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Ex vivo aorta patch model for analysis of cellular adhesion
Tissue and Cell 47 (2015) 266–272
Contents lists available at ScienceDirect
Tissue and Cell journal homepage: www.elsevier.com/locate/tice
Ex vivo aorta patch model for analysis of cellular adhesion Yuan-Na Lin a,b,1 , Raymond Nqobizitha Thata b,1 , Antonio Virgilio Failla c , Markus Geissen d , Guenter Daum d,1 , Sabine Windhorst b,∗,1 a
Department of General, Visceral and Thoracic Surgery, University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany Department of Biochemistry and Signal Transduction, University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany Microscopy Imaging Facility, University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany d Department of Vascular Medicine, University Heart Center, Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany b c
a r t i c l e
i n f o
Article history: Received 26 December 2014 Received in revised form 11 March 2015 Accepted 13 March 2015 Available online 23 March 2015 Keywords: Endothelial cells Vascular model Aorta Cell adhesion
a b s t r a c t The vascular endothelium as well as subendothelium are objects of many researches as it is directly involved in a multiplicity of physiological and pathological settings. Detailed study of endothelial function became feasible with the development of techniques to culture endothelial cells (EC) in vitro. Limitations of this approach have become apparent with the realization that cell culture dedifferentiate with time and do not exhibit properties of intact tissue. Here we describe the development of a novel ex vivo tissue model to study cell–vascular wall interactions by using isolated mouse aorta patches. Validation of this model was performed by demonstrating cell attachment and changes in cell shape typical for cell spreading during adhesion. A major advantage of this model is that cell–endothelium interaction and its molecular backgrounds can now be studied more feasibly on an intact and native tissue. © 2015 Elsevier Ltd. All rights reserved.
1. Introduction The functions of the endothelium (EC) and subendothelium (SEC), which built up the inner layer of all blood vessels, are multiple and do not just form a passive barrier. EC and SEC are actively involved in fluid and solute exchange, regulation of perfusion, inflammatory responses, blood coagulation and not at last in malignant cell arrest and extravasation processes (reviewed in Cines et al., 1998; Pinkney et al., 1997). Aberrant EC function can therefore lead to pathological conditions involving atherosclerosis, hypertension, vasculitis, thromboembolic diseases, tumor onset and metastasis (reviewed in Bouïs et al., 2001; Cines et al., 1998; Gallagher and Sumpio, 1997; Kramer and Nicolson, 1979; Ruoslahti and Rajotte, 2000). The growing interest for the endothelium in physiological and pathological conditions has led to an increased
Abbreviations: EC, endothelial cells; SEC, subendothelial cells; MEC, primary mouse endothelial cells. ∗ Corresponding author at: Institute of Biochemistry and Signal Transduction, University Medical Center Hamburg-Eppendorf, Martinistraße 52, D-20246 Hamburg, Germany. Tel.: +49 40 7410 56341; fax: +49 40 7410 56818. E-mail addresses: [email protected] (Y.-N. Lin), [email protected] (R.N. Thata), [email protected] (A.V. Failla), [email protected] (M. Geissen), [email protected] (G. Daum), [email protected] (S. Windhorst). 1 These authors contributed equally to the work. http://dx.doi.org/10.1016/j.tice.2015.03.004 0040-8166/© 2015 Elsevier Ltd. All rights reserved.
demand for representative model systems. Unfortunately, the direct assessment of vascular tissue in vivo was shown to be difficult. But without such tissue, the endothelial cell’s contribution to disease development can only be deduced. In the last decades, in vitro EC cultures have emerged to be important tools for studying vascular physiology and disease pathology (Jaffe et al., 1973; Lewis et al., 1973; Gimbrone et al., 1974; Bachetti and Morbidelli, 2000; reviewed in Cines et al., 1998). Due to numerous transgenic mouse lines, the isolation and culture of mouse ECs is of particular interest. And as many in vivo data were derived from mice, the results obtained with animal cells may be more easily compared with. However, the isolation and maintenance of primary mouse endothelial cells (MEC) continues to be challenging and time-consuming 1 (Bachetti and Morbidelli, 2000). Moreover, primary EC feature a series of disadvantages, such as cell senescence, heterogeneity of EC donors, the loss of their primary characteristics and responsiveness to various stimuli (Erusalimsky, 2009; Bouïs et al., 2001). Therefore, development of stable, immortalized MEC lines that retain the characteristics of endothelial cells was expected to greatly facilitate endothelial biology and pathology research (reviewed in Bouïs et al., 2001). Since in general immortalization is achieved by transfection with an oncogene, immortalized cells show tumor cell characteristics. Thus, an immortalized cell line is a compromise between the presentation of the desired primary characteristics and tumor cell traits (Hohenwarter et al., 1994).
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Fig. 1. Schematic illustration of the ex vivo aorta tissue model for analysis of tumor cell adhesion. (1) Isolation of Carotid artery from mice; (2a) cutting of blood vessel in roughly rectangular shapes; (2b) pouring of silicone elastomer; (3) using metal pins to secure blood vessel patches onto the wells; (4) patches were kept in DMEM (10% FCS, 1% P/S) at 37 ◦ C and 5% CO2 ; a metal ring was used to isolate the patches from the well; (5) cancer cells (HCT-116) were added to the patches; (6) incubation over several hours or overnight; (7) removal of metal ring; (8) careful staining of actin; (9) mounting of the patches on slides; (10) microscopy of slides.
The objective of this study was to develop a new model for investigation of mouse aortic endothelium by overcoming the disadvantages of hitherto primary and immortalized endothelial cell lines. For that purpose, we established an ex vivo aorta pin tissue model of mouse aortic patches to perform analysis of cell-toendothelium interactions. We conclude that this model is not only applicable for analysis of cell adhesion processes on the vascular endothelium, but would also enable further studies with prospect of more intense investigations of the vascular endothelium.
2. Materials and methods 2.1. Cell line HCT-116 cells were obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (Braunschweig, Germany). The cell line HCT-116 was cultured in Dulbecco‘s modified Eagle‘s medium (DMEM, Gibco), supplemented with 10% (v/v) fetal calf serum (FCS), 100 g/ml streptomycin and 100 units/ml penicillin. Cells were cultured at 37 ◦ C in a humidified incubator with 5% CO2 .
2.2. Isolation and plating of mouse aortal patches Mice (FVB/NCrl (Charles River, USA); male; 11 weeks) were obtained from the animal facility (UKE, Hamburg, Germany). All experimental procedures comply with the German Animal Welfare Act and the European Guideline EU 2010/63. The procedures have been approved by the Animal Welfare Officer of the Institution and the Authorities the ı´Behörde für Gesundheit und Verbraucherschutz´ı of the City of Hamburg. Mice were killed by CO2 intoxication, the sternum was opened and blood was removed by injection of Ringer solution into the left ventricle after opening of the Vena cava inferior. The aorta thoracalis was detached from the spinal cord by cutting the branches and attaching tissue was removed with care. The aorta was collected in Ringer solution on ice, cut into three equal pieces and then the ring was opened carefully by a longitudinal incision. The aorta tissue from mice was transferred to a plate containing Hanks Balanced Salt Solution (HBSS). Extraneous tissue surrounding the aorta was removed, and the ends of the aorta were cut and discarded. Additionally the aorta was cut into three nearly identical pieces. The aorta pieces were then relocated into 6-well plates coated with a silicone elastomer (SylgardTM 184 Silicone Elastomer; Dow Corning; Belgium).
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Fig. 2. Cellular integrity of aortic tissue patches. (A) Aorta tissue cells were transfected with a retroviral vector encoding for eGFP to label the cells. 24 h after treating the tissue with the virus, the patches were fixed and analyzed by fluorescence microscopy. Left panel: Outer cell layer, containing the endothelial cells. Middle panel: The presence of endothelial cells was validated by staining the tissue for the endothelial marker protein CD31. Left panel: Subendothelial layer, containing fibroblasts and collagen fibers. (B) Alamar blue Assay was performed to analyze proliferation rate within 72 h of incubation. Shown is one representative experiment. Bar: 10 m.
Using insect pins the aorta patches were pinned on the coated wells, keeping them roughly half a centimeter above the surface. HBSS was then removed and a metal ring was placed around the tissue. 200 l of culture media (DMEM, 10% FCS, 1% P/S, 1% NEAA) was added into the metal ring. HCT-116 cells were counted using a CASY Counter and then 5 × 104 cells were added to the wells. Pin culture with added cell suspension were incubated at 37 ◦ C in a humidified incubator with 5% CO2 (Fig. 1). 2.3. Staining of mouse aortal patches All steps were performed in the culture dish. The metal ring was removed and the tissue was washed (3×; PBS; 37 ◦ C) with caution by swinging the culture plate (∼40◦ ) 2 times. After fixation (10 min; 3.7% Formalin in phosphate buffer; pH 7.0) and three washing
steps the cultured aorta was incubated in TBST (3×; 5 min). Prior to antibody staining tissues were incubated for 20 min with Antibody diluent (Antibody Diluent Reagent Solution Ready-to-use; Invitrogen; USA). Primary antibody (anti-CD31 antibody; abcam, ab28364; anti-E-cadherin antibody; BD Biosciences, 610181; antiIntegrin 1 antibody; Santa Cruz, sc-73610; all 1/200 in Antibody diluent) was incubated overnight at 4 ◦ C to identify the human cells. After 3 washes (5 min; PBS) incubation with the secondary antibody (anti-mouse Rhodamine (goat; Jackson ImmunoResearch (115-295-062); 1/1000 in Antibody diluent) was performed. After 3 washes (5 min; TBST) the tissues were transferred to glass slides (inside up) and mounted with Fluoromount-G (SouthernBiotech; USA). For staining actin filaments, patches were incubated with Alexa Fluor® 568 (Life Technologies; diluted 1:500) and incubated at room temperature for 30 min. After 3-times washing with PBS,
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Fig. 3. Colon carcinoma cells adhere to aorta patches. (A) HCT-116 cells expressing eGFP were seeded to freshly prepared aorta patches and their eGFP fluorescence was monitored by fluorescence microscopy 24 h after seeding. eGFP-fluorescence of HCT-116 cells (shown in green) was separated from auto-fluorescence of fibroblasts (shown in red) at confocal microscope Olympus SP5. (B) Comparison of cell–cell contacts (anti-E-cadherin) and cell–matrix contacts (anti-integrin beta1) of HCT-116 cells on the aorta patches (left panels) and on collagen-coated chamber slides (right panels). Bars: 10 m. (For interpretation of reference to color in this figure legend, the reader is referred to the web version of this article.)
the aorta pieces were transferred to glass slides (inside up) and mounted with Fluoromount-G.
2.4. Cell viability assay For the Alamar blue cell viability assay, the isolated aorta patches were incubated up to 3 days. Cell medium alone served as negative control, from which the background fluorescence was subtracted. Measurements were performed after incubation of the patches for 24, 48 and 72 h at 37 ◦ C in a humidified atmosphere. Fluorescence based absorption was measured at 540 nm on a micro plate reader (Tekan, Switzerland).
2.5. Generation of viral particles and eGFP transduction of HCT-116 cells Viral particles were produced as cell-free supernatants by transient transfection of the packaging cell line HEK-293T as described (Weber et al., 2008, 2012). In brief, the lentiviral vector LeGO-iG2Luc2 expressing eGFP-cDNA (Firefly luciferase, Promega) was packaged using the third-generation packaging plasmids pMDLg/pRRE (Addgene.org #12251), pRSV-Rev (Addgene.org #12253) and phCMV-VSV-G expressing the envelope protein of vesicular stomatitis virus. The supernatant was harvested 24 h after transfection, 0.45 m filtrated and stored at −80 ◦ C. Titration on 293T cells using 8 g/ml polybrene and spin-inoculation (1000 × g, 1 h, 25 ◦ C) resulted in a functional titer of 5.2 × 107 TU per ml. HCT-116 target
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Fig. 4. Spreading of HCT-116 cells on aorta patches. HCT-116 cells were seeded to aorta patches, fixed after 2, 6 or 16 h (overnight) and stained with Alexa Fluor® 568-coupled phalloidin. The right panels show higher magnifications. Bars: 10 m.
cells were plated at 5 × 104 cells in 0.5 ml medium in each well of a 24 well plate. After addition of 30 l of viral particle containing supernatant to the cells, the medium was replaced the next day. eGFP-ITPKA transduction of the patches was performed with the retroviral vector MigRI as described in Windhorst et al. (2010). 2.6. Microscopy For analysis of eGFP-fluorescence by fluorescence microscopy, the patch was carefully transferred into a chamber slide and cells were fixed with 4% paraformaldehyde. All the light microscopy images were acquired with a Leica TCS SP5 confocal microscope permitting the acquisition of accurate three-dimensional data set. Dual channel confocal imaging was performed. In channel number one eGFP positive cell were imaged using a 488 nm Argon laser
line as excitation source and a novel state of the art GaAsP hybrid detection system Leica HyDTM set to collect all the light coming with the 500–520 nm spectral band. In this channel both specific eGFP signals and unspecific auto fluorescence signals can be differentiated however as eGFP specific signals are one order of magnitude stronger than auto fluorescence, which made it possible to clearly the identify eGFP-expressing cells. In the second channel auto fluorescence from all cells was excited by a Dipole Pumped Solid State DPSS 561 nm laser source and detected with a second state-ofthe-art GaAsP hybrid detection system Leica HyDTM set to collect all the light coming with the 565–747 nm spectral band. In this spectral band only auto fluorescence is detectable being the eGFP positive signal negligible. Auto fluorescence emission spectra upon excitation at 488 and 561 nm were acquired (data not shown) to characterize the optical properties of the system. For accurate 3D
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reconstruction images in 246 × 246 × (20–50) m3 volume were acquired using a pixel size of 240 nm (1024 × 1024) pixel images and a voxel height either of 500 nm or of 1 m. The pixel size of (0.240 × 0.240 × (0.500–1)) m3 is sufficiently accurate to resolve cellular structures. The acquired image stacks were processed by mean of Bitplane Imaris software that allowed 3D reconstruction and accurate image visualization after automatic segmentation routine. 3. Results and discussion 3.1. Establishment of an adhesion assay using aortic patches as substrate To develop a cell adhesion assay that is feasible for routine analyses and that shows all the benefits of intact tissue, we have chosen in situ mouse aortic endothelium as a substrate. Mouse aorta is a robust vessel of suitable size, it is readily available and easy to prepare. Isolated thoracic mouse aortas were longitudinally cut and pinned inside out onto petri dishes that had been coated with a silicone elastomer. Next, a metal ring was placed to enclose the patch and defined numbers (5 × 104 ) of tumor cells were then seeded onto the patch (Fig. 1). In order to prevent de-differentiation of vessel cells, we did not cultivate the patches longer than three days. In conclusion, we established a vessel model that is easily to prepare and stable for three days. 3.2. Validation of cellular integrity of aortic tissue patches In order to analyze if the different cell layers of the aortic patch remained intact after preparation, the tissue cells were transfected with a retroviral vector encoding for eGFP to label the cells. 24 h after treating the tissue with the virus, the patches were fixed and analyzed by fluorescence microscopy. Thereby, we first focused on the outer cell layer containing the endothelial cells, and thereafter on the subendothelial layer containing fibroblasts and collagen fibers. As shown in Fig. 2A the transfection efficiency of tissue cells was very high, and thus the endothelial were well visible at the outer cell layer (left panel). Verification was performed by staining the cells with the endothelial marker protein CD31 (middle panel; Miettinen et al., 1994). The subendothelial layer containing fibroblasts and collagen fibers was detectable when focusing behind the endothelial cells (right panel). However, it was not possible to focus behind the subendothelial layer to detect smooth muscle cells. To show if the tissue cells start to dedifferentiate during cultivation, proliferation was analyzed by the Alamar blue Assay for 72 h, every 24 h. We found no proliferation within 72 h of incubation, but after 72 h cell viability decreased (Fig. 2B). Therefore, we recommend keeping the tissue only for 48 h in culture. In summary our data show that the aortic tissue cells do not dedifferentiate and remain stable for at least 48 h. Furthermore, the endothelial and subendothelial cell layers remain intact after preparation. Thus, the ex vivo model should be suitable to analyze interaction of blood cells (circulating tumor cells, immune cells and plates) with endothelial and with subendothelial cells. In addition, our model enables to genetically modify the vascular cells, thus provides the possibility to analyze the function of proteins of interest for endothelial–blood cell interaction. 3.3. Interaction of vascular cells with tumor cells Since the interaction of circulating tumor cells with endothelial and subendothelial cells is one essential factor that determines if the tumor cells adhere to the vascular wall or not, we analyzed if our
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Table 1 Spreading of HCT-116 cells on aorta patches. HCT-116 cells were seeded on aorta patches and filopodia length and cell area were measured 2 h, 6 h and 16 h after seeding. Values obtained from cells incubated for 16 h were set to 100%. Measurements were done at least twice, at least 80 cells were analyzed per measurement. Time (h)
Filopodia length, %
Cell area, %
2 6 16
Not detectable 88 ± 6 100
53 ± 9 65 ± 17 100
model is suitable to observe tumor cell–endothelial/subendothelial interactions. The prerequisite for such an analysis is that suspended tumor cells adhere to the isolated aortic tissue. Therefore, we first tested adherence of the established colon cancer cell line HCT-116 on aortic patches (Lin et al., 2014). Suspended eGFP-expressing HCT-116 cells were seeded on freshly isolated tissue and one day after incubation, the cells were fixed with paraformaldehyde and embedded in mountain-medium. We found that one day after seeding the cells had adhered to the tissue (Fig. 3A), the apparently visible invasion in Fig. 3A (lower panel – Z-axis) results from folding of the tissue. To analyze potential differences of HCT-116 cell adhesion when seeded on collagen-coated chamber slides or on aorta patches, we stained the cells for E-cadherin to analyze cell–cell contacts. Cell–matrix contacts were examined by staining the cells for integrin-, the main protein of focal adhesions in motile cells (Ivaska et al., 2002). Our data show that cell–cell interactions are similar when cells were seeded on collagen or on vascular tissue (Fig. 3B). Under both conditions, the cells grow as tight clusters. However, the interaction of tumor cells with the matrix seemed to be different. When seeded on collagen-coated chamber slides, the single cells formed multiple focal adhesions that were equally distributed over the cells and additionally, they form large and very distinct focal adhesions at the periphery of the clusters. These additional focal adhesions were not formed when the cells were seeded on vascular tissue. In conclusion, our results reveal that suspended epithelial tumor cells adhere to our vascular patch tissue and show that cell-to-matrix interactions are different when the cells grow on collagen-covered chamber slides or on vascular tissue. 3.4. Spreading of tumor cells on vascular tissue Next, we examined if it is possible to monitor the process of tumor cell spreading (cell flattening and formation of cellular protrusions (Nürnberg et al., 2011) on the vascular patch tissue. Therefore, cells were fixed 2 h, 6 h and 16 h after seeding, and for labeling F-actin the cells were stained with Alexa Fluor® 568 phalloidin. Analysis by fluorescence microscopy revealed that 2 h after seeding most cells were round-shaped and did not form filopodia. After 6 h, most of the cells were still rounded up, but cells had formed filopodia. When the cells were incubated for 16 h, they flattened onto the substratum. Although filopodia-length did not significantly increase in comparison to cells cultivated for six hours, cells started to form lamellipodia (Fig. 4, Table 1). In conclusion, these data show that our ex vivo model provides the possibility to precisely analyze morphological changes during epithelial cell spreading on the aortic endothelial cell layer. In summary, our ex vivo model provides the possibility to analyze interaction of blood cells with endothelial and subendothelial cells. The main advantage of this model is that, in contrast to cultivated endothelial cells, interaction of blood cells with vascular tissue can be examined. In addition, our model provides the possibility to genetically modify both, the cells of the vascular tissue and the blood cells, enabling to analyze the impact of proteins on vascular wall–blood cell interactions. We demonstrated that suspended
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tumor cells adhere to the vascular tissue very well, and we show that it is possible to analyze the process of cell spreading. Since the mechanism of tumor cell spreading on vascular cells has not been elucidated in detail yet, our model now provides the possibility to analyze the impact of proteins involved in this process. Thus, our ex vivo model provides a feasible alternative for analyzing vascular wall–blood cell interactions of cultivated endothelial cells under static conditions. Acknowledgments We thank Christine Blechner for excellent technical support. This work was supported by a UCCH grant (University Cancer Center Hamburg). References Bachetti, T., Morbidelli, L., 2000. Endothelial cells in culture: a model for studying vascular functions. Pharmacol. Res. 42 (1), 9–19. Bouïs, D., Hospers, G.A., Meijer, C., et al., 2001. Endothelium in vitro: a review of human vascular endothelial cell lines for blood vessel-related research. Angiogenesis 4 (2), 91–102. Cines, D.B., Pollak, E.S., Buck, C.A., et al., 1998. Endothelial cells in physiology and in the pathophysiology of vascular disorders. Blood 91 (10), 3527–3561. Erusalimsky, J.D., 2009. Vascular endothelial senescence: from mechanisms to pathophysiology. J. Appl. Physiol. 106, 326–332. Gallagher, G.L., Sumpio, B.E., 1997. In: Kisco, Mt. (Ed.), Endothelial Cells. Futura Publishing Company, New York, pp. 151–186. Gimbrone, M.A., Cotran, R.S., Folkman, J., 1974. Human vascular endothelial cells in culture. Growth and DNA synthesis. J. Cell Biol. 60, 673.
Hohenwarter, O., Jakoubek, A., Schmatz, C., et al., 1994. Expression of SV40 tumor antigens enables human endothelial cells to grow independently from foetal calf serum end exogenous growth factors. J. Biotechnol. 34, 205–211. Ivaska, J., Whelan, R.D., Watson, R., et al., 2002. PKC epsilon controls the traffic of beta1 integrins in motile cells. EMBO J. 21, 3608–3619. Jaffe, E.A., Nachman, R.L., Becker, C.G., Minick, C.R., 1973. Culture of human endothelial cells derived from umbilical veins: Identification by morphologic criteria. J. Clin. Invest. 52, 2745. Kramer, R.H., Nicolson, G.L., 1979. Interactions of tumor cells with vascular endothelial cell monolayers: a model for metastatic invasion. Proc. Natl. Acad. Sci. U. S. A. 76, 5704–5708. Lewis, L.J., Hoak, J.C., Maca, R.D., Fry, G.L., 1973. Replication of human endothelial cells in culture. Science 181, 453. Lin, Y., Izbicki, J.R., König, A., et al., 2014. Expression of DIAPH1 is up-regulated in colorectal cancer and its down-regulation strongly reduces the metastatic capacity of colon carcinoma cells. Int. J. Cancer 134, 1571–1582. Miettinen, M., Lindenmayer, A.E., Chaubal, A., 1994. Endothelial cell markers CD31, CD34, and BNH9 antibody to H- and Y-antigens-evaluation of their specificity and sensitivity in the diagnosis of vascular tumors and comparison with von Willebrand factor. Mod. Pathol. 7 (1), 82–90. Nürnberg, A., Kitzing, T., Grosse, R., 2011. Nucleating actin for invasion. Nat. Rev. Cancer 11 (3), 177–187. Pinkney, J.H., Stehouwer, C.D., Coppack, S.W., Yudkin, J.S., 1997. Endothelial dysfunction: cause of the insulin resistance syndrome. Diabetes 46 (Suppl. 2), S9–S13. Ruoslahti, E., Rajotte, D., 2000. An address system in the vasculature of normal tissues and tumors. Annu. Rev. Immunol. 18, 813–827. Weber, K., Bartsch, U., Stocking, C., Fehse, B., 2008. A multicolor panel of novel lentiviral gene ontology (LeGO) vectors for functional gene analysis. Mol. Ther. 16, 698–706. Weber, K., Thomaschewski, M., Benten, D., Fehse, B., 2012. RGB marking with lentivirale vectors for multicolor clonal cell tracking. Nat. Protoc. 5, 839–849. Windhorst, S., Fliegert, R., Blechner, C., et al., 2010. Inositol 1,4,5-trisphosphate 3kinase-A is a new cell motility-promoting protein that increases the metastatic potential of tumor cells by two functional activities. J. Biol. Chem. 285 (8), 5541–5554.
Contents lists available at ScienceDirect
Tissue and Cell journal homepage: www.elsevier.com/locate/tice
Ex vivo aorta patch model for analysis of cellular adhesion Yuan-Na Lin a,b,1 , Raymond Nqobizitha Thata b,1 , Antonio Virgilio Failla c , Markus Geissen d , Guenter Daum d,1 , Sabine Windhorst b,∗,1 a
Department of General, Visceral and Thoracic Surgery, University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany Department of Biochemistry and Signal Transduction, University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany Microscopy Imaging Facility, University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany d Department of Vascular Medicine, University Heart Center, Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany b c
a r t i c l e
i n f o
Article history: Received 26 December 2014 Received in revised form 11 March 2015 Accepted 13 March 2015 Available online 23 March 2015 Keywords: Endothelial cells Vascular model Aorta Cell adhesion
a b s t r a c t The vascular endothelium as well as subendothelium are objects of many researches as it is directly involved in a multiplicity of physiological and pathological settings. Detailed study of endothelial function became feasible with the development of techniques to culture endothelial cells (EC) in vitro. Limitations of this approach have become apparent with the realization that cell culture dedifferentiate with time and do not exhibit properties of intact tissue. Here we describe the development of a novel ex vivo tissue model to study cell–vascular wall interactions by using isolated mouse aorta patches. Validation of this model was performed by demonstrating cell attachment and changes in cell shape typical for cell spreading during adhesion. A major advantage of this model is that cell–endothelium interaction and its molecular backgrounds can now be studied more feasibly on an intact and native tissue. © 2015 Elsevier Ltd. All rights reserved.
1. Introduction The functions of the endothelium (EC) and subendothelium (SEC), which built up the inner layer of all blood vessels, are multiple and do not just form a passive barrier. EC and SEC are actively involved in fluid and solute exchange, regulation of perfusion, inflammatory responses, blood coagulation and not at last in malignant cell arrest and extravasation processes (reviewed in Cines et al., 1998; Pinkney et al., 1997). Aberrant EC function can therefore lead to pathological conditions involving atherosclerosis, hypertension, vasculitis, thromboembolic diseases, tumor onset and metastasis (reviewed in Bouïs et al., 2001; Cines et al., 1998; Gallagher and Sumpio, 1997; Kramer and Nicolson, 1979; Ruoslahti and Rajotte, 2000). The growing interest for the endothelium in physiological and pathological conditions has led to an increased
Abbreviations: EC, endothelial cells; SEC, subendothelial cells; MEC, primary mouse endothelial cells. ∗ Corresponding author at: Institute of Biochemistry and Signal Transduction, University Medical Center Hamburg-Eppendorf, Martinistraße 52, D-20246 Hamburg, Germany. Tel.: +49 40 7410 56341; fax: +49 40 7410 56818. E-mail addresses: [email protected] (Y.-N. Lin), [email protected] (R.N. Thata), [email protected] (A.V. Failla), [email protected] (M. Geissen), [email protected] (G. Daum), [email protected] (S. Windhorst). 1 These authors contributed equally to the work. http://dx.doi.org/10.1016/j.tice.2015.03.004 0040-8166/© 2015 Elsevier Ltd. All rights reserved.
demand for representative model systems. Unfortunately, the direct assessment of vascular tissue in vivo was shown to be difficult. But without such tissue, the endothelial cell’s contribution to disease development can only be deduced. In the last decades, in vitro EC cultures have emerged to be important tools for studying vascular physiology and disease pathology (Jaffe et al., 1973; Lewis et al., 1973; Gimbrone et al., 1974; Bachetti and Morbidelli, 2000; reviewed in Cines et al., 1998). Due to numerous transgenic mouse lines, the isolation and culture of mouse ECs is of particular interest. And as many in vivo data were derived from mice, the results obtained with animal cells may be more easily compared with. However, the isolation and maintenance of primary mouse endothelial cells (MEC) continues to be challenging and time-consuming 1 (Bachetti and Morbidelli, 2000). Moreover, primary EC feature a series of disadvantages, such as cell senescence, heterogeneity of EC donors, the loss of their primary characteristics and responsiveness to various stimuli (Erusalimsky, 2009; Bouïs et al., 2001). Therefore, development of stable, immortalized MEC lines that retain the characteristics of endothelial cells was expected to greatly facilitate endothelial biology and pathology research (reviewed in Bouïs et al., 2001). Since in general immortalization is achieved by transfection with an oncogene, immortalized cells show tumor cell characteristics. Thus, an immortalized cell line is a compromise between the presentation of the desired primary characteristics and tumor cell traits (Hohenwarter et al., 1994).
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Fig. 1. Schematic illustration of the ex vivo aorta tissue model for analysis of tumor cell adhesion. (1) Isolation of Carotid artery from mice; (2a) cutting of blood vessel in roughly rectangular shapes; (2b) pouring of silicone elastomer; (3) using metal pins to secure blood vessel patches onto the wells; (4) patches were kept in DMEM (10% FCS, 1% P/S) at 37 ◦ C and 5% CO2 ; a metal ring was used to isolate the patches from the well; (5) cancer cells (HCT-116) were added to the patches; (6) incubation over several hours or overnight; (7) removal of metal ring; (8) careful staining of actin; (9) mounting of the patches on slides; (10) microscopy of slides.
The objective of this study was to develop a new model for investigation of mouse aortic endothelium by overcoming the disadvantages of hitherto primary and immortalized endothelial cell lines. For that purpose, we established an ex vivo aorta pin tissue model of mouse aortic patches to perform analysis of cell-toendothelium interactions. We conclude that this model is not only applicable for analysis of cell adhesion processes on the vascular endothelium, but would also enable further studies with prospect of more intense investigations of the vascular endothelium.
2. Materials and methods 2.1. Cell line HCT-116 cells were obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (Braunschweig, Germany). The cell line HCT-116 was cultured in Dulbecco‘s modified Eagle‘s medium (DMEM, Gibco), supplemented with 10% (v/v) fetal calf serum (FCS), 100 g/ml streptomycin and 100 units/ml penicillin. Cells were cultured at 37 ◦ C in a humidified incubator with 5% CO2 .
2.2. Isolation and plating of mouse aortal patches Mice (FVB/NCrl (Charles River, USA); male; 11 weeks) were obtained from the animal facility (UKE, Hamburg, Germany). All experimental procedures comply with the German Animal Welfare Act and the European Guideline EU 2010/63. The procedures have been approved by the Animal Welfare Officer of the Institution and the Authorities the ı´Behörde für Gesundheit und Verbraucherschutz´ı of the City of Hamburg. Mice were killed by CO2 intoxication, the sternum was opened and blood was removed by injection of Ringer solution into the left ventricle after opening of the Vena cava inferior. The aorta thoracalis was detached from the spinal cord by cutting the branches and attaching tissue was removed with care. The aorta was collected in Ringer solution on ice, cut into three equal pieces and then the ring was opened carefully by a longitudinal incision. The aorta tissue from mice was transferred to a plate containing Hanks Balanced Salt Solution (HBSS). Extraneous tissue surrounding the aorta was removed, and the ends of the aorta were cut and discarded. Additionally the aorta was cut into three nearly identical pieces. The aorta pieces were then relocated into 6-well plates coated with a silicone elastomer (SylgardTM 184 Silicone Elastomer; Dow Corning; Belgium).
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Fig. 2. Cellular integrity of aortic tissue patches. (A) Aorta tissue cells were transfected with a retroviral vector encoding for eGFP to label the cells. 24 h after treating the tissue with the virus, the patches were fixed and analyzed by fluorescence microscopy. Left panel: Outer cell layer, containing the endothelial cells. Middle panel: The presence of endothelial cells was validated by staining the tissue for the endothelial marker protein CD31. Left panel: Subendothelial layer, containing fibroblasts and collagen fibers. (B) Alamar blue Assay was performed to analyze proliferation rate within 72 h of incubation. Shown is one representative experiment. Bar: 10 m.
Using insect pins the aorta patches were pinned on the coated wells, keeping them roughly half a centimeter above the surface. HBSS was then removed and a metal ring was placed around the tissue. 200 l of culture media (DMEM, 10% FCS, 1% P/S, 1% NEAA) was added into the metal ring. HCT-116 cells were counted using a CASY Counter and then 5 × 104 cells were added to the wells. Pin culture with added cell suspension were incubated at 37 ◦ C in a humidified incubator with 5% CO2 (Fig. 1). 2.3. Staining of mouse aortal patches All steps were performed in the culture dish. The metal ring was removed and the tissue was washed (3×; PBS; 37 ◦ C) with caution by swinging the culture plate (∼40◦ ) 2 times. After fixation (10 min; 3.7% Formalin in phosphate buffer; pH 7.0) and three washing
steps the cultured aorta was incubated in TBST (3×; 5 min). Prior to antibody staining tissues were incubated for 20 min with Antibody diluent (Antibody Diluent Reagent Solution Ready-to-use; Invitrogen; USA). Primary antibody (anti-CD31 antibody; abcam, ab28364; anti-E-cadherin antibody; BD Biosciences, 610181; antiIntegrin 1 antibody; Santa Cruz, sc-73610; all 1/200 in Antibody diluent) was incubated overnight at 4 ◦ C to identify the human cells. After 3 washes (5 min; PBS) incubation with the secondary antibody (anti-mouse Rhodamine (goat; Jackson ImmunoResearch (115-295-062); 1/1000 in Antibody diluent) was performed. After 3 washes (5 min; TBST) the tissues were transferred to glass slides (inside up) and mounted with Fluoromount-G (SouthernBiotech; USA). For staining actin filaments, patches were incubated with Alexa Fluor® 568 (Life Technologies; diluted 1:500) and incubated at room temperature for 30 min. After 3-times washing with PBS,
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Fig. 3. Colon carcinoma cells adhere to aorta patches. (A) HCT-116 cells expressing eGFP were seeded to freshly prepared aorta patches and their eGFP fluorescence was monitored by fluorescence microscopy 24 h after seeding. eGFP-fluorescence of HCT-116 cells (shown in green) was separated from auto-fluorescence of fibroblasts (shown in red) at confocal microscope Olympus SP5. (B) Comparison of cell–cell contacts (anti-E-cadherin) and cell–matrix contacts (anti-integrin beta1) of HCT-116 cells on the aorta patches (left panels) and on collagen-coated chamber slides (right panels). Bars: 10 m. (For interpretation of reference to color in this figure legend, the reader is referred to the web version of this article.)
the aorta pieces were transferred to glass slides (inside up) and mounted with Fluoromount-G.
2.4. Cell viability assay For the Alamar blue cell viability assay, the isolated aorta patches were incubated up to 3 days. Cell medium alone served as negative control, from which the background fluorescence was subtracted. Measurements were performed after incubation of the patches for 24, 48 and 72 h at 37 ◦ C in a humidified atmosphere. Fluorescence based absorption was measured at 540 nm on a micro plate reader (Tekan, Switzerland).
2.5. Generation of viral particles and eGFP transduction of HCT-116 cells Viral particles were produced as cell-free supernatants by transient transfection of the packaging cell line HEK-293T as described (Weber et al., 2008, 2012). In brief, the lentiviral vector LeGO-iG2Luc2 expressing eGFP-cDNA (Firefly luciferase, Promega) was packaged using the third-generation packaging plasmids pMDLg/pRRE (Addgene.org #12251), pRSV-Rev (Addgene.org #12253) and phCMV-VSV-G expressing the envelope protein of vesicular stomatitis virus. The supernatant was harvested 24 h after transfection, 0.45 m filtrated and stored at −80 ◦ C. Titration on 293T cells using 8 g/ml polybrene and spin-inoculation (1000 × g, 1 h, 25 ◦ C) resulted in a functional titer of 5.2 × 107 TU per ml. HCT-116 target
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Fig. 4. Spreading of HCT-116 cells on aorta patches. HCT-116 cells were seeded to aorta patches, fixed after 2, 6 or 16 h (overnight) and stained with Alexa Fluor® 568-coupled phalloidin. The right panels show higher magnifications. Bars: 10 m.
cells were plated at 5 × 104 cells in 0.5 ml medium in each well of a 24 well plate. After addition of 30 l of viral particle containing supernatant to the cells, the medium was replaced the next day. eGFP-ITPKA transduction of the patches was performed with the retroviral vector MigRI as described in Windhorst et al. (2010). 2.6. Microscopy For analysis of eGFP-fluorescence by fluorescence microscopy, the patch was carefully transferred into a chamber slide and cells were fixed with 4% paraformaldehyde. All the light microscopy images were acquired with a Leica TCS SP5 confocal microscope permitting the acquisition of accurate three-dimensional data set. Dual channel confocal imaging was performed. In channel number one eGFP positive cell were imaged using a 488 nm Argon laser
line as excitation source and a novel state of the art GaAsP hybrid detection system Leica HyDTM set to collect all the light coming with the 500–520 nm spectral band. In this channel both specific eGFP signals and unspecific auto fluorescence signals can be differentiated however as eGFP specific signals are one order of magnitude stronger than auto fluorescence, which made it possible to clearly the identify eGFP-expressing cells. In the second channel auto fluorescence from all cells was excited by a Dipole Pumped Solid State DPSS 561 nm laser source and detected with a second state-ofthe-art GaAsP hybrid detection system Leica HyDTM set to collect all the light coming with the 565–747 nm spectral band. In this spectral band only auto fluorescence is detectable being the eGFP positive signal negligible. Auto fluorescence emission spectra upon excitation at 488 and 561 nm were acquired (data not shown) to characterize the optical properties of the system. For accurate 3D
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reconstruction images in 246 × 246 × (20–50) m3 volume were acquired using a pixel size of 240 nm (1024 × 1024) pixel images and a voxel height either of 500 nm or of 1 m. The pixel size of (0.240 × 0.240 × (0.500–1)) m3 is sufficiently accurate to resolve cellular structures. The acquired image stacks were processed by mean of Bitplane Imaris software that allowed 3D reconstruction and accurate image visualization after automatic segmentation routine. 3. Results and discussion 3.1. Establishment of an adhesion assay using aortic patches as substrate To develop a cell adhesion assay that is feasible for routine analyses and that shows all the benefits of intact tissue, we have chosen in situ mouse aortic endothelium as a substrate. Mouse aorta is a robust vessel of suitable size, it is readily available and easy to prepare. Isolated thoracic mouse aortas were longitudinally cut and pinned inside out onto petri dishes that had been coated with a silicone elastomer. Next, a metal ring was placed to enclose the patch and defined numbers (5 × 104 ) of tumor cells were then seeded onto the patch (Fig. 1). In order to prevent de-differentiation of vessel cells, we did not cultivate the patches longer than three days. In conclusion, we established a vessel model that is easily to prepare and stable for three days. 3.2. Validation of cellular integrity of aortic tissue patches In order to analyze if the different cell layers of the aortic patch remained intact after preparation, the tissue cells were transfected with a retroviral vector encoding for eGFP to label the cells. 24 h after treating the tissue with the virus, the patches were fixed and analyzed by fluorescence microscopy. Thereby, we first focused on the outer cell layer containing the endothelial cells, and thereafter on the subendothelial layer containing fibroblasts and collagen fibers. As shown in Fig. 2A the transfection efficiency of tissue cells was very high, and thus the endothelial were well visible at the outer cell layer (left panel). Verification was performed by staining the cells with the endothelial marker protein CD31 (middle panel; Miettinen et al., 1994). The subendothelial layer containing fibroblasts and collagen fibers was detectable when focusing behind the endothelial cells (right panel). However, it was not possible to focus behind the subendothelial layer to detect smooth muscle cells. To show if the tissue cells start to dedifferentiate during cultivation, proliferation was analyzed by the Alamar blue Assay for 72 h, every 24 h. We found no proliferation within 72 h of incubation, but after 72 h cell viability decreased (Fig. 2B). Therefore, we recommend keeping the tissue only for 48 h in culture. In summary our data show that the aortic tissue cells do not dedifferentiate and remain stable for at least 48 h. Furthermore, the endothelial and subendothelial cell layers remain intact after preparation. Thus, the ex vivo model should be suitable to analyze interaction of blood cells (circulating tumor cells, immune cells and plates) with endothelial and with subendothelial cells. In addition, our model enables to genetically modify the vascular cells, thus provides the possibility to analyze the function of proteins of interest for endothelial–blood cell interaction. 3.3. Interaction of vascular cells with tumor cells Since the interaction of circulating tumor cells with endothelial and subendothelial cells is one essential factor that determines if the tumor cells adhere to the vascular wall or not, we analyzed if our
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Table 1 Spreading of HCT-116 cells on aorta patches. HCT-116 cells were seeded on aorta patches and filopodia length and cell area were measured 2 h, 6 h and 16 h after seeding. Values obtained from cells incubated for 16 h were set to 100%. Measurements were done at least twice, at least 80 cells were analyzed per measurement. Time (h)
Filopodia length, %
Cell area, %
2 6 16
Not detectable 88 ± 6 100
53 ± 9 65 ± 17 100
model is suitable to observe tumor cell–endothelial/subendothelial interactions. The prerequisite for such an analysis is that suspended tumor cells adhere to the isolated aortic tissue. Therefore, we first tested adherence of the established colon cancer cell line HCT-116 on aortic patches (Lin et al., 2014). Suspended eGFP-expressing HCT-116 cells were seeded on freshly isolated tissue and one day after incubation, the cells were fixed with paraformaldehyde and embedded in mountain-medium. We found that one day after seeding the cells had adhered to the tissue (Fig. 3A), the apparently visible invasion in Fig. 3A (lower panel – Z-axis) results from folding of the tissue. To analyze potential differences of HCT-116 cell adhesion when seeded on collagen-coated chamber slides or on aorta patches, we stained the cells for E-cadherin to analyze cell–cell contacts. Cell–matrix contacts were examined by staining the cells for integrin-, the main protein of focal adhesions in motile cells (Ivaska et al., 2002). Our data show that cell–cell interactions are similar when cells were seeded on collagen or on vascular tissue (Fig. 3B). Under both conditions, the cells grow as tight clusters. However, the interaction of tumor cells with the matrix seemed to be different. When seeded on collagen-coated chamber slides, the single cells formed multiple focal adhesions that were equally distributed over the cells and additionally, they form large and very distinct focal adhesions at the periphery of the clusters. These additional focal adhesions were not formed when the cells were seeded on vascular tissue. In conclusion, our results reveal that suspended epithelial tumor cells adhere to our vascular patch tissue and show that cell-to-matrix interactions are different when the cells grow on collagen-covered chamber slides or on vascular tissue. 3.4. Spreading of tumor cells on vascular tissue Next, we examined if it is possible to monitor the process of tumor cell spreading (cell flattening and formation of cellular protrusions (Nürnberg et al., 2011) on the vascular patch tissue. Therefore, cells were fixed 2 h, 6 h and 16 h after seeding, and for labeling F-actin the cells were stained with Alexa Fluor® 568 phalloidin. Analysis by fluorescence microscopy revealed that 2 h after seeding most cells were round-shaped and did not form filopodia. After 6 h, most of the cells were still rounded up, but cells had formed filopodia. When the cells were incubated for 16 h, they flattened onto the substratum. Although filopodia-length did not significantly increase in comparison to cells cultivated for six hours, cells started to form lamellipodia (Fig. 4, Table 1). In conclusion, these data show that our ex vivo model provides the possibility to precisely analyze morphological changes during epithelial cell spreading on the aortic endothelial cell layer. In summary, our ex vivo model provides the possibility to analyze interaction of blood cells with endothelial and subendothelial cells. The main advantage of this model is that, in contrast to cultivated endothelial cells, interaction of blood cells with vascular tissue can be examined. In addition, our model provides the possibility to genetically modify both, the cells of the vascular tissue and the blood cells, enabling to analyze the impact of proteins on vascular wall–blood cell interactions. We demonstrated that suspended
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tumor cells adhere to the vascular tissue very well, and we show that it is possible to analyze the process of cell spreading. Since the mechanism of tumor cell spreading on vascular cells has not been elucidated in detail yet, our model now provides the possibility to analyze the impact of proteins involved in this process. Thus, our ex vivo model provides a feasible alternative for analyzing vascular wall–blood cell interactions of cultivated endothelial cells under static conditions. Acknowledgments We thank Christine Blechner for excellent technical support. This work was supported by a UCCH grant (University Cancer Center Hamburg). References Bachetti, T., Morbidelli, L., 2000. Endothelial cells in culture: a model for studying vascular functions. Pharmacol. Res. 42 (1), 9–19. Bouïs, D., Hospers, G.A., Meijer, C., et al., 2001. Endothelium in vitro: a review of human vascular endothelial cell lines for blood vessel-related research. Angiogenesis 4 (2), 91–102. Cines, D.B., Pollak, E.S., Buck, C.A., et al., 1998. Endothelial cells in physiology and in the pathophysiology of vascular disorders. Blood 91 (10), 3527–3561. Erusalimsky, J.D., 2009. Vascular endothelial senescence: from mechanisms to pathophysiology. J. Appl. Physiol. 106, 326–332. Gallagher, G.L., Sumpio, B.E., 1997. In: Kisco, Mt. (Ed.), Endothelial Cells. Futura Publishing Company, New York, pp. 151–186. Gimbrone, M.A., Cotran, R.S., Folkman, J., 1974. Human vascular endothelial cells in culture. Growth and DNA synthesis. J. Cell Biol. 60, 673.
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