Elsevier

Colloids and Surfaces B: Biointerfaces

Volume 169, 1 September 2018, Pages 462-469
Colloids and Surfaces B: Biointerfaces

Determination of the molecular assembly of actin and actin-binding proteins using photoluminescence

https://doi.org/10.1016/j.colsurfb.2018.05.043Get rights and content

Highlights

  • Modification of actin samples with different structures using binding protein.

  • Method for monitoring of actin structure from monomer to polymerized protein.

  • Polymerization rate of actin under several temperatures and heat energy.

  • Label-free detection of protein in both of in vivo and in vitro conditions.

Abstract

Actin, the most abundant protein in cells, polymerizes into filaments that play key roles in many cellular dynamics. To understand cell dynamics and functions, it is essential to examine the cytoskeleton structure organized by actin and actin-binding proteins. Here, we pave the way for determining the molecular assembly of the actin cytoskeleton using direct photonic in situ analysis, providing the photoluminescence characteristics of actin as a function of filament length and bundling, without labeling. In experiments for monomeric and filamentous actin reconstituted in vitro, structural forms of actin are identified from the peak positions and intensities of photoluminescence. Actin monomers exhibited small intensity emission peaks at 334 nm, whereas filamentous and bundled actin showed a shifted peak at 323 nm with higher intensity. The peak shift was found to be proportional to the length of the actin filament. With probing structural change of actin in various cells in vivo, our study provides an efficient and precise analytical in situ tool to examine the cytoskeleton structure. It will be beneficial for elucidating the mechanism of various cellular functions such as cell migration, differentiation, cytokinesis and adhesion. Furthermore, our technique can be applied to detect the alterations in the cell cytoskeleton that can occur during many pathological processes.

Graphical abstract

Actin’s structure and assembly behavior are determined by light absorption and photoluminescence characteristics under both in vitro and in vivo conditions.

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Introduction

The cytoskeleton is a primary structural component that determines the cell morphology. The cytoskeleton is also dynamic and changes its shape to play a role in many cellular functions. For example, filopodia and lamellipodia, dynamic cytoplasmic protrusions formed during cell migration, consist of thick bundles and orthogonal networks of actin filaments [1]. Networks of actin filaments aligned along cell boundaries provide mechanical support in epithelia [2]. A main constituent of the cytoskeleton is actin which is the most abundant protein in cells. The globular actin (G-actin) polymerizes into the filamentous actin (F-actin) in the presence of adenosine triphosphate (ATP) and divalent cations [3], [4], [5].

Both G-actin and F-actin interact with various actin-binding proteins (ABPs) and they assemble into higher-order structures. ABP α-actinin and fascin bind several actin filaments together to form thick bundles that are observed in filopodia and stress fibers, respectively [6], [7], [8]. Homogeneous network structures are organized by actin filaments cross-linked with ABP filamin [9]. The concentration of cross-linking ABP relative to actin determines the pore size of the actin network. Recent studies found that the mechanical responses of actin networks depended on the type of ABP as well as the constituent concentration [10], [11], [12]. The average length of actin filaments is regulated by a capping or severing ABP, such as gelsolin [4], [13]. Since actin in cells regulates organization of the cytoskeleton architecture according to the functional demands, investigating actin assembly in situ is essential to understanding cellular behaviors.

However, it is difficult to probe the actin cytoskeleton in cells due to the complexity of the cell and the lack of appropriate technique. Immuno-staining and transfection techniques using light microscopy have been widely used to detect actin in cells at high selectivity. However, the required processes of fixation and labeling can damage the cells and continuous monitoring of actin dynamics is limited with these techniques [14], [15]. A myopathy diagnostic tool employed label-free Raman spectroscopy to monitor muscle mutants by examining the biochemical responses of actin, myosin and other skeletal proteins [16]. Compared to optical measurements, electron microscopy allows the imaging of actin without a fluorescent tag at a higher spatial resolution of ∼ 50 pm [17], [18], [19]. However, it also requires a particular step in sample preparation, such as metal coating, and further requires a high vacuum condition for operation.

In this study, we obtained the absorbance and photoluminescence (PL) of actin polymerized with various types of actin-binding proteins in order to probe the actin structures in situ without labeling. Both optical measurements have been useful in examining biomaterials such as DNA and proteins without any detrimental indicator [20]. Absorption spectroscopy, which measures the light transmittance of a suspended sample solution in the ultraviolet (UV) to visible range, has been utilized to determine the presence and concentration of proteins in a sample solution [21], [22]. PL, which is light emission from a sample that has been excited by a photon, has been used to mark the biomolecules with fluorescent indicators. Using PL, bioimaging has been studied with various fluorescent dyes using charge-coupled devices for light detection. We measured the PL spectrum under UV excitation to determine the actin structure. G-actin, F-actin and bundled actin were regulated by adding ABPs such as gelsolin and α-actinin [7], [23], which showed remarkably different features in absorbance and PL. The actin monomer showed a PL emission peak located at 334 nm and polymerized actins provided a shifted peak at 323 nm with a higher intensity. Bundled actin exhibited the strongest PL emission among samples due to the bulk structure of the protein. We investigated the actin polymerization process according to the reaction time using PL characteristics. Furthermore, the actin structure transformation by cytochalasin D in cells, NIH-3T3s and HaCaTs, was also detected by the label-free PL spectrum, which includes the fluorescence of other biomaterials. The PL based determination in vivo was highly consistent with the result of dye-based fluorescence images.

Section snippets

Methods

Actin samples were prepared using the actin polymerization from G-actin and modified with regulatory proteins [9], [24]. G-actin, α-actinin and gelsolin were obtained from rabbit skeletal muscle and were purchased from Cytoskeleton Inc. Actin monomer was reconstituted at 20 μM in a fresh G-buffer (5 mM Tris-HCl, 0.2 mM CaCl2, 0.2 mM adenosine triphosphate (ATP), 0.01% (w/v) NaN3, pH 8.0) and incubated at 4 °C for 2 h. A tenth of the final volume of F-buffer (50 mM Tris-HCl, 500 mM KCl, 2 mM MgCl

Results and discussion

The morphology and structure of actin samples were observed using an SEM, as shown in Fig. 1. The length and thickness of the actin filaments were regulated with ABPs to mimic the various actins observed in cells. We prepared three types of F-actin with or without gelsolin to control the length of actin: intrinsic F-actin polymerized from G-actin without gelsolin (Fig. 1b) and two different volume ratios of G-actin to gelsolin, which were 2000:1 (Fig. 1c) and 200:1 (Fig. 1d), respectively. We

Conclusion

Using the label-free PL measurements, we were able to determine the alterations in length and thickness of protein assembly consisting of actins and actin binding proteins. We revealed that G-actin and polymerized actins exhibit their intrinsic PL characteristics with the specific peak position and intensity. As G-actin were assembled into a filament and filaments bundles, the PL emission peak shifted from 323 nm to 334 nm and its intensity increased significantly. The results indicate that the

Author contributions

B.P., J.L., H.L., and S.C.J. designed research; B.P., S.O., D.K., and S.J. performed experiments; B.P. and S.O. analyzed data; B.P., S.O., Y.J. H.L., and S.C.J. wrote the paper.

Acknowledgements

We thank F. Nakamura and for providing gelsolin proteins. This work was partially supported by Industrial Strategic Technology Development Program through the Ministry of Trade, Industry and Energy (MOTIE, Korea) (2MR4090), the National Research Foundation of Korea (NRF) (NRF-2015R1A5A1037668) funded by the Ministry of Education, Science and Technology (MEST), the ICT R&D program of MSIP/IITP (R0101-15-0034), the NRF (NRF-2015R1A2A2A01007602), Nano-Material Technology Development Program (

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