Med Mol Morphol DOI 10.1007/s00795-013-0063-7
ORIGINAL PAPER
Observation of collagen fibrils produced by osteosarcoma cells using atomic force microscopy Osamu Hoshi
Received: 11 June 2013 / Accepted: 22 October 2013 Ó The Japanese Society for Clinical Molecular Morphology 2013
Abstract The present study examined the three-dimensional process of collagen fibril formation in the human osteosarcoma cell line NOS-1 by conventional scanning electron microscopy (SEM) and atomic force microscopy (AFM). SEM images showed collagen fibril formation on the bottom of culture dishes after 1 week of culture. The collagen fibrils had diameters of 30–100 nm. The surfaces of individual fibrils had characteristic grooves and ridges with periodicities of 60–70 nm. AFM images showed that the newly formed collagen fibrils were 30–300 nm in diameter and possessed characteristic grooves and ridges with periodicities of 60–70 nm. The thicker collagen fibrils contained thinner (approximately 30 nm thick) subfibrils that ran in a helical direction along the long axis of the thicker fibrils. Furthermore, twisted structures of collagen fibrils, which possessed a characteristic rope-like structure, were also identified. The ultrastructure of the collagen fibrils was clearly imaged in liquid medium by AFM, and the process of collagen fibril assembly was successfully analyzed under conditions much closer to the physiological state than those afforded by transmission electron microscopy or SEM. AFM also provided a precise morphological measurement, particularly of the vertical distance, of collagen fibrils with nanometer-scale resolution in liquid conditions. Keywords Collagen fibril assembly Osteosarcoma Cell culture Atomic force microscopy Scanning electron microscopy
O. Hoshi (&) Anatomy and Physiological Science, Graduate School of Health Care Science, Tokyo Medical and Dental University, Tokyo 113-8519, Japan e-mail: [email protected]
Introduction Collagen fibrils are a major protein component of the extracellular matrix in animals and are involved in various important biological functions. Collagens have received interest because of their involvement in not only tissue structuring and cell attachment [1] but also various human diseases [2]. Furthermore, collagen matrices are used as platforms for cell biological and tissue engineering applications [3]. Thus, understanding the mechanisms of collagen fibril formation is of significance for both biological and biotechnological research. As collagen fibrils are assemblies of collagen molecules, many researchers have been interested in observing the assembly process by transmission electron microscopy (TEM) [4, 5]. Previous TEM studies indicated that long and thick collagen fibrils are formed by gathering of short and thin collagen fibrils [6, 7]. Scanning electron microscopy (SEM), particularly in-lens field emission SEM, has also been used to examine the ultrastructure of collagen fibrils with high resolution [8]. However, the detailed structure of collagen fibrils and mechanisms of fibril formation are poorly understood. Atomic force microscopy (AFM), developed in 1986 [9], has an advantage over electron microscopy in that it allows the investigation of biological samples without metal coating or conductive treatment in a non-vacuous (i.e., air or liquid) environment at high resolution, thereby preserving the physiological conformations and functions [10]. AFM also has the potential for imaging biological specimens at higher resolution than TEM or SEM. Consequently, collagen assembly has been studied in several biological systems using AFM [11–16]. Collagen fibril formation has been mainly studied by observation of the growth process of collagen self-assembly
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[7, 11]. In vivo, tendon collagen fibrils have been studied using rotator shadowing or reconstruction methods [17, 18]. However, these studies only showed the structures of a small number of collagen fibrils in a restricted region. In the present study, we attempted to provide methodological insight into the three-dimensional process of collagen fibril formation in NOS-1 human osteosarcoma cells through application of AFM under liquid conditions.
Materials and methods Cell culture The human osteosarcoma cell line NOS-1 was used in this study. This cell line was established by Hotta et al. [19] and is known to produce large amounts of type I collagen fibrils in culture dishes [19, 20]. NOS-1 cells (3 9 105) were incubated in 35-mm plastic dishes (Iwaki Glass, Tokyo, Japan) and maintained in 2 ml of growth medium (Media-I; Immuno-Biological Laboratories, Fujioka, Japan) with 5 % fetal bovine serum (Mitsubishi Kasei, Tokyo, Japan) and 50 lM ascorbic acid [21, 22] at 37 °C in a 5 % CO2 humidified incubator. After 1 week of culture, the cells and their products were prepared for SEM and AFM observation. SEM After a brief rinse in 0.05 M phosphate buffer (pH 7.4), the cells and collagen fibrils were fixed with 1 % osmium tetroxide in the same buffer for 3 h at 4 °C. They were then rinsed again with buffer, conductively stained with 0.5 % tannic acid for 10 min and 1 % osmium tetroxide for 30 min, and dehydrated using a graded ethanol series. The fixation, rinsing, conductive staining, and dehydration were Fig. 1 SEM images of NOS-1 human osteosarcoma cells and collagen fibrils on a culture dish after 1 week of culture. a NOS1 cells scattered on the dish. The cells appear flattened or spherical. Collagen fibrils are seen around some of the NOS-1 cells (arrow). b Highmagnification view of NOS-1 cells with newly formed collagen fibrils. c Highmagnification view of collagen fibrils. The individual fibrils are 30–100 nm in diameter and have grooves and ridges with a periodicity of approximately 70 nm. The periodic grooves and ridges are synchronized with adjacent fibrils
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performed by gently changing the solutions using a micropipette. Immediately after the removal of 100 % ethanol, the dishes were directly placed in a critical point dryer (HCP-2; Hitachi, Tokyo, Japan) and dried in liquid CO2. The bottoms of the dried dishes were cut into small 2-mm2 sections using scissors and affixed to a small metal plate with silver paste. The squares were then coated with platinum–palladium (Pt–Pd) for 45 s in a magnetic ion coater (E-1030; Hitachi) with a distance of 30 mm between the sample and the Pt–Pd target under 5 Pa of argon. The coated samples were observed using in-lens field emission SEM (S-5000; Hitachi) under an accelerating voltage of 5 kV. AFM After a brief rinse in 0.05 M phosphate buffer (pH 7.4), the cells and collagen fibrils were fixed with 1 % glutaraldehyde and treated with 1 % osmium tetroxide in the same buffer for 3 h at 4 °C. A commercial AFM (SPI 3700 with a 4000 probe station; SII NanoTechnology, Chiba, Japan) was used. Because the apparatus was equipped with a light microscope above the microscope unit, the observed NOS-1 cells could be accurately determined in AFM by comparison with phase-contrast micrographs taken earlier. For imaging, AFM was operated in the dynamic force (i.e., intermittent contact) mode in phosphate-buffered saline (PBS) solution. The maximum scan range of the piezo scanner used was approximately 20 lm in width (x–y) and 1.2 lm in height (z). Reduction in the oscillation amplitude was used as a feedback parameter by the slope detection method. Commercially available V-shaped silicon nitride cantilevers with oxide-sharpened tips and a nominal spring constant of 0.32 N/m (SNL-10; Veeco Inc., Santa Barbara,
Med Mol Morphol
CA, USA) were used for imaging, and the normal resonance frequency was approximately 12.5 kHz in the liquid medium. The dimensions of the images obtained were 512 9 512 pixels. The height mode images were displayed by a computer in the ‘‘gradation mode,’’ which presents the height of specimens as color gradients.
Results
directions and formed a network-like structure. Characteristic grooves and ridges with periodicities of 60–70 nm on the surfaces of the collagen fibrils were confirmed in the AFM images (Figs. 2b, 3). The depth of grooves with periodic patterns of 60–70 nm along the long axis of the collagen fibrils was approximately 15 nm (Fig. 3). Thinner collagen fibrils (approximately 30 nm) were often observed entwined in a thicker bundle, which appeared to a thicker collagen fibril. The thinner fibrils that formed the thicker fibrils often ran in a helical direction relative to the long axis of the thicker fibrils (Figs. 2b, 3). Some thicker fibrils were also periodically constricted because of the presence of shallow, oblique grooves (Fig. 4). The vertical distance from the top to the bottom of the shallow oblique grooves was approximately 30 nm, and the helical angle of the groove was 60°.
Under SEM, NOS-1 cells appeared flattened or spherical and were present in groups or singly (Fig. 1a). Some cells had formed new bundles of fibrils around themselves after 1 week of culture (Fig. 1b). The diameters of these fibrils were 30–100 nm (Fig. 1c). At high magnification, the surfaces of the individual fibrils possessed characteristic grooves and ridges with periodicities of 60–70 nm (Fig. 1c), indicating that they were collagen fibrils. Neighboring fibrils often ran parallel to one another or were fused together, with their periodic grooves and ridges synchronized with adjacent fibrils. A tapering end was occasionally observed in some fibrils. The developing collagen fibrils from NOS-1 cells were observed using AFM in the dynamic mode in PBS solution (Fig. 2a, b). The AFM images showed that the diameters of the collagen fibrils varied from 30 to 300 nm (n = 55 collagen fibrils). The average diameter was 145.0 nm with a standard deviation of 78.0 nm. The fibrils ran in various
The present study revealed, for the first time, the process of new collagen fibril formation in cultured NOS-1 human osteosarcoma cells using AFM in a liquid environment. In a previous study, the process of collagen fibril formation in NOS-1 cells was investigated using high-resolution SEM [23]. Short and thin collagen fibrils (approximately 1 lm long and 30 nm thick) were observed twisting with one another at tapered ends in a right helical
Fig. 2 a AFM image of collagen fibrils in phosphate-buffered saline solution. b The diameters of the collagen fibrils vary from 30 to 200 nm. The fibrils run in various directions and cross one another or entwine together to form a network-like structure on the dish. Note that some of the thinner fibrils appear to form thicker bundles. The
surfaces of the collagen fibrils show characteristic grooves and ridges with periodicities of 60–70 nm. Note the presence of some thinner collagen fibrils (arrows) on the surface of the thicker fibril. These thinner fibrils run in a right helical direction relative to the long axis of the thicker collagen fibril
Discussion
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Fig. 3 High-magnification view of a thicker collagen fibril. Characteristic grooves and ridges with periodicities of 60–70 nm are clearly observed on the surface of the collagen fibril. This fibril appears to contain thinner collagen fibrils running in a left helical direction relative to the long axis of the fibril
direction with synchronized periodicity, thereby forming longer and thicker fibrils. The present AFM observations indicate that the thicker collagen fibrils contained thinner subfibrils (approximately 30 nm thick) that ran in right (Fig. 2b) or left (Fig. 3) helical directions along the long axis of the thicker fibrils. The periodic grooves of the thinner fibrils almost corresponded to those of thicker fibrils (Fig. 3). These findings indicate that thinner collagen subfibrils twist around thicker fibrils, leading to the formation of even thicker fibrils (Fig. 5). However, the direction in which thinner collagen subfibrils twist around the thicker collagen fibrils remains uncertain and requires further detailed studies. The constriction of collagen fibrils with oblique grooves at regular intervals was confirmed by the AFM images. This conformation implies local unwinding of the collagen fibrils, a phenomenon previously reported from the observation of tendon collagen fibrils using AFM [24]. In that study, the authors showed data supporting a twisted structure of collagen fibrils similar to the appearance of a classical rope. The present data also support an arrangement characteristic of a rope-like structure. The reason why the helical angle of the shallow obliquely running grooves was 60° remains unclear. It may reflect local unwinding of collagen fibrils as reported by Bozec et al. [24]. Taken together, two points were proposed as the possible mechanisms of collagen formation; one is the twisting of thinner collagen subfibrils
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Fig. 4 AFM image of a slightly constricted collagen fibril with oblique grooves. Note that the grooves (arrows) run obliquely relative to the long axis of the collagen fibril at approximately regular intervals
around thicker fibrils, while the other is the formation of a rope-like structure, i.e., the spiral formation of collagen fibrils. In the present study, fixed cells were observed using AFM in a liquid environment. If the process of collagen fibril formation by a living cell could be imaged using time-lapse AFM, it has the potential to elucidate the mechanisms underlying collagen formation in a more physiological state. In addition, the tumor cell line used for this investigation of collagen fibril formation may have a different mode of fibril formation compared with that in the physiological state, such as that seen in normal human fibroblast cells. In the preliminary experiment, human fibroblasts were cultured in the almost identical conditions to NOS-1 cells, and the bottom of the culture dishes was observed by SEM. However, collagen fibrils could not be identified to the same extent as in NOS-1 cells. Thus, NOS1 cells were chosen for this study. With regard to fibroblasts, the nature of the collagen synthesized and the influence of L-ascorbic acid 2-phosphate on collagen formation have been previously reported in detail [25–27]. The effects of pH on collagen assembly were also studied in embryonic chick corneal epithelium [28]. The new mechanism for fibril formation suggested for NOS-1 cells should be further studied in relation to the nature of collagen, L-ascorbic acid 2-phosphate and the pH of the medium.
Med Mol Morphol Fig. 5 Schematic representation of a collagen fibril produced by NOS-1 cells. The collagen fibril is formed by the assembly of thin collagen fibrils, which are arranged in a helical direction relative to the long axis. The periodic grooves and ridges of thin fibers are synchronized with adjacent fibrils. Bar 30 nm
Conclusion The present study investigated collagen fibril formation in a cell culture system of NOS-1 human osteosarcoma cells using SEM and AFM. The newly formed collagen fibrils from the osteosarcoma cells were successfully observed using AFM in a liquid environment. Thus, the threedimensional structure of wet collagen fibrils was precisely analyzed in a condition closer to the physiological state than that afforded by SEM. Acknowledgments The author is very grateful to Professor T. Ushiki (Niigata University Graduate School of Medical and Dental Sciences) for his continuous support and encouragement. This study was supported in part by a Grant-in-Aid for Scientific Research (No. 20590186 to O.H.) from the Japan Society for the Promotion of Science. Conflict of interest The author reports no conflicts of interest. The author alone is responsible for the content and writing of the paper.
References 1. Grinnell F (2003) Fibroblast biology in three-dimensional collagen matrices. Trends Cell Biol 13:264–269 2. Kadler KE (1993) Learning how mutations in type I collagen genes cause connective tissue disease. Int J Exp Pathol 74:319–323 3. Guidry C, Grinnell F (1985) Studies on the mechanism of hydrated collagen gel reorganization by human skin fibroblast. J Cell Sci 79:67–81 4. Marchini M, Morocutti A, Ruggeri MH, Koch MHJ, Bigi A, Roveri N (1986) Differences in the fibril structure of corneal and tendon collagen. An electron microscopy and X-ray diffraction investigation. Connect Tissue Res 15:269–281 5. Raspanti M, Ottani V, Ruggeri A (1990) Subfibrillar architecture and functional properties of collagen: a comparative study in rat tendons. J Anat 172:157–164 6. Birk DE, Nurminskaya MV, Zycband EI (1995) Collagen fibrillogenesis in situ: fibril segments undergo post-depositional modifications resulting in linear and lateral growth during matrix development. Dev Dyn 202:229–243 7. Kadler KE, Holmes DF, Trotter JA, Chapman JA (1996) Collagen fibril formation. Biochem J 316:1–11 8. Raspanti M, Alessandrini A, Gobbi P, Ruggeri A (1996) Collagen fibril surface: TMAFM, FEG-SEM and freeze-etching observations. Microsc Res Tech 35:87–93
9. Binnig G, Quate CF, Gerber C (1986) Atomic force microscope. Phys Rev Lett 56:930–933 10. Drake B, Prater CB, Weisenhorn AL, Gould SAC, Albrecht TR, Quate CF, Cannel DS, Hansma HG, Hansma PK (1989) Imaging crystals, polymers, and processes in water with the atomic force microscope. Science 243:1586–1589 11. Cisneros DA, Hung C, Franz CM, Muller DJ (2006) Observing growth steps of collagen self-assembly by time-lapse high-resolution atomic force microscopy. J Struct Biol 154:232–245 12. Franz CM, Muller DJ (2011) Studying collagen self-assembly by time-lapse high-resolution atomic force microscopy. Methods Mol Biol 736:97–107 13. Liang Z, Zhou C, Zeng R, Cai H, Guo Z (2010) Visualization of the nanoscale assembly of type I collagen on PLA by AFM. Scanning 32:104–111 14. Raspanti M, Protasoni M, Manelli M, Guizzardi S, Mantovani V, Sala A (2006) The extracellular matrix of the human aortic wall: ultrastructural observations by FEG-SEM and by tapping-mode AFM. Micron 37:81–86 15. Tilley JMR, Carr AJ, Czernuszka JT (2011) Atomic Force Microscopy of bulk tendon samples: affect of location and fixation on tissue ultrastructure. Micron 42:531–535 16. Yamamoto S, Hitomi J, Shigeno M, Sawaguchi S, Abe H, Ushiki T (1997) Atomic force microscopic studies of isolated collagen fibrils of the bovine cornea and sclera. Arch Histol Cytol 60:371–378 17. Birk DE, Zycband EI, Winkelmann DA, Trelstad RL (1989) Collagen fibrillogenesis in situ: fibril segments are intermediates in matrix assembly. Proc Natl Acad Sci USA 86:4549–4553 18. Birk DE, Zycband EI, Woodruff S, Winkelmann DA, Trelstad RL (1997) Collagen fibrillogenesis in situ: fibril segments become long fibrils as the developing tendon matures. Dev Dyn 208:291–298 19. Hotta T, Motoyama T, Watanabe H (1992) Three human osteosarcoma cell lines exhibiting different phenotypic expressions. Acta Pathol Jpn 42:595–603 20. Ogose A, Motoyama T, Hotta T, Waranabe H, Takahashi HE (1995) Bone formation in vitro and in nude mice by human osteosarcoma cells. Virchows Arch 426:117–125 21. Grinnell F, Fukamizu H, Pawelek P, Nakagawa S (1989) Collagen processing, crosslinking, and fibril bundle assembly in matrix produced by fibroblasts in long-term cultures supplemented with ascorbic acid. Exp Cell Res 181:483–491 22. Lamande SR, Barteman JF (1993) A mouse 3T6 fibroblast cell culture model for the study of normal and protein-engineered collagen synthesis and deposition into the extracellular matrix. Matrix 13:323–330 23. Hashizume H, Hitomi J, Ushiki T (1999) Growth of collagen fibrils produced by human osteosarcoma cells: high-resolution scanning electron microscopy. Arch Histol Cytol 62:327–335 24. Bozec L, van der Heijden G, Horton M (2007) Collagen fibrils: nanoscale ropes. Biophys J 92:70–75
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27. Hata R, Senoo H (1989) L-ascorbic acid 2-phosphate stimulates collagen accumulation, cell proliferation, and formation of a three-dimensional tissue-like substance by skin fibroblasts. J Cell Physiol 138:8–16 28. Bard JB, Hulmes DJ, Purdom IF, Ross AS (1993) Chick corneal development in vitro: diverse effects of pH on collagen assembly. J Cell Sci 105:1045–1055
ORIGINAL PAPER
Observation of collagen fibrils produced by osteosarcoma cells using atomic force microscopy Osamu Hoshi
Received: 11 June 2013 / Accepted: 22 October 2013 Ó The Japanese Society for Clinical Molecular Morphology 2013
Abstract The present study examined the three-dimensional process of collagen fibril formation in the human osteosarcoma cell line NOS-1 by conventional scanning electron microscopy (SEM) and atomic force microscopy (AFM). SEM images showed collagen fibril formation on the bottom of culture dishes after 1 week of culture. The collagen fibrils had diameters of 30–100 nm. The surfaces of individual fibrils had characteristic grooves and ridges with periodicities of 60–70 nm. AFM images showed that the newly formed collagen fibrils were 30–300 nm in diameter and possessed characteristic grooves and ridges with periodicities of 60–70 nm. The thicker collagen fibrils contained thinner (approximately 30 nm thick) subfibrils that ran in a helical direction along the long axis of the thicker fibrils. Furthermore, twisted structures of collagen fibrils, which possessed a characteristic rope-like structure, were also identified. The ultrastructure of the collagen fibrils was clearly imaged in liquid medium by AFM, and the process of collagen fibril assembly was successfully analyzed under conditions much closer to the physiological state than those afforded by transmission electron microscopy or SEM. AFM also provided a precise morphological measurement, particularly of the vertical distance, of collagen fibrils with nanometer-scale resolution in liquid conditions. Keywords Collagen fibril assembly Osteosarcoma Cell culture Atomic force microscopy Scanning electron microscopy
O. Hoshi (&) Anatomy and Physiological Science, Graduate School of Health Care Science, Tokyo Medical and Dental University, Tokyo 113-8519, Japan e-mail: [email protected]
Introduction Collagen fibrils are a major protein component of the extracellular matrix in animals and are involved in various important biological functions. Collagens have received interest because of their involvement in not only tissue structuring and cell attachment [1] but also various human diseases [2]. Furthermore, collagen matrices are used as platforms for cell biological and tissue engineering applications [3]. Thus, understanding the mechanisms of collagen fibril formation is of significance for both biological and biotechnological research. As collagen fibrils are assemblies of collagen molecules, many researchers have been interested in observing the assembly process by transmission electron microscopy (TEM) [4, 5]. Previous TEM studies indicated that long and thick collagen fibrils are formed by gathering of short and thin collagen fibrils [6, 7]. Scanning electron microscopy (SEM), particularly in-lens field emission SEM, has also been used to examine the ultrastructure of collagen fibrils with high resolution [8]. However, the detailed structure of collagen fibrils and mechanisms of fibril formation are poorly understood. Atomic force microscopy (AFM), developed in 1986 [9], has an advantage over electron microscopy in that it allows the investigation of biological samples without metal coating or conductive treatment in a non-vacuous (i.e., air or liquid) environment at high resolution, thereby preserving the physiological conformations and functions [10]. AFM also has the potential for imaging biological specimens at higher resolution than TEM or SEM. Consequently, collagen assembly has been studied in several biological systems using AFM [11–16]. Collagen fibril formation has been mainly studied by observation of the growth process of collagen self-assembly
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[7, 11]. In vivo, tendon collagen fibrils have been studied using rotator shadowing or reconstruction methods [17, 18]. However, these studies only showed the structures of a small number of collagen fibrils in a restricted region. In the present study, we attempted to provide methodological insight into the three-dimensional process of collagen fibril formation in NOS-1 human osteosarcoma cells through application of AFM under liquid conditions.
Materials and methods Cell culture The human osteosarcoma cell line NOS-1 was used in this study. This cell line was established by Hotta et al. [19] and is known to produce large amounts of type I collagen fibrils in culture dishes [19, 20]. NOS-1 cells (3 9 105) were incubated in 35-mm plastic dishes (Iwaki Glass, Tokyo, Japan) and maintained in 2 ml of growth medium (Media-I; Immuno-Biological Laboratories, Fujioka, Japan) with 5 % fetal bovine serum (Mitsubishi Kasei, Tokyo, Japan) and 50 lM ascorbic acid [21, 22] at 37 °C in a 5 % CO2 humidified incubator. After 1 week of culture, the cells and their products were prepared for SEM and AFM observation. SEM After a brief rinse in 0.05 M phosphate buffer (pH 7.4), the cells and collagen fibrils were fixed with 1 % osmium tetroxide in the same buffer for 3 h at 4 °C. They were then rinsed again with buffer, conductively stained with 0.5 % tannic acid for 10 min and 1 % osmium tetroxide for 30 min, and dehydrated using a graded ethanol series. The fixation, rinsing, conductive staining, and dehydration were Fig. 1 SEM images of NOS-1 human osteosarcoma cells and collagen fibrils on a culture dish after 1 week of culture. a NOS1 cells scattered on the dish. The cells appear flattened or spherical. Collagen fibrils are seen around some of the NOS-1 cells (arrow). b Highmagnification view of NOS-1 cells with newly formed collagen fibrils. c Highmagnification view of collagen fibrils. The individual fibrils are 30–100 nm in diameter and have grooves and ridges with a periodicity of approximately 70 nm. The periodic grooves and ridges are synchronized with adjacent fibrils
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performed by gently changing the solutions using a micropipette. Immediately after the removal of 100 % ethanol, the dishes were directly placed in a critical point dryer (HCP-2; Hitachi, Tokyo, Japan) and dried in liquid CO2. The bottoms of the dried dishes were cut into small 2-mm2 sections using scissors and affixed to a small metal plate with silver paste. The squares were then coated with platinum–palladium (Pt–Pd) for 45 s in a magnetic ion coater (E-1030; Hitachi) with a distance of 30 mm between the sample and the Pt–Pd target under 5 Pa of argon. The coated samples were observed using in-lens field emission SEM (S-5000; Hitachi) under an accelerating voltage of 5 kV. AFM After a brief rinse in 0.05 M phosphate buffer (pH 7.4), the cells and collagen fibrils were fixed with 1 % glutaraldehyde and treated with 1 % osmium tetroxide in the same buffer for 3 h at 4 °C. A commercial AFM (SPI 3700 with a 4000 probe station; SII NanoTechnology, Chiba, Japan) was used. Because the apparatus was equipped with a light microscope above the microscope unit, the observed NOS-1 cells could be accurately determined in AFM by comparison with phase-contrast micrographs taken earlier. For imaging, AFM was operated in the dynamic force (i.e., intermittent contact) mode in phosphate-buffered saline (PBS) solution. The maximum scan range of the piezo scanner used was approximately 20 lm in width (x–y) and 1.2 lm in height (z). Reduction in the oscillation amplitude was used as a feedback parameter by the slope detection method. Commercially available V-shaped silicon nitride cantilevers with oxide-sharpened tips and a nominal spring constant of 0.32 N/m (SNL-10; Veeco Inc., Santa Barbara,
Med Mol Morphol
CA, USA) were used for imaging, and the normal resonance frequency was approximately 12.5 kHz in the liquid medium. The dimensions of the images obtained were 512 9 512 pixels. The height mode images were displayed by a computer in the ‘‘gradation mode,’’ which presents the height of specimens as color gradients.
Results
directions and formed a network-like structure. Characteristic grooves and ridges with periodicities of 60–70 nm on the surfaces of the collagen fibrils were confirmed in the AFM images (Figs. 2b, 3). The depth of grooves with periodic patterns of 60–70 nm along the long axis of the collagen fibrils was approximately 15 nm (Fig. 3). Thinner collagen fibrils (approximately 30 nm) were often observed entwined in a thicker bundle, which appeared to a thicker collagen fibril. The thinner fibrils that formed the thicker fibrils often ran in a helical direction relative to the long axis of the thicker fibrils (Figs. 2b, 3). Some thicker fibrils were also periodically constricted because of the presence of shallow, oblique grooves (Fig. 4). The vertical distance from the top to the bottom of the shallow oblique grooves was approximately 30 nm, and the helical angle of the groove was 60°.
Under SEM, NOS-1 cells appeared flattened or spherical and were present in groups or singly (Fig. 1a). Some cells had formed new bundles of fibrils around themselves after 1 week of culture (Fig. 1b). The diameters of these fibrils were 30–100 nm (Fig. 1c). At high magnification, the surfaces of the individual fibrils possessed characteristic grooves and ridges with periodicities of 60–70 nm (Fig. 1c), indicating that they were collagen fibrils. Neighboring fibrils often ran parallel to one another or were fused together, with their periodic grooves and ridges synchronized with adjacent fibrils. A tapering end was occasionally observed in some fibrils. The developing collagen fibrils from NOS-1 cells were observed using AFM in the dynamic mode in PBS solution (Fig. 2a, b). The AFM images showed that the diameters of the collagen fibrils varied from 30 to 300 nm (n = 55 collagen fibrils). The average diameter was 145.0 nm with a standard deviation of 78.0 nm. The fibrils ran in various
The present study revealed, for the first time, the process of new collagen fibril formation in cultured NOS-1 human osteosarcoma cells using AFM in a liquid environment. In a previous study, the process of collagen fibril formation in NOS-1 cells was investigated using high-resolution SEM [23]. Short and thin collagen fibrils (approximately 1 lm long and 30 nm thick) were observed twisting with one another at tapered ends in a right helical
Fig. 2 a AFM image of collagen fibrils in phosphate-buffered saline solution. b The diameters of the collagen fibrils vary from 30 to 200 nm. The fibrils run in various directions and cross one another or entwine together to form a network-like structure on the dish. Note that some of the thinner fibrils appear to form thicker bundles. The
surfaces of the collagen fibrils show characteristic grooves and ridges with periodicities of 60–70 nm. Note the presence of some thinner collagen fibrils (arrows) on the surface of the thicker fibril. These thinner fibrils run in a right helical direction relative to the long axis of the thicker collagen fibril
Discussion
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Fig. 3 High-magnification view of a thicker collagen fibril. Characteristic grooves and ridges with periodicities of 60–70 nm are clearly observed on the surface of the collagen fibril. This fibril appears to contain thinner collagen fibrils running in a left helical direction relative to the long axis of the fibril
direction with synchronized periodicity, thereby forming longer and thicker fibrils. The present AFM observations indicate that the thicker collagen fibrils contained thinner subfibrils (approximately 30 nm thick) that ran in right (Fig. 2b) or left (Fig. 3) helical directions along the long axis of the thicker fibrils. The periodic grooves of the thinner fibrils almost corresponded to those of thicker fibrils (Fig. 3). These findings indicate that thinner collagen subfibrils twist around thicker fibrils, leading to the formation of even thicker fibrils (Fig. 5). However, the direction in which thinner collagen subfibrils twist around the thicker collagen fibrils remains uncertain and requires further detailed studies. The constriction of collagen fibrils with oblique grooves at regular intervals was confirmed by the AFM images. This conformation implies local unwinding of the collagen fibrils, a phenomenon previously reported from the observation of tendon collagen fibrils using AFM [24]. In that study, the authors showed data supporting a twisted structure of collagen fibrils similar to the appearance of a classical rope. The present data also support an arrangement characteristic of a rope-like structure. The reason why the helical angle of the shallow obliquely running grooves was 60° remains unclear. It may reflect local unwinding of collagen fibrils as reported by Bozec et al. [24]. Taken together, two points were proposed as the possible mechanisms of collagen formation; one is the twisting of thinner collagen subfibrils
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Fig. 4 AFM image of a slightly constricted collagen fibril with oblique grooves. Note that the grooves (arrows) run obliquely relative to the long axis of the collagen fibril at approximately regular intervals
around thicker fibrils, while the other is the formation of a rope-like structure, i.e., the spiral formation of collagen fibrils. In the present study, fixed cells were observed using AFM in a liquid environment. If the process of collagen fibril formation by a living cell could be imaged using time-lapse AFM, it has the potential to elucidate the mechanisms underlying collagen formation in a more physiological state. In addition, the tumor cell line used for this investigation of collagen fibril formation may have a different mode of fibril formation compared with that in the physiological state, such as that seen in normal human fibroblast cells. In the preliminary experiment, human fibroblasts were cultured in the almost identical conditions to NOS-1 cells, and the bottom of the culture dishes was observed by SEM. However, collagen fibrils could not be identified to the same extent as in NOS-1 cells. Thus, NOS1 cells were chosen for this study. With regard to fibroblasts, the nature of the collagen synthesized and the influence of L-ascorbic acid 2-phosphate on collagen formation have been previously reported in detail [25–27]. The effects of pH on collagen assembly were also studied in embryonic chick corneal epithelium [28]. The new mechanism for fibril formation suggested for NOS-1 cells should be further studied in relation to the nature of collagen, L-ascorbic acid 2-phosphate and the pH of the medium.
Med Mol Morphol Fig. 5 Schematic representation of a collagen fibril produced by NOS-1 cells. The collagen fibril is formed by the assembly of thin collagen fibrils, which are arranged in a helical direction relative to the long axis. The periodic grooves and ridges of thin fibers are synchronized with adjacent fibrils. Bar 30 nm
Conclusion The present study investigated collagen fibril formation in a cell culture system of NOS-1 human osteosarcoma cells using SEM and AFM. The newly formed collagen fibrils from the osteosarcoma cells were successfully observed using AFM in a liquid environment. Thus, the threedimensional structure of wet collagen fibrils was precisely analyzed in a condition closer to the physiological state than that afforded by SEM. Acknowledgments The author is very grateful to Professor T. Ushiki (Niigata University Graduate School of Medical and Dental Sciences) for his continuous support and encouragement. This study was supported in part by a Grant-in-Aid for Scientific Research (No. 20590186 to O.H.) from the Japan Society for the Promotion of Science. Conflict of interest The author reports no conflicts of interest. The author alone is responsible for the content and writing of the paper.
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