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Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

There is increasing evidence that the physical environment is a critical mediator of tumor behavior. Hepatocellular carcinoma (HCC) develops within an altered biomechanical environment, and increasing matrix stiffness is a strong predictor of HCC development. The aim of this study was to establish whether changes in matrix stiffness, which are characteristic of inflammation and fibrosis, regulate HCC cell proliferation and chemotherapeutic response. Using an in vitro system of “mechanically tunable” matrix-coated polyacrylamide gels, matrix stiffness was modeled across a pathophysiologically relevant range, corresponding to values encountered in normal and fibrotic livers. Increasing matrix stiffness was found to promote HCC cell proliferation. The proliferative index (assessed by Ki67 staining) of Huh7 and HepG2 cells was 2.7-fold and 12.2-fold higher, respectively, when the cells were cultured on stiff (12 kPa) versus soft (1 kPa) supports. This was associated with stiffness-dependent regulation of basal and hepatocyte growth factor–stimulated mitogenic signaling through extracellular signal-regulated kinase, protein kinase B (PKB/Akt), and signal transducer and activator of transcription 3. β1-Integrin and focal adhesion kinase were found to modulate stiffness-dependent HCC cell proliferation. Following treatment with cisplatin, we observed reduced apoptosis in HCC cells cultured on stiff versus soft (physiological) supports. Interestingly, however, surviving cells from soft supports had significantly higher clonogenic capacity than surviving cells from a stiff microenvironment. This was associated with enhanced expression of cancer stem cell markers, including clusters of differentiation 44 (CD44), CD133, c-kit, cysteine-X-cysteine receptor 4, octamer-4 (CXCR4), and NANOG. Conclusion: Increasing matrix stiffness promotes proliferation and chemotherapeutic resistance, whereas a soft environment induces reversible cellular dormancy and stem cell characteristics in HCC. This has implications for both the treatment of primary HCC and the prevention of tumor outgrowth from disseminated tumor cells. (HEPATOLOGY 2011;)

Hepatocellular carcinoma (HCC) is the third most common cause of cancer-related mortality worldwide.1 The majority (80%) of HCCs develop in the context of advanced liver fibrosis or cirrhosis and liver cirrhosis is the single most important risk factor for HCC development.2 Liver fibrosis is defined by stereotypical changes in both the biochemical and physical properties of the cellular microenvironment. However, the role of mechanical factors in modulating the growth and progression of HCC remain poorly defined. Recent studies involving ultrasound elastography (FibroScan) demonstrate that liver stiffness measurements are a strong predictor of HCC development.3 Furthermore, once established, tumor development is associated with further increases in matrix stiffness to values greater than those of the surrounding hepatic parenchyma.4 It is therefore evident that HCC develops in a niche with mechanical properties distinct from those encountered in the normal liver.

Cancer development and progression is dependent on both intrinsic genetic abnormalities and external structural determinants.5 Matrix stiffness has recently been directly implicated in aiding tumor development. Increases in matrix stiffness that enhance cell contractility have been found to be sufficient to enhance the transformation of mammary epithelial cells.6 Conversely, a reduction in tissue stiffness by inhibition of collagen cross-linking impedes malignant growth and tumor development in a murine model of breast cancer.7 Cellular stiffness sensing relies upon intracellular tension, which is determined by a dynamic equilibrium between forces generated by a contractile cytoskeleton and the elastic resistance (stiffness) provided by the extracellular matrix (ECM). In this context, cancer progression (tumor growth, invasion, and dissemination) is accompanied by changes in both the mechanical properties of the cancer cell niche and changes in cellular contractility (modified by genetic and epigenetic factors) that regulate tumor cell behavior.

HCC continues to have a poor prognosis (median survival less than 12 months), reflecting its late presentation and lack of effective therapies.8, 9 The effectiveness of both hepatic resection and liver transplantation for HCC is limited by tumor recurrence, which can occur months or years following resection of a primary tumor.10 Furthermore, systemic chemotherapy has proved ineffective both for the treatment of advanced HCC and in an adjuvant/neoadjuvant setting for the eradication of disseminated (dormant) tumor cells, the progenitors of clinical metastases. The mechanisms underlying chemotherapy resistance in HCC have not been fully elucidated. Although it has previously been demonstrated that the composition of the matrix can enhance chemotherapy resistance in a range of epithelial cancers,11, 12 the role of matrix stiffness has not been specifically addressed for HCC or other epithelial cancers.

Here, we demonstrate that mechanical factors regulate both the proliferation and chemotherapeutic response of HCC cells. In addition, we show that both tumor cell differentiation and cancer stem cell characteristics are influenced by the mechanical properties (that is, stiffness) of the cancer cell niche.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Cell Culture and Microscopy.

Human HCC cell lines Huh7 and HepG2 (kindly provided by S. Wigmore, Edinburgh, UK) were cultured in Dulbecco's modified Eagle medium (Gibco, Paisley, UK) supplemented with 10% fetal bovine serum, penicillin/streptomycin and L-glutamine. For all experiments, cells were plated at semiconfluent density in 1% fetal bovine serum. Chemical reagents were purchased from Sigma (Poole, UK) unless otherwise stated. Transforming growth factor-beta (TGFβ) and hepatocyte growth factor (HGF) (Peprotech, London, UK) were used at concentrations of 5 ng/mL and 10 ng/mL, respectively. Anti–β1-integrin, clone 6S6 (Millipore, Watford, UK) and control immunoglobulin G1 (IgG1; AbD Serotec, Oxford, UK) were used for cell culture experiments at 50 μg/mL. Echistatin (Tocris, Bristol, UK) was used in cell culture experiments at 100 nM concentration. The chemical focal adhesion kinase (FAK) inhibitor, PF573228 (Tocris, Bristol, UK) was solubilized in dimethylsulfoxide (DMSO) and used for cell culture experiments at a concentration of 1-5 μM. A detailed description of microscopy and morphological analysis can be found in the Supporting Methods online.

Preparation of Polyacrylamide Gel Supports.

Polyacrylamide (PA) gels of variable stiffness were prepared on glass coverslips using modifications13 to the method initially described by Pelham and Wang.14 A detailed description can be found in the Supporting Methods online.

Immunofluorescence Staining.

Cells were fixed in 4% paraformaldehyde in phosphate-buffered saline and permeabilized with 0.2% Triton-X-100 in phosphate-buffered saline. Slides were stained with anti-Ki67 (Novocastra, Newcastle, UK) and anti-vinculin (Sigma, Poole, UK); corresponding Alexa Fluor-555 secondary antibodies were used for detection (Invitrogen, Paisley, UK). Actin stress fibers were stained with Alexa-488 phalloidin (Invitrogen). Nuclear DNA was counterstained with 4′,6′-diamidino-2-phenyl-indole dihydrochloride (DAPI; Dako, Ely, UK). Cellular proliferative index (Ki67-positive cells/total cells) was calculated by direct cell counting from 15 randomly selected high magnification photomicrographs from Ki67-stained slides (n = 3).

Western Blot.

A detailed description of western blotting and a complete list of antibodies are provided in the Supporting Methods.

Immunohistochemistry.

Human HCC specimens were obtained from archived tissue held by Tayside Tissue Bank and the Department of Pathology, University Medical Center Hamburg-Eppendorf with appropriate ethical approval (UK-LREC: TR000216). Immunohistochemistry was performed as previously described.15 The primary antibodies and antigen retrieval regimes used were anti-pFAK (pY397) (Invitrogen, Paisley, UK/Microwave pH9) and anti-β1-integrin (Abcam, Cambridge, UK/Microwave pH9). Negative controls with isotype immunoglobulins (Santa Cruz, Heidelberg, Germany) and species-specific serum alone showed no specific staining.

Clonogenic Assay.

Cells were plated at semi-confluent density onto PA gels. After 48 hours, cells either received cisplatin (HepG2, 10 μM/Huh7, 20 μM) or 5-fluorouracil (5FU; 25 μM) or were left untreated in plating medium. After 24 hours, the medium was changed to normal culture medium and the cells were incubated further for 48 hours, for a total of 5 days of culture. Cells were then retrieved by trypsinization, counted and plated at clonal density (10,000 cells/well) into 12-well plates in normal culture medium. Cells were fixed at between 5-10 days in 4% paraformaldehyde and stained with 0.5% crystal violet solution. Colonies were visualized with a VersaDoc system and analyzed with Quantify-One (Bio-Rad Laboratories, Hercules, CA).

Flow-Cytometric Analysis.

Cells were harvested by trypsinization and single cell suspension generated by passing cells through a 40 μm cell strainer. Cells were stained with the following antibodies: CD44-phycoerythrin (PE), CD117-PE (c-kit), CD133-PE, CD184-PE (cysteine-X-cysteine receptor 4 [CXCR-4]), and corresponding PE-labeled isotype controls (E-Bioscience, Hatfield, UK). After staining, cells were washed, post-fixed in 1% paraformaldehyde and analyzed on a FACScan (BD Biosciences, Franklin Lakes, NJ). Data analysis was performed using FlowJo software (TreeStar, Inc., Ashland, OR).

cDNA Synthesis and Real-Time PCR.

Relative mRNA expression for genes of interest was determined by real-time PCR using an Applied Biosystems 7700 Sequence Detection System. Primer sequences for the genes of interest and the 18S housekeeping gene were purchased from Applied Biosystems (Warrington, UK).

Statistics.

Data are expressed as mean ± standard error of the mean (SEM) of at least three independent experiments unless stated otherwise. Comparisons between groups were performed using a two-tailed Student t test.

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Increasing Matrix Stiffness Is Associated With a Mesenchymal Shift in HCC Cells.

The response of HCC cells to alterations in matrix stiffness was investigated using a system of mechanically-tunable ligand-coated PA gels.13, 14 In this system, matrix stiffness is altered by modulating the bis-acrylamide crosslink density of thin PA gels without altering the surface composition or density of ligands to which the cells are exposed.13 Matrix stiffness (expressed as shear modulus, G′) was modeled across a range of pathophysiologically-relevant stiffness values (1-12 kPa) corresponding to values encountered in normal and fibrotic livers.16 The PA gels used in this study were coated with collagen-I, representing the predominant ECM protein encountered in the fibrotic liver.

For both Huh7 and HepG2 cells we observed a consistent morphological response to changes in support stiffness. HCC cells on soft (1 kPa) supports were small and rounded in contrast to the well-spread and flattened cells seen on stiff (12 kPa) supports (Fig. 1). Differences in cellular spreading as a function of support stiffness develop rapidly (within 1 hour) and are maximal at 24 hours (Supporting Fig. 1). Confocal microscopy showed that increasing matrix stiffness was associated with the development of prominent actin stress fibers and mature (vinculin-positive) focal adhesions (Fig. 2). These features were absent in cells cultured on soft supports. The presence of stress fibers is linked to acquisition of mesenchymal properties (mesenchymal-shift) and de-differentiation in epithelial cells. In accordance with this we demonstrated up-regulation of the mesenchymal markers N-cadherin (Huh7/HepG2) and vimentin (shown for Huh7; vimentin is not expressed in HepG2 cells under either condition) in HCC cells cultured on stiff supports (Fig. 3A). There was no change in the expression of the epithelial marker E-cadherin. HepG2 and Huh7 cells cultured on soft supports expressed higher levels of albumin, hepatocyte nuclear factor-4α (HNF4α), alpha-1-antitrypsin and alpha-fetoprotein (AFP) than cells cultured on stiff supports (Fig. 3B). This suggests that a soft environment promotes a differentiated hepatocyte phenotype, whereas increasing support stiffness is associated with cellular de-differentiation toward a mesenchymal phenotype.

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Figure 1. Changes in matrix stiffness regulate HCC cell morphology and spreading. Huh7 and HepG2 cells were cultured on collagen-I–coated polyacrylamide (PA) gels with “tunable stiffness” (expressed as shear modulus, G′) in the range of 1-12 kPa and collagen-I–coated glass. The stiffness values of the PA gel supports used were selected in order to reflect range of stiffness values encountered in normal and fibrotic livers. Phase-contrast photomicrographs demonstrate the regulation of cellular morphology by support stiffness in both (A) Huh7 and (B) HepG2 cells. The surface area (square micrometers) of (C) Huh7 and (D) HepG2 cells was calculated by digital image analysis of phase-contrast images of cells on PA gel supports. In each case, values reflect the mean (±SEM) of measurements from 50 cells in three independent experiments (*P < 0.05, **P < 0.01, and ***P < 0.001).

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Figure 2. Changes in matrix stiffness regulate the formation of actin stress fibers and focal adhesion maturation in HCC cells. Confocal microscopy (×320 magnification) of (A) Huh7 and (B) HepG2 cultured on soft (1 kPa) and stiff (12 kPa) collagen-I–coated PA supports as indicated. The photomicrographs displayed are of cells stained for the presence of actin stress fibers (phalloidin: green), mature focal adhesions (anti-vinculin: red), and nuclear DNA (DAPI: blue). The merged image (right panel) demonstrates the spatial relationship between actin stress fibers and mature focal adhesions. Insets display high-magnification images for Huh7 and HepG2 cells cultured on stiff (12 kPa) supports demonstrating the insertion of actin stress fibers into mature focal adhesions. In each image, the scale bars represent 20 μm.

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Figure 3. Increased matrix stiffness is associated with mesenchymal shift in HCC cells. (A) Western blot from whole-cell lysates showing expression of E-cadherin, N-cadherin, and vimentin in Huh7 and HepG2 cells cultured on soft (1 kPa) and stiff (12 kPa) collagen-I–coated PA gel supports (as indicated). (B) Western blots from whole-cell lysates showing expression of albumin, hepatocyte nuclear factor 4 alpha (HNF4α), alpha-1-antitrypsin, and alpha-fetoprotein (AFP) in Huh7 and HepG2 cells. (C) Western blots from whole-cell lysates from Huh7 cells showing the expression of phospho-Smad2, phospho-Smad3, and total Smad2/3 following stimulation with transforming growth factor beta (TGFβ; 5 ng/mL). In each western blot, equal quantities of protein were loaded and equal loading was confirmed in relation to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression. In each case, the western blots shown are representative examples from three independent experiments. (D) The line graphs show a schematic representation of densitometry analysis of phosphorylated Smad2 and Smad3, expressed relative to GAPDH. Each time point represents the mean of three independent experiments.

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TGFβ is a potent inducer of mesenchymal changes in both primary and transformed epithelial cells. We therefore investigated whether support stiffness regulated TGFβ-induced Smad signaling activity in HCC cells. The Huh7 cell line demonstrated increased basal activity of the TGFβ signaling pathway (as indicated by increased Smad3 phosphorylation) in cells cultured on stiff supports (Fig. 3C,D). In addition, upon stimulation with TGFβ there was enhanced Smad2 and Smad3 phosphorylation in cells from stiff supports.

Increasing Matrix Stiffness Promotes HCC Proliferation.

In both HCC cell lines, matrix stiffness regulated HCC cell proliferation (Fig. 4A). The proliferative indices of Huh7 and HepG2 cells (assessed by nuclear localization of Ki67) were 2.7-fold (P < 0.001) and 12.2-fold (P < 0.001) higher, respectively, when the cells were cultured on stiff (12 kPa) versus soft (1 kPa) supports. Maximal proliferative index was seen when cells were cultured on collagen-I–coated glass, which has a shear modulus several orders of magnitude higher than any physiological matrix. Both MTT assay (Supporting Fig. 2) and direct cell counting (data not shown) confirmed an increase in total cell number with increasing support stiffness. A similar trend for cellular proliferation was observed in primary mouse hepatocytes (Supporting Fig. 3). Matrix stiffness had a corresponding effect on the expression of cell cycle regulators of G1 progression (Fig. 4B,C). We observed a strong reduction in the expression of cyclin-D1 and cyclin-D3 in cells cultured on soft supports. There was no evidence of up-regulation of the cyclin-dependent kinase inhibitors p21cip or p27kip on soft gels and indeed a moderate down-regulation of p27kip was observed on soft gels. Induction of terminal senescence on soft supports was excluded by showing that upon transfer to a stiff matrix, cells resumed proliferation to levels comparable to cells coming from a stiff matrix (Supporting Fig. 4A). Furthermore, cells on both soft and stiff supports showed no evidence of beta-galactosidase accumulation (data not shown). In each cell line, differences in cellular proliferation as a function of stiffness were evident across a wide range of plating densities (Supporting Fig. 5).

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Figure 4. Increased matrix stiffness promotes HCC cell proliferation. (A) Graphs showing the mean proliferative index (Ki67 positivity) of Huh7 and HepG2 cells cultured on collagen-I–coated polyacrylamide gel supports across a range of stiffness values (1-12 kPa), as indicated, and collagen-I–coated glass (n = 3). (B) Western blots from whole-cell lysates showing expression of cyclin-D1 and cyclin-D3 in Huh7 and HepG2 cells cultured on soft (1 kPa) and stiff (12 kPa) supports, as indicated. (C) Western blots from whole-cell lysates showing expression of cyclin-dependent kinase inhibitors, p21cip, and p27kip in Huh7 and HepG2, cultured on soft (1 kPa) and stiff (12 kPa) supports, as indicated. In each western blot, equal quantities of protein were loaded and equal loading confirmed in relation to GAPDH expression. The western blots shown are representative examples from three to six independent experiments. (D). Graphs showing the mean proliferative index (Ki67 positivity) of Huh7 (left panel) and HepG2 (right panel) cells cultured on both soft (1 kPa) and stiff (12 kPa) polyacrylamide supports coated with collagen-I, collagen-IV, laminin, or fibronectin (n = 3). In each case, error bars represent SEM, *P < 0.05, **P < 0.01, and ***P < 0.001.

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Figure 5. Matrix stiffness regulates mitogenic signaling in HCC. (A) Western blots showing basal expression of phosphorylated and total FAK, ERK, protein kinase B (PKB/Akt), and STAT3 in Huh7 and HepG2 cells cultured on soft (1 kPa) and stiff (12 kPa) supports, as indicated. (B) Western blots showing cyclin-D1 expression in Huh7 and HepG2 cells cultured for 24 hours on soft (1 kPa) and stiff (12 kPa) supports in the presence (+) or absence (−) of HGF (10 ng/mL). (C) Western blots showing a time-course analysis for expression of phosphorylated and total ERK, PKB/Akt, and STAT3 in Huh7 cells cultured on soft (1 kPa) and stiff (12 kPa) supports. Whole-cell lysates were harvested at baseline and specific time points (as indicated) following the addition of HGF (10 ng/mL) to culture media. In each western blot, equal quantities of protein were loaded and equal loading was confirmed in relation to GAPDH expression. The line graphs (right panel) show a schematic representation of densitometry analysis of phosphorylated ERK, PKB/Akt, and STAT3 expressed relative to GAPDH. Each time point represents the mean of three independent experiments.

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In order to exclude a specific effect related to collagen-I, we investigated the effect of different ECM coatings on HCC cell proliferation on PA gels (Fig. 4D). Although minor differences were observed with respect to cellular morphology and spreading (Supporting Fig. 6) when cells were plated on collagen-I, collagen-IV, laminin and fibronectin-coated gels, the biochemical composition of the surface coating did not significantly alter the stiffness-dependent regulation of cell proliferation. In other words, the physical rather than the biochemical properties of the PA gels exerted the predominant effect on HCC cell proliferation.

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Figure 6. β1-Integrin and phospho-FAK are expressed in human HCC tumors and regulate the stiffness-dependent proliferation of human HCC cells in vitro. (A) Low magnification (×50) photomicrographs from a human HCC resection specimen stained with hematoxylin and eosin (left panel), anti–β1-integrin (middle panel) and anti-phospho-FAK (right panel). Negative control staining is represented by the indented images in the top righthand corner of each image. β1-integrin is expressed in both the HCC tumor (HCC) and surrounding hepatic parenchyma (P), as indicated. Phospho-FAKTyr397 is strongly expressed in the HCC tissue relative to the hepatic parenchyma. Scale bars represent 200 micrometers. (B) Western blots showing the expression of phospho-FAKTyr397 in Huh7 and HepG2 cells either left untreated or treated for 24 hours with the focal adhesion kinase (FAK) inhibitor PF573228 at concentrations of 1 μM and 5 μM, as indicated. In each western blot, equal quantities of protein were loaded and equal loading confirmed in relation to GAPDH expression. (C) Graphs showing the mean proliferative index (Ki67 positivity) of Huh7 and HepG2 cells cultured on 12 kPa (stiff) collagen-I-coated polyacrylamide supports. Cells were treated with the anti-β1-integrin antibody 6S6 (50 μg/mL), isotype control IgG1 antibody (50 μg/mL), echistatin (100 nm), PF573228 (1 μM), PF573228 (5μM), DMSO (vehicle control), or left in media alone (untreated control) for 24 hours, as indicated (n = 3-5). In each case, error bars represent SEM, *P < 0.05, **P < 0.01, and ***P < 0.001.

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Matrix Stiffness Modulates Basal and HGF-Induced Signaling Responses.

Using immunoblotting with phosphorylation-specific antibodies we analyzed stiffness-dependent differences in the activity of critical mitogenic signaling pathways. Growth on stiff (12 kPa) versus soft (1 kPa) supports was associated with enhanced FAK, extracellular signal-regulated kinase (ERK), protein kinase B (PKB/Akt) (Huh7 cells only), and signal transducer and activator of transcription 3 (STAT3) phosphorylation (Fig. 5A). Substrate stiffness significantly modulated growth factor-induced mitogenic signaling in response to HGF. Upon stimulating cells plated on both soft and stiff PA gels with HGF, we observed an increase in the magnitude of ERK, PKB/Akt, and STAT3 activation in cells cultured on stiff gels (Fig. 5C, Supporting Fig. 7). Substrate stiffness also modulated cyclin-D1 expression in response to HGF stimulation. Following HGF stimulation in HepG2 and Huh7 cells, there was up-regulation of cyclin-D1 expression in cells cultured on both soft and stiff supports (Fig. 5B). Importantly, the magnitude of cyclin-D1 induction following HGF stimulation was substantially higher in cells cultured on stiff supports.

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Figure 7. Matrix stiffness regulates apoptosis and clonogenic capacity following chemotherapy. (A) Western blot showing full-length (116 kDa) and cleaved poly-ADP-ribose polymerase (PARP) (89 kDa) expression in Huh7 and HepG2 cells following treatment with cisplatin on soft (1 kPa) and stiff (12 kPa) supports, as indicated. In each western blot, equal quantities of protein were loaded and equal loading was confirmed in relation to GAPDH expression. The western blots shown are representative examples from three independent experiments. (B) Colony formation potential of Huh7 and HepG2 cells following chemotherapy. Huh7 and HepG2 cells were cultured for 48 hours on either soft (1 kPa) or stiff (12 kPa) polyacrylamide supports. Cells were then left untreated (left panel) or treated with cisplatin (middle panel) or 5-fluorouracil (5-FU) (right panel) for 24 hours prior to media change. After a further 48 hours in culture, the cells were trypsinized and equal numbers re-plated at clonal density in 12-well plates. Clonogenic capacity was calculated by direct counting of the resulting colonies. The results are expressed as the percentage colony formation relative to the number of colonies obtained from 1 kPa supports from three independent experiments. In each case, error bars represent SEM, *p<0.05, ***p<0.001 and ns=not significant.

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β1-Integrin and Phospho-FAK Are Expressed in Human HCC Tumors and Regulate the Stiffness-Dependent Proliferation of Human HCC Cells In Vitro.

Integrins and integrin-associated focal adhesions are known to be important mediators of mechanotransduction. We therefore used immunohistochemistry to investigate the prevalence of β1-integrin and phospho-FAKTyr397 expression in HCC tissue from an unselected cohort of 15 HCC specimens obtained at the time of tumor resection or biopsy (Fig. 6A). β1-Integrin was expressed in tumor tissue in all 15 of 15 HCC specimens tested. In addition, we found up-regulation of FAK expression in tumor tissue relative to the surrounding parenchyma in 8/15 (53%) HCC specimens tested. These results are consistent with published histological studies.17, 18 We subsequently investigated the effect of the β1-integrin inhibition on HCC cell proliferation in vitro using a function blocking anti–β1-integrin antibody (6S6) and the disintegrin echistatin (Fig. 6C). Anti–β1-integrin antibody and echistatin promoted cellular rounding in both the Huh7 and HepG2 cells cultured on collagen-I–coated 12 kPa (stiff) supports. Huh7 cell proliferation was reduced by treatment with both 6S6 antibody (38% reduction, P < 0.05) and echistatin (29% reduction, P = 0.07) relative to relevant controls. Similarly, in HepG2 cells, cell proliferation was reduced by treatment with both 6S6 antibody (92% reduction, P < 0.001) and echistatin (21% reduction, P < 0.01). The effect of FAK activation on HCC cell proliferation was investigated in experiments with the small molecular FAK inhibitor PF573228 (Fig. 6B,C). Treatment with PF573228 (5 μM) was associated with a reduction in the proliferation of both Huh7 (42% reduction, P < 0.01) and HepG2 cells (45% reduction, P < 0.001) cultured on collagen-I–coated 12 kPa polyacrylamide gels. Furthermore, inhibition of β1-integrin or FAK expression in HepG2 cells with siRNA resulted in a significant reduction in cellular proliferation relative to control siRNA transfection (Supporting Fig. 8). A similar trend in respect to cellular proliferation was observed following siRNA-dependent inhibition of β1-integrin or FAK expression in Huh7 cells, although in this case the reduction was not statistically significant.

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Figure 8. Matrix stiffness and chemotherapy regulate stem cell marker expression in HepG2 cells. (A) Quantification by flow cytometric analysis of putative cancer stem cell markers CD133, c-kit, CD44 and CXCR-4 in HepG2 cells cultured for 5 days on soft (1 kPa) or stiff (12 kPa) supports. Cells were either left untreated (black) or treated for 24 hours with cisplatin (white). Results are representative of three independent experiments. (B) Real-time quantitative PCR analysis of octamer-4 (OCT4) (left panel) and NANOG (right panel) expression in HepG2 cells cultured for 5-days on soft (1 kPa) or stiff (12 kPa) supports. Cells were either left untreated (black) or treated for 24 hours with cisplatin (white). Expression is relative to the 18S housekeeping gene. In each case, error bars represent SEM, *P < 0.05, **P < 0.01, and ***P < 0.001.

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Matrix Stiffness Modulates HCC Apoptosis and Clone-Forming Capability Following Chemotherapy.

HCC is resistant to treatment with conventional chemotherapeutic agents. We therefore investigated whether the stiffness of the cancer cell niche regulates the susceptibility of HCC cells to chemotherapy-induced apoptosis. In both cell lines, there was decreased apoptosis in cells cultured on stiff supports, as indicated by reduced poly-ADP-ribose polymerase (PARP) cleavage (Fig. 7A). There was a nonsignificant trend toward increased numbers of surviving cells on stiff supports (data not shown). We also performed a series of clonogenic assays to investigate whether changes in matrix stiffness modulate the survival and behavior of tumor-initiating cells after chemotherapy. Following cisplatin treatment, the surviving cell population included an increased frequency of clone-initiating cells for both HepG2 (2.4-fold, P < 0.001) and Huh7 cells (2.2-fold, P < 0.05) cultured on soft (1 kPa) versus stiff (12 kPa) supports (Fig. 7B). In addition, there was a nonsignificant trend toward an absolute increase in the total number of clone-forming cells from soft supports (data not shown). To assess the validity of this finding, experiments were repeated using a second, unrelated chemotherapeutic agent, 5-fluorouracil (5-FU). Consistent with our findings with cisplatin, following 5-FU chemotherapy there was an increased frequency of clone-initiating cells from HepG2 (3.6-fold, P < 0.001) and Huh7 cells (1.9-fold, P < 0.05) cultured on soft versus stiff supports. There was no difference in the frequency of clone-initiating cells in untreated HepG2 or Huh7 cells after culture on soft or stiff supports in the absence of chemotherapy.

The paradoxical increase in the clone-initiating capability of chemotherapy-treated cells from a low stiffness environment could be explained by selective enrichment for clone-initiating cells with stem cell characteristics. We therefore performed flow-cytometric analyses for putative cancer stem cell markers in HCC cells cultured on soft (1 kPa) and stiff (12 kPa) supports, both without and following cisplatin treatment (Fig. 8A). Culture on soft versus stiff supports was associated with an enrichment for the cell surface markers CD133 (1.5-fold, P < 0.001), c-kit (1.3-fold, P = 0.78), CD44 (6.4-fold, P < 0.001), and CXCR-4 (2.9-fold, P < 0.01). Following cisplatin treatment, there was statistically significant up-regulation of CD44 (1.7-fold, P < 0.01), CD133 (1.6-fold, P < 0.01) and c-kit (15.8-fold, P < 0.01) for cells maintained on soft but not stiff supports. Additionally, real-time PCR demonstrated up-regulation of stem cell-associated transcription factors OCT4 and NANOG in HepG2 cells cultured on soft versus stiff supports, both in untreated controls (OCT4 2.0-fold increase, P < 0.05; NANOG 2.7-fold increase, P < 0.05) and following cisplatin treatment (OCT4 2.0-fold increase, P < 0.05; NANOG 3.4-fold increase, P < 0.05) (Fig. 8B).

Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

In this study, we demonstrated that the stiffness of the subcellular matrix profoundly alters the phenotype and behavior of HCC cells in vitro. Pathophysiological increases in matrix stiffness, as encountered in fibrotic and cirrhotic livers,19 promote the proliferation of HCC cells. Our work defines novel mechanisms linking the physical properties of the fibrotic liver and the malignant behavior of HCC. Our data is consistent with in vivo evidence, not only of de novo HCC development and progression against a background of cirrhosis, but also animal studies showing that the induction of liver fibrosis is associated with accelerated tumor growth following orthotopic HCC implantation.20, 21 Furthermore, histological examination of human HCC specimens demonstrates a significant association between the presence of hepatic fibrosis and enhanced tumor cell proliferation.22 Critically, our findings suggest that a reduction in the stiffness of the cancer cell niche, as would be encountered by a disseminated tumor cell entering an unaffected secondary site, would be sufficient to promote reversible cellular quiescence and cancer cell dormancy.

It has previously been demonstrated that matrix stiffness can regulate proliferation in nontransformed cells. More recently increased matrix stiffness has been shown to promote cellular proliferation in glioma cells.23 We have extended these findings to a range of epithelial malignancies, including HCC (Supporting Fig. 9). Furthermore, we have shown that β1-integrin and FAK (the canonical mediator of integrin-related signaling) regulate stiffness-dependent proliferation in HCC cells. In both fibroblasts and nontransformed mammary epithelial cells, a critical role for ERK-induced cyclin-D induction has been established in the stiffness-dependent regulation of cell proliferation.24 In accordance with these findings we demonstrate both an up-regulation of cyclin-D levels and increased mitogenic signaling through ERK in HCC cells on stiff substrates. Interestingly, reduced ERK activation has previously been linked to cellular quiescence in a cell line-specific model of cancer dormancy.25 Additionally, for the first time, we demonstrate a role for matrix stiffness in modulating the activation of the STAT3 pathway. STAT3 has recently been identified as a central component in tumor progression and a potential target of cancer therapy in HCC and other epithelial malignancies.26 The STAT3 pathway is activated in response to multiple cytokines and growth factors during cancer-associated inflammation (e.g., interleukin-6, interleukin-10, epidermal growth factor, and HGF). Our findings demonstrate that matrix stiffness has a substantial impact upon the intrinsic and extrinsic (growth factor-induced) activation of the STAT3 pathway. This indicates an additional role for biophysical factors in regulating this critical signaling pathway. Interactions between pathways conveying information from both soluble mediators and the ECM are integrated at the level of the cytoskeleton. In this context, the role of cytoskeletal tension has been likened to a cellular rheostat, acting to dampen or augment the responses of multiple signaling pathways to growth factor stimulation, thereby blocking or facilitating mitogenic responses.

It has been proposed that the ECM is a critical regulator of cellular dormancy27; however the role of matrix stiffness in regulating this process has not been specifically addressed. The growth, invasion and dissemination of tumor cells are accompanied by dramatic changes in the mechanical properties (stiffness) of the cancer cell niche. The bone marrow, a common reservoir site for disseminated tumor cells, provides a microenvironment with stiffness significantly lower than that encountered in most epithelial tumors.28 Our findings suggest that a reduction in the stiffness of the cancer cell niche would be sufficient to promote reversible cellular quiescence (dormancy). Furthermore, increases in environmental stiffness (as may occur with inflammation, surgery or stromal reaction to tumor) or alteration in the stiffness-sensing machinery of the cell (as a result of genomic instability) might facilitate reactivation. Indeed, early work on cancer cell dormancy in animal models established inflammation and surgical trauma as a mechanism of reactivation of dormant cells.29, 30 More recently, fibrosis-associated collagen-I has been linked to reactivation of tumor cells in an in vivo model of cancer cell dormancy.31 With respect to the liver, tumor growth and intrahepatic metastasis have been shown to be enhanced in a fibrotic environment.20-22

The precise phenotype of disseminated tumor cells derived from epithelial malignancies remains poorly defined.32 During tumor dissemination, tumor cells are believed to acquire mesenchymal properties, enabling them to migrate through and invade surrounding tissues and enter the bloodstream.33 However, at secondary sites, tumor cells are primarily detected by their epithelial characteristics and outgrowing metastases recapitulate the epithelial phenotype of the primary tumor.34 Despite our increasing understanding of the regulation of epithelial-to-mesenchymal transition, the reverse process—mesenchymal-to-epithelial transition—is largely uncharacterized. We have demonstrated that HCC cells lose mesenchymal features, including stress fibers, N-cadherin, and vimentin expression, and the cells up-regulate markers of hepatocyte differentiation when maintained in a soft environment. This is consistent with previous findings showing that nontransformed mammary epithelial cells revert to an organized epithelial phenotype in a soft environment6 and that hepatocytes retain an epithelial phenotype on soft collagen gels.35 FAK activation has been implicated in the process of epithelial-to-mesenchymal transition and responsiveness to TGFβ.36 It remains unclear whether reduced FAK activation and TGFβ signaling in cells in a low stiffness environment is a mechanistic link to mesenchymal-to-epithelial transition.

The high rate of chemotherapy resistance in HCC is a major obstacle in treating patients with advanced disease. Identifying the mechanism of this resistance has the potential to reveal new treatment options for this group of patients. We have provided evidence that increasing ECM stiffness, as encountered by cells within an established tumor,4 reduces chemotherapy-induced apoptosis. However, the clinical utility of systemic chemotherapy is also limited by the failure of adjuvant/neoadjuvant chemotherapy to target disseminated tumor cells that give rise to late tumor recurrence and metastases.32, 37 Intriguingly, we have been able to demonstrate an increase in clone-initiating capability following chemotherapy in cells from a low stiffness environment. This was accompanied by an increase of cells positive for cancer stem cell markers (CD44, CD133, c-kit, CXCR-4, OCT4, and NANOG).38 This provides a potential mechanism for long-term survival and clone-initiating capability of disseminated tumor cells in a soft environment (e.g., bone marrow) following chemotherapy. Whether the higher frequency of cells with a cancer stem cell phenotype is due to positive selection or active induction of cancer stem cell characteristics needs to be determined.

In summary, we have provided evidence that the biomechanical composition of the ECM is a critical regulator of HCC behavior. We suggest that the high stiffness environment encountered in chronic fibrotic liver disease fosters HCC progression by promoting cellular proliferation, a mesenchymal phenotype and resistance to chemotherapy. Conversely, a soft physiological environment (as might be encountered by a disseminated tumor cell) induces cellular dormancy, a stem cell phenotype and enhanced clonogenic capacity following chemotherapy. Indeed, we propose that alterations in the stiffness of the cancer cell niche are responsible for regulating cancer cell proliferation and phenotype throughout the natural history of HCC. Manipulation of environmental stiffness or interference with the stiffness-sensing apparatus of HCC cells has the potential to impede both tumor growth and reactivation of dormant tumor cells, thereby limiting recurrence. In concert with future in vivo models of HCC, these findings will provide a platform for the future design of therapies targeting the biomechanical properties of the cancer cell niche.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

The authors would like to acknowledge the assistance of Dr. David Hay (University of Edinburgh), who was supported by a fellowship from the Research Council UK, for his intellectual input with respect to experimental design. They would also like to thank Prof. Margaret Frame (University of Edinburgh) and Dr. Jim Norman (University of Glasgow) for advice in respect to experimental reagents.

References

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Additional Supporting Information may be found in the online version of this article.

FilenameFormatSizeDescription
HEP_24108_sm_suppinfofigure1.tif104KSupporting Information Figure 1; Figure S1: The effect of matrix stiffness on HCC cell spreading: a time-course analysis
HEP_24108_sm_suppinfofigure2.tif97KSupporting Information Figure 2
HEP_24108_sm_suppinfofigure3.tif1115KSupporting Information Figure 3
HEP_24108_sm_suppinfofigure4.tif84KSupporting Information Figure 4
HEP_24108_sm_suppinfofigure5.tif105KSupporting Information Figure 5
HEP_24108_sm_suppinfofigure6.tif107KSupporting Information Figure 6
HEP_24108_sm_suppinfofigure7.tif595KSupporting Information Figure 7
HEP_24108_sm_suppinfofigure8.tif468KSupporting Information Figure 8
HEP_24108_sm_suppinfofigure9.tif109KSupporting Information Figure 9
SupplementalFigureLegends.pdf42KSupplementary Figure legends
SupplementalMethods.pdf39KSupplemental Methods

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