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Single-Molecule Imaging of Proteoglycans in the Pericellular Matrix
Biophysical Letter
Single-Molecule Imaging of Proteoglycans in the Pericellular Matrix Jan Scrimgeour,1,2,* Louis T. McLane,3,4 Patrick S. Chang,4 and Jennifer E. Curtis4,5 1 Department of Physics and 2Center for Advanced Materials Processing, Clarkson University, Potsdam, New York; 3School of Physics and Astronomy, Rochester Institute of Technology, Rochester, New York; and 4School of Physics and 5Parker H. Petit Institute for Bioengineering and Biosciences, Georgia Institute of Technology, Atlanta, Georgia
ABSTRACT The pericellular matrix is a robust, hyaluronan-rich polymer brush-like structure that controls access to the cell surface, and plays an important role in cell adhesion, migration, and proliferation. We report the observation of single bottlebrush proteoglycan dynamics in the pericellular matrix of living chondrocytes. Our investigations show that the pericellular matrix undergoes gross extension on the addition of exogenous aggrecan, and that this extension is significantly in excess of that observed in traditional particle exclusion assays. The mean-square displacement of single, bound proteoglycans increases with distance to cell surface, indicating reduced confinement by neighboring hyaluronan-aggrecan complexes. This is consistent with published data from quantitative particle exclusion assays that show openings in the pericellular matrix microstructure ranging from 150 nm near the cell surface to 400 nm near the cell edge. In addition, the mobility of tethered aggrecan drops significantly when the cell coat is enriched with bottlebrush proteoglycans. Single-molecule imaging in this thick polysaccharide matrix on living cells has significant promise in the drive to elucidate the role of the pericellular coat in human health.
The pericellular matrix (PCM) is a robust polymer layer that is directly anchored to the membrane of many living cells (1). The key components of this extended, often micronsthick cell coat are the polymer hyaluronan, hyaluronanbinding chondroitin sulfate bottlebrushes like versican or neurocan, and the hyaluronan (HA) surface receptor CD44 (2). A simple model for PCM organization is shown in Fig. 1 A. The ubiquity of the PCM’s components, and its appearance in diverse biological contexts, suggest that there is no single, defining function for the PCM, but rather it is involved in a range of processes that are important to the function of single cells and tissues. Previous work has shown that the PCM is involved in fundamental biophysical processes such as cell adhesion, migration, and proliferation (3–6). The role of the PCM in human health remains to be fully explored, but there is evidence to suggest that it is involved in the onset and progression of cancer (7,8), wound healing (9), and infertility (10). The challenges of determining the function of the PCM are strongly linked to two of its defining properties: low molecular density and very high levels of hydration. Even in vitro, investigations into the interaction between PCM Submitted October 5, 2016, and accepted for publication September 19, 2017. *Correspondence: [email protected] Editor: Philip LeDuc. https://doi.org/10.1016/j.bpj.2017.09.030 Ó 2017 Biophysical Society.
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components have required the use of novel methods, such as confocal fluorescence recovery after photobleaching (11,12) and quartz crystal microbalance (13). Invisible under traditional optical microscopy except for its ability to prevent large particles from reaching the cell surface (Fig. 1 B), it is only recently that techniques such as optical and atomic force probe microscopy (14–17), environmental scanning electron microscopy (3), fluorescence microscopy (18), and particle tracking (19–21) have begun to offer a glimpse into the structure of the PCM on living cells. Fig. 1 C shows fluorescently labeled proteoglycans incorporated into the PCM of a living chondrocyte cell, cultured on glass. The earliest particle tracking experiments, based on the gold labeling of proteoglycans, found that the PCM was likely to consist of HA molecules that are individually tethered to the cell surface, and showed that the chondroitin sulfate proteoglycan aggrecan (ACAN) was responsible for the extension of the PCM away from the cell surface (19). Other work looked at how size-dependent particle transport to the cell surface depends on bottlebrush proteoglycan content (22,23), and the role of the PCM in filtering, capturing, and possibly concentrating nanoparticles (24) and positively charged molecules (22,23). The outcomes of these studies depend on the dynamics of the bottlebrushes, as well as their total number. In this article, we use single-molecule fluorescence microscopy to track the dynamics of the bottlebrush proteoglycan
Biophysical Letter
FIGURE 1 (A) Structure of the PCM showing hyaluronan (HA) bound to the cell surface and the binding of proteoglycans to the HA in the extracellular space (nucleus, blue). (B) Brightfield image of a particle exclusion assay where the PCM was visualized by the introduction of fixed red blood cells into the culture. (C) x-z cross section of fluorescent proteoglycans in the PCM of a living chondrocyte imaged using confocal microscopy (PCM, green; membrane, white). (D) Schematic illustrating how PCM is illuminated using a highly angled laser beam (unilluminated PCM, gray; illuminated PCM, green).
ACAN in the PCM of living rat chondrocytes (RCJ-P). We investigate how the motion of tethered ACAN varies with location within the PCM and distance from the cell surface, and then quantify how these dynamics are suppressed by the crowding from other HA-tethered bottlebrushes at several different concentrations. These detailed localized measurements of single ACAN dynamics indirectly map out the PCM microstructure; we therefore compare the results with other structural characterizations of the PCM on the same cell type. Achieving single-molecule imaging in the PCM is challenging because of the large background signal produced by fluorescent ACAN that is free in the culture medium. To counter this, we adopted highly inclined laser illumination (see Fig. 1 D), after Tokunaga et al. (25), which minimizes out-of-focus excitation and allows for high-contrast imaging at depths of a few micrometers into the sample. Details of the single-molecule imaging set-up and sample preparation are available in the Supporting Information. Images of single fluorescent proteoglycans bound to the PCM are presented in Fig. 2. Single fluorescent ACAN (ACAN-488) molecules were embedded in the native RCJ-P PCM by incubation of the cell culture with an imaging buffer containing 5 fM ACAN-488 for 20 min before experiments began. In Fig. 2, three fluorescent proteoglycans tethered to the PCM are
identified. Traces of their x-y motion show that their range of motion increases with distance from the cell surface, ds. No bright fluorescence sources were observed in the PCM during control experiments (Fig. S1), in which the ACAN488 was left out of the imaging buffer. To investigate the dynamics of ACAN-488 in the native PCM, we first examined the mean-square displacements (MSDs) versus distance from the cell surface at three distinct lag times, t. This data is presented in Fig. 3 A and represents measurements from a total of 14 RCJ-P cells. Our first observation is that the MSDs measured in these experiments are substantially smaller than would be expected for ACAN diffusing freely in solution (diffusion coefficient 4.25 10 8 cm2/s), which would exhibit MSDs in the range of 0.53–2.12 mm2 for the relevant lag times (11). Here, the maximum reported MSD is 0.2 mm2. Previous investigations into the kinetics of this interaction showed that the dissociation rate for ACAN-binding HA is 3.96 10 3 s 1 (26). This corresponds to a bond lifetime of 252 s, which is substantially longer than our experiments, which lasted between 1 and 5 s. Therefore, we hypothesize that the tracked ACAN-488 are bound to HA in the PCM, and that our observations report directly on the dynamics of the HA-ACAN aggregates that make up the bulk of the chondrocyte PCM.
FIGURE 2 Brightfield and fluorescence images of two cultured RCJ-P cells. Arrows highlight single ACAN-488 at different distances (ds) from the cell surface. The trajectories of these proteoglycans are shown to the right of the images. The MSDs for these trajectories, stated with their SEs, are (i) 0.031 5 0.001 mm2, (ii) 0.057 5 0.001 mm2, and (iii) 0.103 5 0.003 mm2. To see this figure in color, go online.
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FIGURE 3 (A) MSD versus ds for ACAN-488 in native PCM for three distinct lag times. (B) MSD versus lag time the three ACAN-488 (those identified in Fig. 2) in the native PCM of a single cell. (C) MSD versus lag time for three ACAN-488 in the native PCM of a second cell. All error bars represent the mean 5 SE. To see this figure in color, go online.
To examine the trends in ACAN-488 dynamics at the single-cell level, the MSDs of individual ACAN-488 molecules in the native PCM of two cells are plotted against lag time in Fig. 3, B and C. Close to the surface of the cell (ds < 3.5 mm), these plots agree with the data presented in Fig. 3 A, showing that the MSD is effectively constant with lag time. This observation is consistent with the motion of a tethered and/or confined molecule. Linear fits to log-log representations of these data sets show that their slope is not statistically different from zero (p > 0.05). In contrast, at larger distances (ds > 3.5 mm), the data shows that the MSDs of molecules initially increase with lag time, before becoming constant at longer lag times. Linear fits to these data sets indicate that the slope is statistically different from zero (p << 0.05), and that the slope increases with distance from the cell surface. Fitting procedures and statistics (Tables S1 and S2) are reported in the Supporting Information. The single-cell data shows strong confinement of ACAN488 motion near the cell surface, whereas at larger distances from the cell, we observe increasingly weak confinement of the probe. This characteristic change in ACAN-488 confine-
ment is likely the result of a changing balance between motion that results directly from shape and bending fluctuations in the tethering HA-ACAN aggregate, and crowding of this aggregate induced by its neighbors. The data, however, does not immediately allow us to separate the contributions of the two factors. The properties of the RCJ-P PCM can be manipulated in a dramatic fashion by the addition of significant concentrations of exogenous ACAN to the imaging buffer (21,22,27). The ACAN binds to the HA strands, stretching them out. At saturation (182 mg/ml), it increases the average thickness of the RCJ-P PCM from 7 to 17.5 mm (22). Fig. 4 A compares the motion of the ACAN in the native PCM to that in the PCM after enrichment with exogenous ACAN. These observations were made by tracking individual tethered molecules in the PCM under native conditions and after the addition of 333 mg/ml of non-fluorescent exogenous ACAN. All MSDs presented were calculated using a lag time of 0.03125 s. The MSD of the molecules under each condition is plotted against distance from the cell surface. The data reveals a gross extension of some HA molecules within the PCM based on the largest distance from the
FIGURE 4 (A) MSD versus ds for ACAN-488 in native and exogenously modified PCM. The lines represent linear fits to the data. (B) MSD versus ds for ACAN-488 in exogenously modified PCM. Additional intermediate concentrations of exogenous ACAN are shown along with the data introduced in (A). The lines represent linear fits to the data. (C) Root-mean-square displacement (RMSD) versus ds for ACAN-488 in native PCM and PCM treated with 333 mg/ml ACAN. (D) Pore size of the PCM versus ds, estimated by a qPEA. To see this figure in color, go online.
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Biophysical Letter
cell surface at which tethered ACAN-488 can be observed. In addition, a considerable reduction in the mobility of molecules in the inner 2–10 mm of the PCM’s extension from the cells surface was observed at the high ACAN concentration. This is illustrated by a substantial change in the slope of linear fits made to each data set. Fig. 4 B extends these measurements with three additional concentrations of exogenous ACAN, showing the MSDs for ACAN tethered in the PCM under native conditions plus the following concentrations of exogenous ACAN: 49, 95, 182, or 333 mg/ml. Again, gross extension of the PCM is observed, with the results showing that the mobility of the ACAN at intermediate concentrations lies between the two extremes presented in Fig. 4 A. This data shows substantially more noise than the data at the extreme conditions. As a result, although the slopes of the linear fits to the new data sets follow the expected trend, with the slope reducing as the exogenous ACAN concentration increases (Table S3), they cannot be assumed to be statistically different from each other. The gross extension of the PCM at even low concentrations of exogenous ACAN is interesting, as it is considerably beyond the extension observed using large particle exclusion assays (15,22). There are several possible explanations that might account for this observation. If one assumes that all the HA is solely bound to the cell surface then the immediate conclusion is that there is at least a small population of HA molecules that are up to 25 mm in length (or 9 MDa). These molecules would be compressed during particle exclusion assays and difficult to detect with confocal microscopy. Although such molecules would typically be considered large even by the standards of HA, this is not improbable for chondrocytes and some other cell types. HA in healthy synovial fluid has an average mass of 7 MDa (28), whereas vitreous humor has 8 MDa HA (29). Further, it is known that HA synthase produces a broad distribution of sizes (30), so 9-MDa polymers in small amounts are not surprising. Another possibility is that there are some HA molecules that are attached to the PCM through entanglement or cross-linking. However, optical force probe assays provide evidence that there is minimal cross-linking within the RCJ-P PCM (15). Our current data, which suggests that these grossly extended gigantic molecules exist in relatively small quantities, unfortunately does not allow us to distinguish between these two options. The reduction in mobility of the tethered ACAN also provides some useful insights into the structure of the PCM. Of particular interest is the significant change observed between 182 and 333 mg/ml. These two points both lie on the plateau of a PCM thickness versus ACAN concentration curve, made using a large particle exclusion assay (22), which by itself suggests that the ACAN-binding sites might be fully occupied above exogenous ACAN concentrations of 182 mg/ml. However, in combination with the results from single-molecule tracking, it appears that full extension
of the PCM is reached well before full occupancy of the HA. This is consistent with confocal microscopy studies of ACAN-488 distribution of RCJ-P cells (22). The reduced MSD at 333 mg/ml is also consistent with reports of reduced openings in the RCJ-P PCM measured by quantitative particle exclusion assay (qPEA) (Fig. 4 C). The increase in MSD versus the distance of the observed ACAN-488 from the cell surface seen here is in qualitatively good agreement with measurements made using both optical force probe assays and qPEA (15,22), which show that the open spaces of the PCM increase with distance from the cell surface. Although the single-molecule measurements do not directly map the size of the open spaces, it is useful to compare the characteristic length scale (confinement length) of the ACAN-488 motion (Fig. 4 C) to measurements of the PCM pore size made using a qPEA (Fig. 4 D). We estimate the confinement length scale for ACAN-488 motion using the root-mean-square displacement of the tracked ACAN. The pore size of the PCM was estimated by the nanoparticle size found to penetrate into the PCM using qPEA analysis. The two distinct measurements provide estimates of the same order of magnitude, and both measure increases with distance to cell surface. However, it is notable that the trends observed are quite distinct. The confinement length for ACAN-488 motion is initially significantly larger than the PCM pore size estimated by the qPEA assay; however, the ACAN motion increases quite slowly with increasing distance to the cell surface (see Fig. S2 for direct comparison of data in Fig. 4, C and D). In contrast, the measured pore size of the PCM increases rapidly with distance to the cell surface, and near the outer edge of the PCM is larger than the characteristic length of the ACAN-488 motion. These comparisons suggest that even near the cell surface the confinement of the ACAN-488 by nearby HA-ACAN aggregates is quite weak, with fluctuations in the PCM’s microstructure enabling a relatively large range of motion compared to the pore size measured via qPEA. In addition, the crossing observed in the data (Fig. S2) correlates well with the appearance of nonzero slopes in the MSD versus lag time data shown in Fig. 3, B and C, suggesting that the effects of confinement weaken as the distance to the cell surface increases. We have used high-contrast, single-molecule imaging to probe the dynamics of HA-tethered ACAN within the pericellular coat of living rat chondrocytes. The data shows gross extension of the PCM on the addition of exogenous ACAN that is greater than measurements made in the same cell type by large particle exclusion assays. These observations may be due to a small population of long polymers grafted to the cell surface or tethered into the structure of the coat. The results show that even after the PCM reaches full extension, there are a significant number of unoccupied ACAN-binding sites along the HA backbone. We believe that single-molecule imaging in the PCM will
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open up new avenues of research into its biophysics and biochemistry by allowing direct access to many of its microscopic properties, and will facilitate a deeper understanding of the role of PCM in human health. SUPPORTING MATERIAL Supporting Materials and Methods, two figures, and three tables are available at http://www.biophysj.org/biophysj/supplemental/S0006-3495(17)31073-1.
AUTHOR CONTRIBUTIONS J.S. and J.E.C designed the research. J.S., L.T.M., and P.S.C. performed the research. J.S. and J.E.C. analyzed data and wrote the manuscript.
ACKNOWLEDGMENTS We are grateful to Hemaa Selvakumar for assistance with the confocal imaging experiments. J.E.C. and L.T.M. were supported by National Science Foundation grant 0955811. The Human Frontier Science Program (grant RGP0013/2010) provided partial support for J.S. and J.E.C.
REFERENCES 1. Toole, B. P. 2004. Hyaluronan: from extracellular glue to pericellular cue. Nat. Rev. Cancer. 4:528–539. 2. Knudson, C. B., and W. Knudson. 1993. Hyaluronan-binding proteins in development, tissue homeostasis, and disease. FASEB J. 7:1233–1241. 3. Cohen, M., E. Klein, ., L. Addadi. 2003. Organization and adhesive properties of the hyaluronan pericellular coat of chondrocytes and epithelial cells. Biophys. J. 85:1996–2005. 4. Cohen, M., D. Joester, ., B. Geiger. 2007. Hyaluronan in the pericellular coat: an additional layer of complexity in early cell adhesion events. Soft Matter. 3:327–332. 5. Evanko, S. P., J. C. Angello, and T. N. Wight. 1999. Formation of hyaluronan- and versican-rich pericellular matrix is required for proliferation and migration of vascular smooth muscle cells. Arterioscler. Thromb. Vasc. Biol. 19:1004–1013. 6. Itano, N., F. Atsumi, ., K. Kimata. 2002. Abnormal accumulation of hyaluronan matrix diminishes contact inhibition of cell growth and promotes cell migration. Proc. Natl. Acad. Sci. USA. 99:3609–3614. 7. Jones, L. M. H., M. J. Gardner, ., G. A. Turner. 1995. Hyaluronic acid secreted by mesothelial cells: a natural barrier to ovarian cancer cell adhesion. Clin. Exp. Metastasis. 13:373–380. 8. Turley, E. A., D. K. Wood, and J. B. McCarthy. 2016. Carcinoma cell hyaluronan as a ‘‘portable’’ cancerized prometastatic microenvironment. Cancer Res. 76:2507–2512. 9. Oksala, O., T. Salo, ., H. Larjava. 1995. Expression of proteoglycans and hyaluronan during wound healing. J. Histochem. Cytochem. 43:125–135. 10. Curtis, J. E. 2016. The mechanics of ovulation depend on an incredibly soft and sugar-rich extracellular matrix. Biophys. J. 110:2566–2567.
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11. Gribbon, P., and T. E. Hardingham. 1998. Macromolecular diffusion of biological polymers measured by confocal fluorescence recovery after photobleaching. Biophys. J. 75:1032–1039. 12. Scrimgeour, J., J. K. Cho, ., J. Curtis. 2011. Microfluidic dialysis cell for characterization of macromolecule interactions. Soft Matter. 7:4762–4767. 13. Richter, R. P., K. K. Hock, ., J. P. Spatz. 2007. Membrane-grafted hyaluronan films: a well-defined model system of glycoconjugate cell coats. J. Am. Chem. Soc. 129:5306–5307. 14. Ng, L., H.-H. Hung, ., A. Grodzinsky. 2007. Nanomechanical properties of individual chondrocytes and their developing growth factorstimulated pericellular matrix. J. Biomech. 40:1011–1023. 15. McLane, L. T., P. Chang, ., J. E. Curtis. 2013. Spatial organization and mechanical properties of the pericellular matrix on chondrocytes. Biophys. J. 104:986–996. 16. Dokukin, M., Y. Ablaeva, ., I. Sokolov. 2016. Pericellular brush and mechanics of guinea pig fibroblast cells studied with AFM. Biophys. J. 111:236–246. 17. Simon, M., M. Dokukin, ., C. Staii. 2016. Load rate and temperature dependent mechanical properties of the cortical neuron and its pericellular layer measured by atomic force microscopy. Langmuir. 32:1111–1119. 18. Rilla, K., R. Tiihonen, ., R. Tammi. 2008. Pericellular hyaluronan coat visualized in live cells with a fluorescent probe is scaffolded by plasma membrane protrusions. J. Histochem. Cytochem. 56:901–910. 19. Lee, G. M., B. Johnstone, ., B. Caterson. 1993. The dynamic structure of the pericellular matrix on living cells. J. Cell Biol. 123:1899–1907. 20. Boehm, H., T. A. Mundinger, ., J. E. Curtis. 2009. Mapping the mechanics and macromolecular organization of hyaluronan-rich cell coats. Soft Matter. 5:4331–4337. 21. Nijenhuis, N., D. Mizuno, ., C. F. Schmidt. 2012. High-resolution microrheology in the pericellular matrix of prostate cancer cells. J. R. Soc. Interface. 9:1733–1744. 22. Chang, P. S., L. T. McLane, ., J. E. Curtis. 2016. Cell surface access is modulated by tethered bottlebrush proteoglycans. Biophys. J. 110:2739–2750. 23. Heldin, P., and H. Pertoft. 1993. Synthesis and assembly of the hyaluronan-containing coats around normal human mesothelial cells. Exp. Cell Res. 208:422–429. 24. Zhou, R., H. Zhou, ., E. S. Yeung. 2012. Pericellular matrix enhances retention and cellular uptake of nanoparticles. J. Am. Chem. Soc. 134:13404–13409. 25. Tokunaga, M., N. Imamoto, and K. Sakata-Sogawa. 2008. Highly inclined thin illumination enables clear single-molecule imaging in cells. Nat. Methods. 5:159–161. 26. Watanabe, H., S. C. Cheung, ., Y. Yamada. 1997. Identification of hyaluronan-binding domains of aggrecan. J. Biol. Chem. 272:28057–28065. 27. Attili, S., and R. P. Richter. 2013. Self-assembly and elasticity of hierarchical proteoglycan–hyaluronan brushes. Soft Matter. 9:10473–10483. 28. Kogan, G., L. Soltes, ., P. Gemeiner. 2007. Hyaluronic acid: a natural biopolymer with a broad range of biomedical and industrial applications. Biotechnol. Lett. 29:17–25. 29. Burdick, J. A., and G. D. Prestwich. 2011. Hyaluronic acid hydrogels for biomedical applications. Adv. Mater. 23:H41–H56. 30. Baggenstoss, B. A., and P. H. Weigel. 2006. Size exclusion chromatography-multiangle laser light scattering analysis of hyaluronan size distributions made by membrane-bound hyaluronan synthase. Anal. Biochem. 352:243–251.
Single-Molecule Imaging of Proteoglycans in the Pericellular Matrix Jan Scrimgeour,1,2,* Louis T. McLane,3,4 Patrick S. Chang,4 and Jennifer E. Curtis4,5 1 Department of Physics and 2Center for Advanced Materials Processing, Clarkson University, Potsdam, New York; 3School of Physics and Astronomy, Rochester Institute of Technology, Rochester, New York; and 4School of Physics and 5Parker H. Petit Institute for Bioengineering and Biosciences, Georgia Institute of Technology, Atlanta, Georgia
ABSTRACT The pericellular matrix is a robust, hyaluronan-rich polymer brush-like structure that controls access to the cell surface, and plays an important role in cell adhesion, migration, and proliferation. We report the observation of single bottlebrush proteoglycan dynamics in the pericellular matrix of living chondrocytes. Our investigations show that the pericellular matrix undergoes gross extension on the addition of exogenous aggrecan, and that this extension is significantly in excess of that observed in traditional particle exclusion assays. The mean-square displacement of single, bound proteoglycans increases with distance to cell surface, indicating reduced confinement by neighboring hyaluronan-aggrecan complexes. This is consistent with published data from quantitative particle exclusion assays that show openings in the pericellular matrix microstructure ranging from 150 nm near the cell surface to 400 nm near the cell edge. In addition, the mobility of tethered aggrecan drops significantly when the cell coat is enriched with bottlebrush proteoglycans. Single-molecule imaging in this thick polysaccharide matrix on living cells has significant promise in the drive to elucidate the role of the pericellular coat in human health.
The pericellular matrix (PCM) is a robust polymer layer that is directly anchored to the membrane of many living cells (1). The key components of this extended, often micronsthick cell coat are the polymer hyaluronan, hyaluronanbinding chondroitin sulfate bottlebrushes like versican or neurocan, and the hyaluronan (HA) surface receptor CD44 (2). A simple model for PCM organization is shown in Fig. 1 A. The ubiquity of the PCM’s components, and its appearance in diverse biological contexts, suggest that there is no single, defining function for the PCM, but rather it is involved in a range of processes that are important to the function of single cells and tissues. Previous work has shown that the PCM is involved in fundamental biophysical processes such as cell adhesion, migration, and proliferation (3–6). The role of the PCM in human health remains to be fully explored, but there is evidence to suggest that it is involved in the onset and progression of cancer (7,8), wound healing (9), and infertility (10). The challenges of determining the function of the PCM are strongly linked to two of its defining properties: low molecular density and very high levels of hydration. Even in vitro, investigations into the interaction between PCM Submitted October 5, 2016, and accepted for publication September 19, 2017. *Correspondence: [email protected] Editor: Philip LeDuc. https://doi.org/10.1016/j.bpj.2017.09.030 Ó 2017 Biophysical Society.
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components have required the use of novel methods, such as confocal fluorescence recovery after photobleaching (11,12) and quartz crystal microbalance (13). Invisible under traditional optical microscopy except for its ability to prevent large particles from reaching the cell surface (Fig. 1 B), it is only recently that techniques such as optical and atomic force probe microscopy (14–17), environmental scanning electron microscopy (3), fluorescence microscopy (18), and particle tracking (19–21) have begun to offer a glimpse into the structure of the PCM on living cells. Fig. 1 C shows fluorescently labeled proteoglycans incorporated into the PCM of a living chondrocyte cell, cultured on glass. The earliest particle tracking experiments, based on the gold labeling of proteoglycans, found that the PCM was likely to consist of HA molecules that are individually tethered to the cell surface, and showed that the chondroitin sulfate proteoglycan aggrecan (ACAN) was responsible for the extension of the PCM away from the cell surface (19). Other work looked at how size-dependent particle transport to the cell surface depends on bottlebrush proteoglycan content (22,23), and the role of the PCM in filtering, capturing, and possibly concentrating nanoparticles (24) and positively charged molecules (22,23). The outcomes of these studies depend on the dynamics of the bottlebrushes, as well as their total number. In this article, we use single-molecule fluorescence microscopy to track the dynamics of the bottlebrush proteoglycan
Biophysical Letter
FIGURE 1 (A) Structure of the PCM showing hyaluronan (HA) bound to the cell surface and the binding of proteoglycans to the HA in the extracellular space (nucleus, blue). (B) Brightfield image of a particle exclusion assay where the PCM was visualized by the introduction of fixed red blood cells into the culture. (C) x-z cross section of fluorescent proteoglycans in the PCM of a living chondrocyte imaged using confocal microscopy (PCM, green; membrane, white). (D) Schematic illustrating how PCM is illuminated using a highly angled laser beam (unilluminated PCM, gray; illuminated PCM, green).
ACAN in the PCM of living rat chondrocytes (RCJ-P). We investigate how the motion of tethered ACAN varies with location within the PCM and distance from the cell surface, and then quantify how these dynamics are suppressed by the crowding from other HA-tethered bottlebrushes at several different concentrations. These detailed localized measurements of single ACAN dynamics indirectly map out the PCM microstructure; we therefore compare the results with other structural characterizations of the PCM on the same cell type. Achieving single-molecule imaging in the PCM is challenging because of the large background signal produced by fluorescent ACAN that is free in the culture medium. To counter this, we adopted highly inclined laser illumination (see Fig. 1 D), after Tokunaga et al. (25), which minimizes out-of-focus excitation and allows for high-contrast imaging at depths of a few micrometers into the sample. Details of the single-molecule imaging set-up and sample preparation are available in the Supporting Information. Images of single fluorescent proteoglycans bound to the PCM are presented in Fig. 2. Single fluorescent ACAN (ACAN-488) molecules were embedded in the native RCJ-P PCM by incubation of the cell culture with an imaging buffer containing 5 fM ACAN-488 for 20 min before experiments began. In Fig. 2, three fluorescent proteoglycans tethered to the PCM are
identified. Traces of their x-y motion show that their range of motion increases with distance from the cell surface, ds. No bright fluorescence sources were observed in the PCM during control experiments (Fig. S1), in which the ACAN488 was left out of the imaging buffer. To investigate the dynamics of ACAN-488 in the native PCM, we first examined the mean-square displacements (MSDs) versus distance from the cell surface at three distinct lag times, t. This data is presented in Fig. 3 A and represents measurements from a total of 14 RCJ-P cells. Our first observation is that the MSDs measured in these experiments are substantially smaller than would be expected for ACAN diffusing freely in solution (diffusion coefficient 4.25 10 8 cm2/s), which would exhibit MSDs in the range of 0.53–2.12 mm2 for the relevant lag times (11). Here, the maximum reported MSD is 0.2 mm2. Previous investigations into the kinetics of this interaction showed that the dissociation rate for ACAN-binding HA is 3.96 10 3 s 1 (26). This corresponds to a bond lifetime of 252 s, which is substantially longer than our experiments, which lasted between 1 and 5 s. Therefore, we hypothesize that the tracked ACAN-488 are bound to HA in the PCM, and that our observations report directly on the dynamics of the HA-ACAN aggregates that make up the bulk of the chondrocyte PCM.
FIGURE 2 Brightfield and fluorescence images of two cultured RCJ-P cells. Arrows highlight single ACAN-488 at different distances (ds) from the cell surface. The trajectories of these proteoglycans are shown to the right of the images. The MSDs for these trajectories, stated with their SEs, are (i) 0.031 5 0.001 mm2, (ii) 0.057 5 0.001 mm2, and (iii) 0.103 5 0.003 mm2. To see this figure in color, go online.
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FIGURE 3 (A) MSD versus ds for ACAN-488 in native PCM for three distinct lag times. (B) MSD versus lag time the three ACAN-488 (those identified in Fig. 2) in the native PCM of a single cell. (C) MSD versus lag time for three ACAN-488 in the native PCM of a second cell. All error bars represent the mean 5 SE. To see this figure in color, go online.
To examine the trends in ACAN-488 dynamics at the single-cell level, the MSDs of individual ACAN-488 molecules in the native PCM of two cells are plotted against lag time in Fig. 3, B and C. Close to the surface of the cell (ds < 3.5 mm), these plots agree with the data presented in Fig. 3 A, showing that the MSD is effectively constant with lag time. This observation is consistent with the motion of a tethered and/or confined molecule. Linear fits to log-log representations of these data sets show that their slope is not statistically different from zero (p > 0.05). In contrast, at larger distances (ds > 3.5 mm), the data shows that the MSDs of molecules initially increase with lag time, before becoming constant at longer lag times. Linear fits to these data sets indicate that the slope is statistically different from zero (p << 0.05), and that the slope increases with distance from the cell surface. Fitting procedures and statistics (Tables S1 and S2) are reported in the Supporting Information. The single-cell data shows strong confinement of ACAN488 motion near the cell surface, whereas at larger distances from the cell, we observe increasingly weak confinement of the probe. This characteristic change in ACAN-488 confine-
ment is likely the result of a changing balance between motion that results directly from shape and bending fluctuations in the tethering HA-ACAN aggregate, and crowding of this aggregate induced by its neighbors. The data, however, does not immediately allow us to separate the contributions of the two factors. The properties of the RCJ-P PCM can be manipulated in a dramatic fashion by the addition of significant concentrations of exogenous ACAN to the imaging buffer (21,22,27). The ACAN binds to the HA strands, stretching them out. At saturation (182 mg/ml), it increases the average thickness of the RCJ-P PCM from 7 to 17.5 mm (22). Fig. 4 A compares the motion of the ACAN in the native PCM to that in the PCM after enrichment with exogenous ACAN. These observations were made by tracking individual tethered molecules in the PCM under native conditions and after the addition of 333 mg/ml of non-fluorescent exogenous ACAN. All MSDs presented were calculated using a lag time of 0.03125 s. The MSD of the molecules under each condition is plotted against distance from the cell surface. The data reveals a gross extension of some HA molecules within the PCM based on the largest distance from the
FIGURE 4 (A) MSD versus ds for ACAN-488 in native and exogenously modified PCM. The lines represent linear fits to the data. (B) MSD versus ds for ACAN-488 in exogenously modified PCM. Additional intermediate concentrations of exogenous ACAN are shown along with the data introduced in (A). The lines represent linear fits to the data. (C) Root-mean-square displacement (RMSD) versus ds for ACAN-488 in native PCM and PCM treated with 333 mg/ml ACAN. (D) Pore size of the PCM versus ds, estimated by a qPEA. To see this figure in color, go online.
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Biophysical Letter
cell surface at which tethered ACAN-488 can be observed. In addition, a considerable reduction in the mobility of molecules in the inner 2–10 mm of the PCM’s extension from the cells surface was observed at the high ACAN concentration. This is illustrated by a substantial change in the slope of linear fits made to each data set. Fig. 4 B extends these measurements with three additional concentrations of exogenous ACAN, showing the MSDs for ACAN tethered in the PCM under native conditions plus the following concentrations of exogenous ACAN: 49, 95, 182, or 333 mg/ml. Again, gross extension of the PCM is observed, with the results showing that the mobility of the ACAN at intermediate concentrations lies between the two extremes presented in Fig. 4 A. This data shows substantially more noise than the data at the extreme conditions. As a result, although the slopes of the linear fits to the new data sets follow the expected trend, with the slope reducing as the exogenous ACAN concentration increases (Table S3), they cannot be assumed to be statistically different from each other. The gross extension of the PCM at even low concentrations of exogenous ACAN is interesting, as it is considerably beyond the extension observed using large particle exclusion assays (15,22). There are several possible explanations that might account for this observation. If one assumes that all the HA is solely bound to the cell surface then the immediate conclusion is that there is at least a small population of HA molecules that are up to 25 mm in length (or 9 MDa). These molecules would be compressed during particle exclusion assays and difficult to detect with confocal microscopy. Although such molecules would typically be considered large even by the standards of HA, this is not improbable for chondrocytes and some other cell types. HA in healthy synovial fluid has an average mass of 7 MDa (28), whereas vitreous humor has 8 MDa HA (29). Further, it is known that HA synthase produces a broad distribution of sizes (30), so 9-MDa polymers in small amounts are not surprising. Another possibility is that there are some HA molecules that are attached to the PCM through entanglement or cross-linking. However, optical force probe assays provide evidence that there is minimal cross-linking within the RCJ-P PCM (15). Our current data, which suggests that these grossly extended gigantic molecules exist in relatively small quantities, unfortunately does not allow us to distinguish between these two options. The reduction in mobility of the tethered ACAN also provides some useful insights into the structure of the PCM. Of particular interest is the significant change observed between 182 and 333 mg/ml. These two points both lie on the plateau of a PCM thickness versus ACAN concentration curve, made using a large particle exclusion assay (22), which by itself suggests that the ACAN-binding sites might be fully occupied above exogenous ACAN concentrations of 182 mg/ml. However, in combination with the results from single-molecule tracking, it appears that full extension
of the PCM is reached well before full occupancy of the HA. This is consistent with confocal microscopy studies of ACAN-488 distribution of RCJ-P cells (22). The reduced MSD at 333 mg/ml is also consistent with reports of reduced openings in the RCJ-P PCM measured by quantitative particle exclusion assay (qPEA) (Fig. 4 C). The increase in MSD versus the distance of the observed ACAN-488 from the cell surface seen here is in qualitatively good agreement with measurements made using both optical force probe assays and qPEA (15,22), which show that the open spaces of the PCM increase with distance from the cell surface. Although the single-molecule measurements do not directly map the size of the open spaces, it is useful to compare the characteristic length scale (confinement length) of the ACAN-488 motion (Fig. 4 C) to measurements of the PCM pore size made using a qPEA (Fig. 4 D). We estimate the confinement length scale for ACAN-488 motion using the root-mean-square displacement of the tracked ACAN. The pore size of the PCM was estimated by the nanoparticle size found to penetrate into the PCM using qPEA analysis. The two distinct measurements provide estimates of the same order of magnitude, and both measure increases with distance to cell surface. However, it is notable that the trends observed are quite distinct. The confinement length for ACAN-488 motion is initially significantly larger than the PCM pore size estimated by the qPEA assay; however, the ACAN motion increases quite slowly with increasing distance to the cell surface (see Fig. S2 for direct comparison of data in Fig. 4, C and D). In contrast, the measured pore size of the PCM increases rapidly with distance to the cell surface, and near the outer edge of the PCM is larger than the characteristic length of the ACAN-488 motion. These comparisons suggest that even near the cell surface the confinement of the ACAN-488 by nearby HA-ACAN aggregates is quite weak, with fluctuations in the PCM’s microstructure enabling a relatively large range of motion compared to the pore size measured via qPEA. In addition, the crossing observed in the data (Fig. S2) correlates well with the appearance of nonzero slopes in the MSD versus lag time data shown in Fig. 3, B and C, suggesting that the effects of confinement weaken as the distance to the cell surface increases. We have used high-contrast, single-molecule imaging to probe the dynamics of HA-tethered ACAN within the pericellular coat of living rat chondrocytes. The data shows gross extension of the PCM on the addition of exogenous ACAN that is greater than measurements made in the same cell type by large particle exclusion assays. These observations may be due to a small population of long polymers grafted to the cell surface or tethered into the structure of the coat. The results show that even after the PCM reaches full extension, there are a significant number of unoccupied ACAN-binding sites along the HA backbone. We believe that single-molecule imaging in the PCM will
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open up new avenues of research into its biophysics and biochemistry by allowing direct access to many of its microscopic properties, and will facilitate a deeper understanding of the role of PCM in human health. SUPPORTING MATERIAL Supporting Materials and Methods, two figures, and three tables are available at http://www.biophysj.org/biophysj/supplemental/S0006-3495(17)31073-1.
AUTHOR CONTRIBUTIONS J.S. and J.E.C designed the research. J.S., L.T.M., and P.S.C. performed the research. J.S. and J.E.C. analyzed data and wrote the manuscript.
ACKNOWLEDGMENTS We are grateful to Hemaa Selvakumar for assistance with the confocal imaging experiments. J.E.C. and L.T.M. were supported by National Science Foundation grant 0955811. The Human Frontier Science Program (grant RGP0013/2010) provided partial support for J.S. and J.E.C.
REFERENCES 1. Toole, B. P. 2004. Hyaluronan: from extracellular glue to pericellular cue. Nat. Rev. Cancer. 4:528–539. 2. Knudson, C. B., and W. Knudson. 1993. Hyaluronan-binding proteins in development, tissue homeostasis, and disease. FASEB J. 7:1233–1241. 3. Cohen, M., E. Klein, ., L. Addadi. 2003. Organization and adhesive properties of the hyaluronan pericellular coat of chondrocytes and epithelial cells. Biophys. J. 85:1996–2005. 4. Cohen, M., D. Joester, ., B. Geiger. 2007. Hyaluronan in the pericellular coat: an additional layer of complexity in early cell adhesion events. Soft Matter. 3:327–332. 5. Evanko, S. P., J. C. Angello, and T. N. Wight. 1999. Formation of hyaluronan- and versican-rich pericellular matrix is required for proliferation and migration of vascular smooth muscle cells. Arterioscler. Thromb. Vasc. Biol. 19:1004–1013. 6. Itano, N., F. Atsumi, ., K. Kimata. 2002. Abnormal accumulation of hyaluronan matrix diminishes contact inhibition of cell growth and promotes cell migration. Proc. Natl. Acad. Sci. USA. 99:3609–3614. 7. Jones, L. M. H., M. J. Gardner, ., G. A. Turner. 1995. Hyaluronic acid secreted by mesothelial cells: a natural barrier to ovarian cancer cell adhesion. Clin. Exp. Metastasis. 13:373–380. 8. Turley, E. A., D. K. Wood, and J. B. McCarthy. 2016. Carcinoma cell hyaluronan as a ‘‘portable’’ cancerized prometastatic microenvironment. Cancer Res. 76:2507–2512. 9. Oksala, O., T. Salo, ., H. Larjava. 1995. Expression of proteoglycans and hyaluronan during wound healing. J. Histochem. Cytochem. 43:125–135. 10. Curtis, J. E. 2016. The mechanics of ovulation depend on an incredibly soft and sugar-rich extracellular matrix. Biophys. J. 110:2566–2567.
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11. Gribbon, P., and T. E. Hardingham. 1998. Macromolecular diffusion of biological polymers measured by confocal fluorescence recovery after photobleaching. Biophys. J. 75:1032–1039. 12. Scrimgeour, J., J. K. Cho, ., J. Curtis. 2011. Microfluidic dialysis cell for characterization of macromolecule interactions. Soft Matter. 7:4762–4767. 13. Richter, R. P., K. K. Hock, ., J. P. Spatz. 2007. Membrane-grafted hyaluronan films: a well-defined model system of glycoconjugate cell coats. J. Am. Chem. Soc. 129:5306–5307. 14. Ng, L., H.-H. Hung, ., A. Grodzinsky. 2007. Nanomechanical properties of individual chondrocytes and their developing growth factorstimulated pericellular matrix. J. Biomech. 40:1011–1023. 15. McLane, L. T., P. Chang, ., J. E. Curtis. 2013. Spatial organization and mechanical properties of the pericellular matrix on chondrocytes. Biophys. J. 104:986–996. 16. Dokukin, M., Y. Ablaeva, ., I. Sokolov. 2016. Pericellular brush and mechanics of guinea pig fibroblast cells studied with AFM. Biophys. J. 111:236–246. 17. Simon, M., M. Dokukin, ., C. Staii. 2016. Load rate and temperature dependent mechanical properties of the cortical neuron and its pericellular layer measured by atomic force microscopy. Langmuir. 32:1111–1119. 18. Rilla, K., R. Tiihonen, ., R. Tammi. 2008. Pericellular hyaluronan coat visualized in live cells with a fluorescent probe is scaffolded by plasma membrane protrusions. J. Histochem. Cytochem. 56:901–910. 19. Lee, G. M., B. Johnstone, ., B. Caterson. 1993. The dynamic structure of the pericellular matrix on living cells. J. Cell Biol. 123:1899–1907. 20. Boehm, H., T. A. Mundinger, ., J. E. Curtis. 2009. Mapping the mechanics and macromolecular organization of hyaluronan-rich cell coats. Soft Matter. 5:4331–4337. 21. Nijenhuis, N., D. Mizuno, ., C. F. Schmidt. 2012. High-resolution microrheology in the pericellular matrix of prostate cancer cells. J. R. Soc. Interface. 9:1733–1744. 22. Chang, P. S., L. T. McLane, ., J. E. Curtis. 2016. Cell surface access is modulated by tethered bottlebrush proteoglycans. Biophys. J. 110:2739–2750. 23. Heldin, P., and H. Pertoft. 1993. Synthesis and assembly of the hyaluronan-containing coats around normal human mesothelial cells. Exp. Cell Res. 208:422–429. 24. Zhou, R., H. Zhou, ., E. S. Yeung. 2012. Pericellular matrix enhances retention and cellular uptake of nanoparticles. J. Am. Chem. Soc. 134:13404–13409. 25. Tokunaga, M., N. Imamoto, and K. Sakata-Sogawa. 2008. Highly inclined thin illumination enables clear single-molecule imaging in cells. Nat. Methods. 5:159–161. 26. Watanabe, H., S. C. Cheung, ., Y. Yamada. 1997. Identification of hyaluronan-binding domains of aggrecan. J. Biol. Chem. 272:28057–28065. 27. Attili, S., and R. P. Richter. 2013. Self-assembly and elasticity of hierarchical proteoglycan–hyaluronan brushes. Soft Matter. 9:10473–10483. 28. Kogan, G., L. Soltes, ., P. Gemeiner. 2007. Hyaluronic acid: a natural biopolymer with a broad range of biomedical and industrial applications. Biotechnol. Lett. 29:17–25. 29. Burdick, J. A., and G. D. Prestwich. 2011. Hyaluronic acid hydrogels for biomedical applications. Adv. Mater. 23:H41–H56. 30. Baggenstoss, B. A., and P. H. Weigel. 2006. Size exclusion chromatography-multiangle laser light scattering analysis of hyaluronan size distributions made by membrane-bound hyaluronan synthase. Anal. Biochem. 352:243–251.