (PAI-1) was used as a positive control for canonical TGF-β signaling activation. Signals were normalized to GAPDH. C, Western immunoblotting for α- and γ-SMA expression in 10T1/2 cells after stimulation with TGF-β1, with or without a TGF-βRI inhibitor (SB431542). Phosphorylation of Smad2 (P-Smad2) was examined as a marker of canonical TGF-β signaling. GAPDH was used as a loading control.

We used 10T1/2 cells cultured on stiffness-controlled PAA gels in 2-D culture (Fig. 7A) to determine whether Prx1-dependent biophysical properties of the ECM are critical factors in controlling mesenchymal cell differentiation and whether Prx1-dependent ECM stiffness alone is sufficient or whether TGF-β signaling needs to be involved in this process.^32,38 By adjusting the concentration of bis-acrylamide within the gels, the bulk physical properties of these substrates were designed to match the stiffness of either Prx1^WT (i.e., 1,500-Pa) or Prx1^null (i.e., 500-Pa) lung scaffolds. Expression of the SMC differentiation markers α-SMA, SM22α, and SMMHC was examined following culture of 10T1/2 cells on these PAA gels. Fluorescence microscopy images showed that in the absence of TGF-β1, 10T1/2 cells cultured on the stiffer substrate developed a relatively elongated morphology and more marker expression (Fig. 7B); with TGF-β1 treatment, characteristics of the SMC-like phenotype, such as cell elongation and SMC marker expression, were further increased (Fig. 7B, top vs. bottom). In contrast, 10T1/2 cells cultivated on the softer Prx1^null-stiffness substrate retained mesenchymal-like morphology and expressed low levels of SMC markers with or without TGF-β1 stimulation (Fig. 7B) compared with cells cultured on stiffer Prx1^WT-stiffness substrates. To quantify the morphological change of cell shape, the aspect ratio was measured (Fig. 7C). Supporting the cell morphological observation of immunocytochemistry, 10T1/2 cells cultured on stiffer substrates had a higher aspect ratio (more elongated SMC-like morphology), and the effect of TGF-β on the aspect ratio was enhanced by a stiffer substrate. Real-time PCR for SMC differentiation markers (α-SMA, SM22α, and SMMHC) demonstrated that mRNA expression of the SMC differentiation markers was induced by stiffer substrates and enhanced by TGF-β signaling (Fig. 7D–7F), which is parallel to the results of immunostaining. We also noted that among the SMC differentiation markers we examined, expression of α-SMA, an initial SMC differentiation marker, showed the most significant differences between the culture settings (Fig. 7B, 7D) compared with the expression of SM22α (Fig. 7B, 7E) and SMMHC (Fig. 7B, 7F), which are more mature SMC markers and are significantly induced only in the cells seeded on the 1,500-Pa substrates with TGF-β stimulation. This indicates that the 24-hour biophysical and biochemical stimulation leads to an initial stage of SMC differentiation from embryonic precursor cells but is not sufficient to achieve full differentiation. Taken together, these results indicate that ECM stiffness is a key determinant in controlling 10T1/2 cell differentiation to SMCs in conjunction with TGF-β stimulation. Our results also indicate that Prx1-dependent ECM stiffness alone is not sufficient to support mesenchymal cells to differentiate to the smooth muscle phenotype and requires coordination with TGF-β signaling. For this event, Prx1 likely promotes SMC differentiation via its effect on both the biophysical and the biochemical properties of the developing lung ECM.

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Figure 7.

Transforming growth factor (TGF)–β–dependent 10T1/2 cell differentiation tightly interacts with Prx1-dependent extracellular matrix (ECM) stiffness. A, Schema for the stiffness-controlled polyacrylamide (PAA) gel 2-D culture system. B, 10T1/2 cells were cultured on PAA gel 2-D substrates with a stiffness of either 500 (top) or 1,500 (bottom) Pa, which is parallel to the E17.5 Prx1-lung stiffness, in the presence or absence of TGF-β stimulation. Smooth muscle cell differentiation was evaluated by expression of α-smooth muscle actin (SMA; left, green), SM22α (middle, green), and smooth muscle myosin heavy chain (SMMHC; right, red). 4′,6-Diamidino-2-phenylindole dihydrochloride (blue) was used for nuclear staining. Scale bar = 10 μm. C, Aspect ratio of cells cultured on 2-D PAA gels under the above-described conditions was examined to quantify cell morphology change. D–F, Real-time polymerase chain reaction results for 10T1/2 cells cultured on 2-D PAA gels of either 500- or 1,500-Pa stiffness with or without TGF-β stimulation. α-SMA (D), SM22α (E), and SMMHC (F) messenger RNA expression was examined and normalized by GAPDH or cyclophilin A (cyclo A) expression. One asterisk indicates P < 0.05, and two asterisks indicate P < 0.01. G, 3-D projection of confocal images of F-actin-stained 10T1/2 cells cultured in soft and stiff 3-D methacrylated hyaluronic acid synthetic scaffolds. F-actin is red, nucleus is blue. Scale bar = 20 μm.

To further examine the effect of ECM stiffness on cell differentiation under more physiological conditions, we cultured 10T1/2 cells within chemically engineered 3-D substrates of different stiffness. Electrospun, MeHA gels were coated with type 1 collagen before being seeded with cells. ³⁵ The stiffness of these synthetic scaffolds was measured by AFM after the scaffolds were rehydrated to determine the physiological stiffness. We used MeHA gels with stiffness of ≃1,000 Pa to represent a soft ECM and of ~3,000–5,000 Pa to represent a stiffer ECM for cell culture. ³⁴ 3-D projections of confocal microscope images outlined by cellular F-actin staining showed that on soft 3-D scaffolds 10T1/2 cells sporadically aggregated and failed to differentiate or assemble functional vascular SMC-like networks (Fig. 7G). Within the stiffer 3-D microenvironment, however, cells grew into the MeHA substrate and demonstrated prominent network formation with elongated cell morphologies (Fig. 7G), consistent with functional SMC differentiation. Although the synthetic scaffold stiffness is not identical to the Prx1^WT and Prx1^null lung scaffold stiffness because of technical limitations, these results suggest that 3-D stiffness cues have a stronger effect on 10T1/2 cell to SMC functional differentiation than 2-D cues.

Prx1-dependent ECM regulates SMC differentiation in a TGF-β-dependent manner

Finally, we asked whether acellular Prx1^WT and Prx1^null lung scaffolds differentially support SMC differentiation. We seeded and cultured 10T1/2 cells in decellularized lung scaffolds derived from Prx1^WT and Prx1^null mice. After 5 days in culture, cells had clearly migrated into both Prx1^WT and Prx1^null scaffolds, yet their organization and morphology greatly differed: recolonization of vascular structures was clearly evident for 10T1/2 cells in Prx1^WT lung scaffolds (Fig. 8A, left) compared with the Prx1^null scaffolds, within which cells appeared to be randomly distributed (Fig. 8A, right). Furthermore, expression of α-SMA of seeded cells in Prx1^WT scaffolds was greater than that in Prx1^null scaffolds (Fig. 8A, 8B). SEM images for ultrafine structures support the immunostaining results that 10T1/2 cells on Prx1^WT scaffolds possess a more elongated SMC-like phenotype compared with those cultivated on Prx1^null scaffolds (Fig. 8E). Consistent with the idea that Prx1-dependent ECM properties play a major role in mesenchymal cell to SMC differentiation, we tested whether 10T1/2 cell differentiation within Prx1^WT scaffolds is simply dependent on the mechanical properties of the ECM or is also dependent on the biochemical properties of the ECM, TGF-β. We pretreated Prx1^WT scaffolds with a function-blocking anti-TGF-β1 antibody and then seeded 10T1/2 cells. Whereas IgY-treated controls adopted the expected elongated and differentiated phenotype, including expression of α-SMA (Fig. 8C), cultures treated with anti-TGF-β1 IgY failed to colonize scaffolds appropriately, adopted a nondifferentiated morphology, and expressed reduced levels of α-SMA (Fig. 8C, 8D). Collectively, these results strongly suggest that Prx1 plays a critical role in lung vascular SMC differentiation during embryonic development by coordinating the biophysical and biochemical properties of the ECM.

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Figure 8.

Prx1-dependent extracellular matrix regulates lung vascular smooth muscle cell differentiation. A, Immunofluorescence staining for α-smooth muscle actin (SMA; green) in 10T1/2 cells seeded within E17.5 Prx1^WT (left) or Prx1^null (right) lung scaffolds. Insets show enlarged 10T1/2 cell images. Nuclei stained blue by 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI) staining. Scale bar = 20 μm. B, Fluorescence intensity of α-SMA immunostaining on frozen sections of 10T1/2 cells cultured in Prx1^WT and Prx1^null scaffolds (A) was quantified using ImageJ. One asterisk indicates P < 0.05. C, Immunofluorescence staining for α-SMA (green) of 10T1/2 cells seeded in Prx1^WT scaffolds, treated with either control immunoglobulin (Ig) Y (left) or transforming growth factor (TGF)-β function-blocking IgY (right). Nuclei stained blue by DAPI. Scale bar = 20 μm. D, Fluorescence intensity of α-SMA immunostaining on frozen sections of 10T1/2 cells cultured in Prx1^WT scaffolds with or without function-blocking TGF-β antibody (C), quantified using ImageJ. Two asterisks indicate P < 0.01. E, Scanning electron photomicrographs showing the ultrastructure of 10T1/2 cells in Prx1^WT (left) and Prx1^null (right) lung scaffolds after 5 days in culture. Cells are highlighted in red. Scale bar = 5 μm. WT: wild type.