Collagen-binding proteoglycan fibromodulin can determine stroma matrix structure and fluid balance in experimental carcinoma
- Åke Oldberg*,
- Sebastian Kalamajski*,
- Alexei V. Salnikov†,‡,§,
- Linda Stuhr¶,
- Matthias Mörgelin‖,
- Rolf K. Reed¶,
- Nils-Erik Heldin**, and
- Kristofer Rubin†,††
- *Department of Experimental Medical Sciences, BMC, B-12, and
- ‖Department of Clinical Sciences, BMC B14, University of Lund, SE-221 84 Lund, Sweden;
- †Department of Medical Biochemistry and Microbiology, Uppsala University, BMC, Box 582, SE-751 23 Uppsala, Sweden;
- ‡Oncology Clinic, University Hospital Lund, SE-221 85 Lund, Sweden;
- ¶Department of Biomedicine, University of Bergen, N-5009 Bergen, Norway and
- **Department of Genetics and Pathology, Uppsala University Hospital, Rudbeck Laboratory, SE-751 85 Uppsala, Sweden
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Edited by Erkki Ruoslahti, Burnham Institute for Medical Research, Santa Barbara, CA, and approved July 16, 2007 (received for review March 7, 2007)
Abstract
Research on the biology of the tumor stroma has the potential to lead to development of more effective treatment regimes enhancing the efficacy of drug-based treatment of solid malignancies. Tumor stroma is characterized by distorted blood vessels and activated connective tissue cells producing a collagen-rich matrix, which is accompanied by elevated interstitial fluid pressure (IFP), indicating a transport barrier between tumor tissue and blood. Here, we show that the collagen-binding proteoglycan fibromodulin controls stroma structure and fluid balance in experimental carcinoma. Gene ablation or inhibition of expression by anti-inflammatory agents showed that fibromodulin promoted the formation of a dense stroma and an elevated IFP. Fibromodulin-deficiency did not affect vasculature but increased the extracellular fluid volume and lowered IFP. Our data suggest that fibromodulin controls stroma matrix structure that in turn modulates fluid convection inside and out of the stroma. This finding is particularly important in relation to the demonstration that targeted modulations of the fluid balance in carcinoma can increase the response to cancer therapeutic agents.
Despite advances in our knowledge of causative factors and genetic changes during development of the malignant genotype, treatment results and mean survival time for patients suffering from advanced cancer have improved only marginally. In recent years, partly in response to disillusion with the available cytotoxic cancer chemotherapeutic agents, attention has turned to the stromal elements of tumors. Abnormal blood vessels and connective tissue cells that produce a fibrotic collagen-rich matrix characterize the stroma of a carcinoma (1–4). Invasiveness and growth of malignant cells are affected by interactions with the stroma (5), and in this respect a carcinoma resembles an inflammatory lesion (6) or a wound that will not heal (7). A mathematical model of cancer invasion recently suggested that a heterogeneous extracellular matrix (ECM) with varying ECM component densities selects aggressive tumor cell phenotypes, whereas tumors with a homogenous ECM are less aggressive and invasive (8).
Carcinomas have a pathologically elevated interstitial fluid pressure (IFP) indicating a transport barrier between tumor tissue and blood (9–11). Indeed, pharmaceutical treatments that lower IFP in experimental carcinoma (12–14) and human cancers (ref 15 and references therein) also increase efficacy of cytostatic treatment. Treatment of mice carrying a xenografted human anaplastic thyroid carcinoma (KAT-4) with the soluble TGF-β receptor type II-murine Fc:IgG2A chimeric protein (Fc:TβRII), which inhibits TGF-β1 and -β3, lowers IFP, reduces plasma protein leakage, and enhances the anti-tumor effect of doxorubicin (16, 17). Microarray analyses (Affymetrix, Santa Clara, CA) of Fc:TβRII-treated KAT-4 carcinomas revealed down-regulations predominantly of macrophage-related genes (17). Furthermore, inhibition of IL-1 reduces IFP in these carcinomas (17). These findings indicate a role of inflammation in the generation of an elevated IFP. Tortuous and leaky blood vessels, and absence of lymph drainage in tumors have also been suggested to be involved in the generation of the elevated tumor IFP. Inhibition of plasma protein leakage by interfering with vascular endothelial growth factor reduces IFP in experimental carcinomas (18, 19) and human rectal carcinoma (15). Little is known, however, about how the structure and composition of the ECM in the stroma influence carcinoma IFP.
The small leucine-rich repeat proteoglycan fibromodulin has a known role in collagen assembly and maintenance (20). Fibromodulin is expressed in dense regular connective tissues, such as tendons, ligaments, and cartilage, but is absent or expressed at low levels in skin, bone, and visceral organs (21, 22). Fibromodulin controls collagen matrix structure in tendons and ligaments, and deficient mice show altered tissue organization with fewer and structurally abnormal collagen fiber bundles. The collagen fibrils are thinner (23), tendons and ligaments have reduced tensile strength, and the animals develop knee-joint instability and osteoarthritis (24–26). Fibromodulin mRNA has been detected in a variety of clinical malignancies, such as lung, breast, and prostate carcinomas (27–29). Here, we report that fibromodulin has a role in regulating stroma ECM structure and fluid balance in carcinoma.
Results
We were led to fibromodulin by the finding that in addition to lower expression of macrophage-related genes, Fc:TβRII significantly down-regulated mRNA encoding fibromodulin. Notably, mRNA levels encoding other ECM proteins, such as collagen type I, fibronectin, and decorin were unchanged [supporting information (SI) Table 3]. Initially, we determined that fibromodulin showed a wide-ranging expression in experimental carcinomas, such as chemically induced rat mammary carcinoma, mouse RIP-Tag insulinoma, transplantable syngeneic colonic carcinoma (PROb), and KAT-4 carcinoma growing as a xenograft in nude mice (Fig. 1). Staining was restricted to the stroma compartment in the carcinomas. This finding is consistent with the absence of fibromodulin in normal murine skin, thyroid, and colon (SI Fig. 5 A–C), which is in agreement with the reported distribution of fibromodulin (21, 22). Cultured KAT-4 and PROb cells did not express fibromodulin mRNA or protein (data not shown). Quantitative PCR analyses showed that treatment of KAT-4 carcinomas with Fc:TβRII reduced fibromodulin mRNA by ≈50% (Fig. 1 D). Immunohistochemical analyses showed that fibromodulin protein expression was also reduced (Figs. 1 E–F). No staining was seen in carcinomas from Fmod-null mice (Fig. 1 H).
Expression of fibromodulin in carcinomas. (A–C) Immunoperoxidase staining of fibromodulin in PROb rat colonic carcinoma (A), DMBA-induced rat breast carcinoma (B), and mouse RIP-Tag insulinomas (C). Quantitative real-time PCR of KAT-4 tumors from mice treated with control IgG2A or Fc:TβRII. (D) The difference in mRNA levels between treated and untreated tumors was significant (P < 0.05, n = 3). (E and F) Staining of fibromodulin in control IgG2A-treated (E) and Fc:TβRII-treated (F) mice carrying KAT-4 carcinomas. (G and H) Fibromodulin stainings of KAT-4 carcinoma xenografted in WT (G) and Fmod-null (H) mice. Mice carrying KAT-4 carcinoma were treated with vehicle or Fc:TβRII (10 mg/kg) 10 days before killing (16). Tumors were frozen, sectioned, stained with a rabbit anti-bovine fibromodulin antiserum, and counterstained with hematoxylin (23). (Scale bar, 100 μm.)
Fmod-null nude/nude mice were used further to study the role of fibromodulin in tumors. Human KAT-4 carcinoma cells were transplanted s.c. in WT, Fmod-null, and heterozygous nude/nude littermates. Tumor take, length of lag phase, and temporal increase in external sizes were identical in the three genotypes. Total carcinoma weights 31 days after transplantation averaged 0.54 ± 0.26 g (average ± SD, n = 8) in WT and 0.54 ± 0.21 g (n = 6) in fibromodulin-deficient mice. DNA content averaged 5.8 mg DNA/g wet weight ± 0.6 (± SD, n = 5) in WT and 5.8 ± 0.3 (n = 7) in fibromodulin-deficient mice, indicating that fibromodulin had no influence on tumor cellularity.
Stroma collagen fibrils in WT and heterozygous mice were similar with an average diameter of 43 nm, whereas those in KAT-4 carcinomas grown in Fmod-null mice were significantly thinner (30 nm) (Fig. 2 A, B, and E). The gross structures of the collagen scaffold also differed, with thicker and more abundant collagen fiber bundles in WT stroma (Fig. 2 G and H). Analyses of entire whole-mount KAT-4 carcinomas showed that the densities of the collagen scaffolds were 25% (P < 0.008) higher in WT (60 ± 4 pixels per area unit, n = 4) compared with Fmod-null carcinomas (45 ± 0.5 pixels per area unit, n = 4). A similar dependence on fibromodulin for collagen scaffold structure was also observed in PROb rat colorectal adenocarcinoma (data not shown). Average hydroxyproline contents in pepsin-resistant triple-helical collagen solubilized from KAT-4 carcinomas were 2.79 ± 0.72 and 1.73 ± 0.13 μg/mg dry weight tumor tissue (± SD, n = 3, P = 0.066) in carcinomas from WT and Fmod-null animals, respectively. This 38% lowering of hydroxyproline content in fibromodulin-deficient carcinomas is in agreement with the difference in the gross structure of the collagen scaffold and the analyses of collagen densities of whole-mount tumors (Fig. 2 G and H). Fc:TβRII treatment, which reduces fibromodulin, resulted in similar collagen fibril diameters and scaffold structures as in the stroma of KAT-4 carcinomas grown in fibromodulin-deficient mice (Fig. 2 C, D, and F). Two discrete populations arose after treatment with Fc:TβRII, characterized by thin fibrils similar to those in Fmod-null stroma and wider fibrils, the latter presumably formed before treatment. The collagen scaffold gross structure after Fc:TβRII treatment and in fibromodulin-deficient stroma were similar (Fig. 2 I and J). Fc:TβRII treatment reduced collagen density by 21% (P < 0.03). Collagen densities in control carcinomas (IgG2A-treated) were 62 ± 4 pixels per unit area (n = 3) compared with 49 ± 3 pixels per unit area (n = 3) in Fc:TβRII-treated carcinomas. These results are consistent with our observation of a 35% reduction in hydroxyproline after Fc:TβRII treatment of KAT-4 carcinomas (16) and suggest a role for fibromodulin in the assembly of the collagen scaffold.
Tumor stroma ultrastructure. (A–D) Transmission electron micrographs of KAT-4 carcinomas grown in WT (A), fibromodulin-deficient (B), control IgG2A-treated (C), and Fc:TβRII-treated (D) mice. (Magnification, A–D, ×25,000; scale bar, 200 nm.) (E) Morphometric analyses of collagen fibrils in carcinomas grown in WT (open bars) and fibromodulin-deficient (filled bars) mice. (F) Collagen fibril diameters in control IgG2A-treated (open bars) and Fc:TβRII-treated (filled bars) carcinomas. (G and H) Scanning electron micrographs of carcinomas grown in WT (G) and Fmod-null (H) mice. (I and J) Control IgG2A-treated (I) and Fc:TβRII-treated (J) mice (Magnification, G–J ×300; scale bar, 100 μm). Quantification by image analyses of entire whole-mount tumors revealed that the collagen scaffold density was significantly higher (P < 0.008) in WT (60 ± 4 pixels per area unit, n = 4) (G) compared with Fmod-null carcinomas (45 ± 0.5 pixels per area unit, n = 4) (H). Collagen densities in control carcinomas (IgG2A-treated) were 62 ± 4 pixels per unit area (n = 3) (I) compared with 49 ± 3 pixels per unit area (n = 3) in Fc:TβRII-treated carcinomas (J) (P < 0.03).
IFP was lower in KAT-4 and PROb experimental carcinomas grown in Fmod-null mice (Fig. 3). Average IFP values were 5.7 ± 2.0 mmHg (1mmHg = 133 Pa) (± SD, n = 10) in WT, 4.7 ± 1.2 mmHg (n = 9) in heterozygous, and 3.7 ± 1.3 mmHg (n = 15, P < 0.05) in fibromodulin-deficient KAT-4 carcinomas. PROb carcinomas showed IFP values of 5.6 ± 1.7 mmHg (n = 4) when grown in WT mice, 4.3 ± 1.6 mmHg in Fmod +/− (n = 4), and 2.6 ± 1.8 mmHg in Fmod −/− (n = 5). The differences between WT and fibromodulin-deficient mice were significant (P < 0.05) assuming a Gaussian distribution of the values. IFP averages in tumors grown in heterozygous mice were intermediate but not significantly different from IFP averages recorded in any of the other genotypes, indicating a dose-dependent effect of fibromodulin.
Effect of fibromodulin deficiency on IFP in carcinomas. KAT-4 (A) or PROb (B) carcinomas were grown in WT, heterozygous (+/−), and Fmod-null (−/−) mice. IFPs were determined by the “wick-in-the-needle” technique described in ref. 16. Average IFP values in carcinomas grown in fibromodulin-deficient animals were significantly different from carcinomas grown in WT animals for both KAT-4 (A) and PROb (B) (P < 0.05).
Fibromodulin-deficiency had no apparent effect on blood vessels in KAT-4 carcinomas. Thus, the densities of CD31-expressing endothelial structures (Fig. 4 A), leakage of Evans' blue dye (EBD)-labeled albumin, and number of perfused vessels per area unit were not significantly different in KAT-4 carcinomas grown in WT or Fmod-null mice (Table 1). Furthermore, plasma volumes in KAT-4 carcinomas, determined from the distribution volume of radio-labeled albumin, were unaffected (Table 2). KAT-4 carcinomas grown in Fmod-null mice had, however, significantly increased extracellular fluid volumes (ECV) (Table 2). The ECV in the skin of the two genotypes were, however, similar (Table 2). Lymph vessels and cells expressing LYVE-1 were not detected in tumor tissue but were abundant in the s.c. tissues surrounding tumors. No apparent differences in distribution or intensity of LYVE-1-stained structures were detected around KAT-4 carcinomas grown in WT or Fmod-null mice (SI Fig. 6). Similar results were obtained when lymphatic vessels were detected by anti-podoplanin antibodies (data not shown).
Analyses of vessel density, macrophage infiltration, and regulation of fibromodulin mRNA by anti-inflammatory drugs in KAT-4 carcinomas. (A–C) KAT-4 carcinomas grown in WT (open bars), fibromodulin-heterozygous (stripped bars), and fibromodulin-deficient (filled bars) mice were subjected to immunohistochemistry to detect CD31-positive vessels (A), tumor-infiltrating macrophages (B), and MHC class II-expression by macrophages (C). Athymic mice carrying KAT-4 carcinomas were treated with 30 mg/kg dexamethasone (n = 3) or rh-IL-1Ra (Kinaret; injected s.c. twice daily for 10 days with 7.5 mg; n = 2) (17). (D) Fibromodulin mRNA levels were determined by real-time PCR, using β-actin or GAPDH as an internal standards. mRNA levels are presented as percent of the levels in IgG2A-treated control tumors (mean ± range).
Plasma protein leakage and dextran-perfused vessel density in KAT-4 tumors grown in fibromodulin-deficient mice
Tumor extracellular fluid and plasma volumes
Fibromodulin deficiency had no effect on the number of macrophages or their expression of MHC class II in KAT-4 tumor viable tissue (Fig. 4 B and C). Anti-inflammatory agents, such as dexamethasone and recombinant human IL-1 receptor antagonist (rh-IL-1Ra) efficiently down-regulated the expression of fibromodulin mRNA in KAT-4 carcinomas (Fig. 4 D). Immunoreactive fibromodulin in KAT-4 carcinomas was also reduced in rh-IL-1Ra-treated carcinomas (SI Fig. 5D).
Discussion
Here, we show that fibromodulin promotes the formation of a dense collagen scaffold in experimental carcinoma. Absence or reduction of fibromodulin resulted in a lowering of IFP. The lower IFP recorded in Fmod-null mice could not be ascribed to enhanced lymphatic drainage because LYVE-1-positive structures were absent in carcinoma stroma independent of fibromodulin. Furthermore, we did not detect any vascular differences between WT and Fmod-null KAT-4 carcinoma. Taken together, these observations strongly suggest that factors beyond the lymphatics and the vasculature are responsible for the reduction in IFP and increase in ECV in fibromodulin-deficient KAT-4 carcinoma. Our results thus indicate that the changes in fluid balance are caused by alterations in stroma collagen architecture.
It is likely that the architecture and functional properties of the collagen scaffold correlate with malignancy (8), although desmoplasia as a prognostic marker remains controversial (30, 31). In carcinomas resembling wounds, collagen turnover is generally high (32). Fibromodulin and other small leucine rich repeat proteoglycans bind and protect collagen fibrils from degradation by collagenases in vitro (33). Recent studies in our laboratories suggest that collagen turnover in tendons is increased in Fmod-null mice indicating a protective role for fibromodulin against degradation also in vivo (A.O., S.K., and K.R. unpublished data). Alternatively, fibromodulin may affect the collagen fibril assembly by binding and aligning monomeric triple-helical collagen and thin fibrils in the growing fibril. A proper alignment of collagen is essential for interactions and the formation of cross-links in the growing fibril. An increased turnover may in part be due to misaligned collagen fibrils that are rapidly degraded in fibromodulin-deficient tendons. The present findings of fibromodulin-directed regulation of collagen assembly in carcinoma suggests that fibromodulin synthesis constitutes a mechanism providing a stable collagen scaffold that otherwise would be rapidly degraded.
The tumor stroma is generated by carcinoma cell-driven activation of normal, i.e., nontransformed host cells, such as endothelial cells, pericytes, and fibroblasts. Carcinoma cells may activate such stroma target cells directly and/or indirectly via infiltrated immune cells. In recent years much focus has been directed to the role of macrophages for the development of a tumor stroma. Activated macrophages in carcinoma modulate their microenvironment (6) and are conversely modulated by a hypoxic microenvironment (34). Our data show that fibromodulin does not affect macrophage numbers or their expression of MHC class II antigens, the latter reportedly reflecting macrophage activation (35). Treatment of KAT-4 carcinomas with Fc:TβRII reduces inflammatory characteristics (17) and down-regulated fibromodulin expression (present findings). In agreement, the anti-inflammatory agents dexamethasone and IL-1 receptor antagonist down-regulated fibromodulin. The same agents and Fc:TβRII reduce IFP in KAT-4 carcinomas (ref. 16 and unpublished data). Dexamethasone reduces IFP also in other types of experimental carcinomas (36). Thus, our findings suggest that inflammatory processes in carcinomas promote fibromodulin synthesis by stroma cells, leading to the formation of a dense and stiff collagen scaffold and a high IFP.
The composition of the ECM determines hydraulic conductivity in tissues (37), and irradiation of experimental tumor increases collagen content and reduces hydraulic conductivity (38). The sparser collagen scaffold and reduced fibril diameters in fibromodulin-deficient carcinoma would by itself raise the hydraulic conductivity. The increased ECV will also contribute to increased conductivity in the stroma by effectively diluting matrix components (37). The increased hydraulic conductivity will favor a flow of fluid from parts of the tumor with a high IFP to the periphery where IFP is lower. Such a redistribution of fluid should result in a general reduction of IFP in the carcinoma as seen in fibromodulin-deficient KAT-4 carcinomas. The increased ECV in fibromodulin-deficient carcinomas also suggests that the collagen scaffold is less stiff, allowing tumors to swell because of the high plasma protein content in the carcinoma interstitium that results from the leaky blood vessels. Indeed, treatment of KAT-4 carcinomas with a specific inhibitor of carcinoma cell-derived vascular endothelial growth factor (Bevacizumab) decreases plasma protein leakage and reduces the ECV (19). An increased hydraulic conductivity and fluid flow in carcinomas may also explain the increased efficacy of chemotherapy in KAT-4 carcinomas pretreated with Fc:TβRII (17). Several agents reduce IFP, and some increase the delivery and efficacy of chemotherapeutics (11). Here, we show that one on these agents, Fc:TβRII, reduces fibromodulin expression. Furthermore, the collagen scaffold restricts diffusive transport of macromolecules in the stroma of experimental carcinoma (10, 39) and thus most likely the delivery of high-molecular weight anti-cancer drugs. The unexpected role for fibromodulin in controlling carcinoma fluid balance may provide a basis for a treatment to improve delivery of anti-cancer drugs by modulation of fibromodulin expression.
Methods
Animals and Tumors.
Fibromodulin-deficient mice were generated and maintained in 129 SV background (23). Fmod-null mice were crossed with C57BL nu/nu mice to generate male (nude/nude, Fmod +/−) and female (nude/wild type, Fmod +/−). All experiments were conducted on nu/nu littermates generated by breeding male (nude/nude, Fmod +/−) and female (nude/wild type, Fmod +/−). Mice were genotyped as described in ref. 26. KAT-4 anaplastic thyroid carcinoma cells (5 × 106) or rat PROb (5 × 106) colorectal cells were injected s.c. in the left flank of the mice and treated with Fc:TβRII (10 mg/kg) as described in refs. 14 and 16. All animal experiments were approved by the Ethical Committee for Animal Experiments in Uppsala and Lund, Sweden. The number of animals was minimized to comply with guidelines from the Ethical Committee.
In Vivo Perfusion, Vascular Permeability, Extracellular Fluid, and Plasma Volumes.
Perfused tumor vessels were visualized by using FITC-dextran (molecular mass 2,000 kDa) (Sigma, St. Louis, MO). Vascular permeability was assessed by determining leakage of EBD into the tumor tissue interstitium. EBD (30 mg/kg) (Sigma) was administered i.v. 30 min and FITC-dextran (100 mg/kg) 2 min before animal killing. To evaluate the perfused area of tumor vasculature and leakage of EBD, 20-μm frozen sections were analyzed by fluorescent microscopy. Images were obtained from 16 fields of vision from eight sections per tumor taken 100 μm apart and assessed under low magnification. The extent of EBD leakage was analyzed by using NIH Image 1.62 software (National Institutes of Health, Bethesda, MD). Digital images were analyzed in a gray-scale mode, and dye density in tumor sections was presented as number of pixels per area of tumor tissue. The density of FITC-dextran-perfused vessels was quantified in 16 fields of vision with high vascular density under low magnification according to Chalkley point-overlap morphometry. Using this method, the mean of perfused vessel densities of the most vascular areas within the tumor (×200 magnification) is calculated manually. Selected areas (“hot spots”) were identified after scanning the whole section at low magnification. This technique has been established as a standard method for quantification of angiogenesis in sections from solid tumors. ECV in KAT-4 carcinomas were measured as extravascular distribution space of 51Cr-EDTA after functional nephrectomy by the bilateral ligation of the renal pedicles via flank incisions. Plasma volumes were measured by using 125I-labeled human serum albumin (125I-HSA; Institute for Energy Techniques, Kjeller, Norway). Both techniques have been detailed elsewhere (19).
Immunohistochemistry and Immunofluorescence.
Six-micrometer frozen sections of KAT-4 carcinomas were subjected to immunohistochemistry with anti-mouse CD31/PECAM-1 antibody (BD PharMingen, San Diego, CA) detecting endothelial cells and anti-mouse F4/80 antibody (Serotec, Oxford, U.K.) detecting macrophages. Blood vessel density and macrophage infiltration of tumor tissue were analyzed under ×200 and ×500 magnification by using a counting grid as described in ref. 19. Pericyte coverage of CD31-positive blood vessels was determined by double immunofluorescence with anti-smooth muscle actin antibody (Sigma). MHC class II antigen expression by F4/80-positive intratumoral macrophages was analyzed with anti-mouse FITC-labeled I-A/A-E (2G9) antibody (BD Phar-Mingen). Lymphatic vessels were detected by double immunofluorescence with rabbit anti-LYVE-1 IgG (Abcam, Cambridge, U.K.) and anti-CD31.
Quantitation of Hydroxyproline and Quantitative Real Time PCR.
Lyophilized tumors were treated with 2 mg/ml porcine stomach mucosa pepsin (Sigma) in 0.5 M acetic acid. Released triple helical collagen was precipitated with 1 M NaCl. The collagen precipitate was hydrolyzed by 6 M HCl at 110°C for 18 h. Hydroxyproline content was determined in the hydrolysate as described in ref. 40. Total RNA was extracted from KAT-4 tumors by using RNAeasy kit (Qiagen, Valencia, CA) and TRIZOL solution (Invitrogen, San Diego, CA) and with DNase I, and a first-strand cDNA synthesis was performed [first-strand synthesis kits (Roche Diagnostics, Penzberg, Germany) or iScript (Bio-Rad Laboratories, Sundbyberg, Sweden)], using oligo(dT) primers. The cDNA samples were mixed with specific primers and SYBR Green and amplified in an Applied Biosystems Prism instrument (Applied Biosystems, Foster City, CA), starting with an initial 10-min heating at 95°C followed by 40 cycles at 95°C for 15 s, 60°C for 30 s, and 72°C for 30 s. The data were analyzed by using SDS software, Version 2.1 (Applied Biosystems). The calculated threshold cycle values were normalized to the threshold cycle-value for β-actin or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) used as internal standard. The primers used were: Fmod, 5′-AGCAGTCCACCTACTACGACC-3′ and 5′-CAGTCGCATTCTTGGGGACA-3′; β-actin, 5′-GGCACCCAGCACAATGAAG-3′ and 5′-GCCGATCCACACGGAGTACT-3′; GAPDH, 5′-AGG T CGGTGTGAACGGATTTG-3′ and 5′-TGTAGACCATGTAGTTGAGGTCA-3′.
Electron Microscopy.
Three similarly sized tumors from WT, fibromodulin-deficient, IgG2A-treated, and Fc:TβRII-treated mice were investigated by transmission electron microscopy. Carcinomas were fixed in 0.15 M sodium cacodylate-buffered 2% glutaraldehyde, postfixed in 0.1 M-collidine-buffered 2% osmium tetroxide, and embedded in epoxy resin (23). Ultrathin sections were analyzed in Philips CM-10 electron microscope (Philips, Amsterdam, The Netherlands). Scanning electron microscopy was performed on whole-mount specimens prepared by alkali maceration (41) and analyzed in a Philips 515 electron microscope. For quantification of matrix amount, the images were converted to white and black pixels that represented matrix and empty areas, respectively. Pixels were quantified with ImageJ software in three whole-mount carcinomas of similar size from control and Fc:TβRII-treated carcinomas and from carcinomas grown in WT and fibromodulin-deficient mice. Collagen densities are expressed as percent of area occupied by matrix.
Statistical Methods.
The unpaired, two-tailed Student's t test was used. The Mann–Whitney U test was used when data failed to fulfill the normality criteria. P < 0.05 was considered statistically significant. Standard deviations of means are indicated in the figures unless otherwise specified.
Acknowledgments
We thank Ann-Marie Gustafson, Annika Hermansson, and Gerd Salvesen for technical assistance. This work was supported by grants from the Swedish Cancer Foundation (N.-E.H. and K.R.), Swedish Research Council (Å.O., N.-E.H., and K.R.), Gustaf V:s 80-Anniversary Fund (Å.O. and K.R.), Greta and Johan Kocks and Alfred Österlunds Funds (Å.O.), Gunnar Nilssons Cancer Organization (K.R.), and the Norwegian Research Council (L.S. and R.K.R.).
Footnotes
- ††To whom correspondence should be addressed. E-mail: kristofer.rubin{at}imbim.uu.se
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Author contributions: Å.O., S.K., and K.R. designed research; Å.O., S.K., A.V.S., L.S., R.K.R., N.-E.H., and K.R. performed research; Å.O., S.K., A.V.S., L.S., M.M., R.K.R., N.-E.H., and K.R. analyzed data; Å.O., R.K.R., and K.R. wrote the paper.
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↵ §Present address: Division of Molecular Immunology, German Cancer Research Center, D-69120 Heidelberg, Germany.
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The authors declare no conflict of interest.
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This article is a PNAS Direct Submission.
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This article contains supporting information online at www.pnas.org/cgi/content/full/0702014104/DC1.
- Abbreviations:
- ECM,
- extracellular matrix;
- EBD,
- Evans' blue dye;
- IFP,
- interstitial fluid pressure;
- ECV,
- extracellular fluid volumes.
- © 2007 by The National Academy of Sciences of the USA