In situ 3D visualization of biomineralization matrix proteins
Graphical abstract
Introduction
Biomineralized skeletons evolved over 660 MYA and by the Ediacaran-Cambrian transition, were present in members of all major animal phyla (Maloof et al., 2010). Research on extant species has revealed that mineralized tissues across taxa comprise many different types of minerals and organic macromolecules that are arranged in different structural motifs, leading to diverse biomineral morphologies (Lowenstam, 1989). These organic macromolecules are a fundamental component of biomineral formation, being responsible for the mediation of mineral precursors, selection of mineral polymorph, and regulation of mineral growth (Mass et al., 2014, Pouget et al., 2009, Söllner et al., 2003). Furthermore, as many of the minerals by themselves are poor building materials, the architecture of the organic matrix, which influences how the mineral and organic molecules are organized, is clearly critical to the unique material properties of biominerals (Weiner and Addadi, 1997).
Identifying the spatial organization of macromolecules within the organic matrix is key to unlocking these unusual material properties but to our knowledge, no method has been developed that allows for both the characterization and 3D visualization of organic molecules in situ. Several techniques can visualize the 2D locations of macromolecules, but only those present in high abundance (Fang et al., 2011, Gal et al., 2016, Mass et al., 2014). Similarly, although organic components are easy to identify through mass spectrometry, this requires separation of the macromolecules from the dominant mineral phase (which can be up to 99% w/w), resulting in the decoupling of macromolecules from their locations and thus destroying any spatial information about their timing of expression.
To address this limitation, we developed a modified clearing protocol (CLARITY, ACT-PRESTO) (Chung et al., 2013, Lee et al., 2016) to examine the organic matrix of biomineral structures, which we then applied to otoliths from Atlantic salmon (Salmo salar). CLARITY has been successfully employed to image brain and bone marrow proteins (Greenbaum et al., 2017, Lee et al., 2016, Tomer et al., 2014). Here, we describe the results of our method, where we completely removed the otolith mineral phase, specifically calcium carbonate in the form of aragonite, while preserving the architecture and spatial integrities of organic matrix proteins. Our method allows for successful antibody penetration and, for the first time, three-dimensional imaging of key biomineral proteins.
Otoliths exist as three pairs of bilaterally oriented structures in the inner ears of all bony fish (Campana and Neilson, 1985, Thomas and Swearer, 2019). They allow for the fine scale detection of minute changes in water pressure – essential for hearing and balance in an aqueous environment. Otoliths are an unusual biomineral because they form in embryo (Campana and Neilson, 1985) and grow continually through the deposition of a double layer of proteinaceous and aragonite-dominant material during the crepuscular periods of the day (Pannella, 1971, Thomas and Swearer, 2019). Despite their widespread application in fish and fisheries research, the circadian mechanisms that regulate daily increment deposition remain unconfirmed (Izzo et al., 2016). We, and others, previously identified two otolith proteins likely involved in increment formation and regulation – Neuroserpin and Secreted protein acidic and rich in cysteine (SPARC/Osteonectin/BM40) (Kang et al., 2008, Thomas et al., 2019). Neuroserpin, a known serine protease inhibitor, likely initiates growth of the proteinaceous layer through the inhibition of a proteolytic enzyme cascade, while SPARC likely plays a dual role, being able to bind both scaffolding collagen proteins and calcium ions during growth of the aragonite-dominant layer (Thomas et al., 2019).
Section snippets
Chemicals
All samples, buffers and solutions were prepared using ultra-pure Milli-Q H2O (18.2 MΩ; Merck Millipore, Australia). Unless otherwise noted, all chemicals were purchased from Sigma Aldrich (Castle Hill, NSW, Australia).
Modified CLARITY and ACT-PRESTO protocols
Otoliths were dissected from post-smolt Atlantic salmon (age = ca. 2.5 y), rinsed in ultra-pure Milli-Q H2O and fixed for 24 h at 4 °C in 4% paraformaldehyde in 0.1 M phosphate buffer solution (PB; pH 7.35). They were then washed 3× in 0.1 M PB, before undergoing a modified
Results and discussion
The overall otolith protein architecture was revealed through Oriole™ staining and light-sheet imaging (Fig. 1). Despite complete removal of mineral, the external morphology and internal microstructure of the otolith were preserved, including the sulcus acusticus, antirostrum, rostrum, and postrostrum (Campana, 2004). Notably, annuli, bands corresponding to seasonal fluctuations in metabolism and growth were clearly evident (Campana and Neilson, 1985). Through shotgun proteomics, we identified
CRediT authorship contribution statement
Oliver R.B. Thomas: Conceptualization, Data curation, Funding acquisition, Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review & editing. Kay L. Richards: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Visualization, Writing - review & editing. Steven Petrou: Conceptualization, Methodology, Resources, Supervision, Writing - review & editing. Blaine R. Roberts: Conceptualization, Methodology, Resources, Writing -
Declaration of Competing Interest
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
Acknowledgements
We thank N. Williamson for assistance with protein sequencing, P. Jusuf for antibodies, E. Kapp for assistance with proteomics and F. Warren-Myers for supply of otoliths. This work was funded by the University of Melbourne.
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