Measuring Protein Dynamics Research Articles

Probing protein structural dynamics at the single-molecule level by nanopore tweezers.

Proteins are dynamic entities that sample multiple conformations over a range of time scales. Understanding how proteins work requires a full description of their conformational dynamics over time. Nuclear magnetic resonance (NMR) spectroscopy can measure protein dynamics at multiple time scales, however, application is laborious and limited to relatively small proteins that can be stably prepared at high concentrations. Conformational heterogeneity, such as multiple weakly populated conformational states, is challenging to resolve from ensemble methods such as NMR due to averaging effects. Integrative approaches and single-molecule techniques can be advantageous for revealing transient protein dynamics and functional heterogeneities. Nanopore tweezers have emerged as a powerful, single-molecule analytical tool to monitor the protein's conformational dynamics. Nanopore tweezers confine a single protein within the pore lumen and measure ionic current changes induced by the protein analyte's motion. Here we will present several studies of ClyA nanopore tweezers to showcase their utility in resolving ligand interaction modes, screening allosteric drugs and investigating intrinsically disordered protein interactions.

Challenges and Opportunities for Characterizing the Assembly of Nuclear Envelope Proteins by Fluorescence Fluctuation Spectroscopy

The nuclear envelope (NE), consisting of two membranes separated by a thin perinuclear space, has proven to be a challenging environment for measuring protein dynamics in living cells. First, exogenously expressed, fluorescently tagged proteins must be properly localized to the NE. Furthermore, undulations of the nuclear membranes complicate the application of fluorescence fluctuation spectroscopy (FFS) to proteins located within the perinuclear space by adding a slowly fluctuating signal. While this signal is absent for proteins associated with the nuclear membranes, distinguishing soluble from membrane-associated protein complexes by FFS presents a significant challenge since the diffusion time depends strongly on molecular size within the perinuclear space. Some proteins, such as the AAA+ ATPase TorsinA, may exist in both a membrane-associated and soluble form depending on the conditions in the cell. Determining the extent to which a protein associates with the membrane can provide valuable insight into its function within the cell. While diffusion time alone is insufficient to differentiate these populations, we explore whether the dependence of diffusion time on temperature can provide insight into a protein's localization within the NE. In addition, we explore how spatial correlations can be used to study the dynamics of proteins within the NE and the membrane undulations we previously reported. Finally, z-scan FFS provides a means to quantify the localization of proteins to the NE. These examples highlight the ways in which FFS techniques can be used to overcome the challenges of quantifying protein behavior in the NE of living cells. This work has been supported by a grant from the National Institutes of Health (R01 GM64589).

Real-Time De Novo Deposition of Centromeric Histone-Associated Proteins Using the Auxin-Inducible Degradation System.

Measuring protein dynamics is essential to uncover protein function and to understand the formation of large protein complexes such as centromeres. Recently, genome engineering in human cells has improved our ability to study the function of endogenous proteins. By combining genome editing techniques with the auxin-inducible degradation (AID) system, we created a versatile tool to study protein dynamics. This system allows us to analyze both protein function and dynamics by enabling rapid protein depletion and reexpression in the same experimental setup. Here, we focus on the dynamics of the centromeric histone-associated protein CENP-C, responsible for the formation of the kinetochore complex. Following rapid removal and reactivation of a fluorescent version of CENP-C by auxin treatment and removal, we could follow CENP-C de novo deposition at centromeric regions during different stages of the cell cycle. In conclusion, the auxin degradation system is a powerful tool to assess and quantify protein dynamics in real time.

Heterogeneity of the Nuclear Environment Investigated by Superresolution Microscopy and Fluorescence Correlation Spectroscopy

Genomes are more than one-dimensional entities purely defined by their linear DNA sequences [Misteli T, Cell (2007)]. A long standing challenge in biology is to decipher the principles of organization of what can be considered, by analogy with human-created libraries, the cell's primary unit of information storage and retrieval. In this respect, the development of super-resolved fluorescence microscopy has provided a new toolbox to peer into the nucleus. For instance, super-resolution microscopy has been recently applied to the investigation of nanoscale chromatin architecture, revealing nucleosome higher-order organization into heterogeneous ‘clutches’ and epigenetically dependent folding motives [Ricci M et al, Cell, (2015); Boettiger A et al, Nature (2016)].However, dynamic properties of proteins in the nucleus are also critical for their function [Misteli T, Cell (2007)]. Fluorescence Correlation Spectroscopy (FCS) has been used to map the dynamics of several proteins in the nucleus with diffraction-limited spatial resolution. Here we combine a novel STED-based super-resolution method [Lanzano’ L et al, Nat Commun (2015)] with FCS to measure protein dynamics in the nucleus with an improved spatial resolution of about 100 nm. We sample several positions within the nucleus by performing line scanning. The measured spatial and temporal heterogeneity of the dynamics, quantified by a recently introduced algorithm [Scipioni L et al, Biophys J (2016)], is discussed in relation to nuclear organization.

Light Sheet Microscopy to Measure Protein Dynamics.

Visualizing protein dynamics is the key to a quantitative understanding of molecular mechanisms in biological systems. Recent developments in fluorescent microscopy techniques allow for novel experimental approaches to study stochastic localization, distribution, and movement of fluorescently labeled molecules in high resolution as they occur over time in the context of living cells and whole organisms. Particularly suitable for such studies is the application of light sheet microscopy, as it enables rapid in vivo imaging of a wide range and size of specimen, from whole organisms and organs, down to the dynamics of subcellular structures and single molecules. This article summarizes the principles of light sheet microscopy and its advantages over other optical imaging techniques, such as light microscopy and confocal microscopy, and highlights recent innovations that significantly enhance spatio-temporal resolution. Also, this manuscript contains basic guidelines for the implementation of light sheet microscopy in the laboratory and presents thus far unpublished light sheet microscopy applications demonstrating the ability of this technology to measure protein dynamics in a whole-organism context. J. Cell. Physiol. 232: 27-35, 2017. © 2016 Wiley Periodicals, Inc.

Rotameric preferences of a protein spin label at edge‐strand β‐sheet sites

Protein spin labeling to yield the nitroxide-based R1 side chain is a powerful method to measure protein dynamics and structure by electron spin resonance. However, R1 measurements are complicated by the flexibility of the side chain. While analysis approaches for solvent-exposed α-helical environment have been developed to partially account for flexibility, similar work in β-sheets is lacking. The goal of this study is to provide the first essential steps for understanding the conformational preferences of R1 within edge β-strands using X-ray crystallography and double electron electron resonance (DEER) distance measurements. Crystal structures yielded seven rotamers for a non-hydrogen-bonded site and three rotamers for a hydrogen-bonded site. The observed rotamers indicate contextual differences in R1 conformational preferences compared to other solvent-exposed environments. For the DEER measurements, each strand site was paired with the same α-helical site elsewhere on the protein. The most probable distance observed by DEER is rationalized based on the rotamers observed in the crystal structure. Additionally, the appropriateness of common molecular modeling methods that account for R1 conformational preferences are assessed for the β-sheet environment. These results show that interpretation of R1 behavior in β-sheets is difficult and indicate further development is needed for these computational methods to correctly relate DEER distances to protein structure at edge β-strand sites.

Tracking molecular dynamics without tracking: image correlation of photo-activation microscopy

Measuring protein dynamics in the plasma membrane can provide insights into the mechanisms of receptor signaling and other cellular functions. To quantify protein dynamics on the single molecule level over the entire cell surface, sophisticated approaches such as single particle tracking (SPT), photo-activation localization microscopy (PALM) and fluctuation-based analysis have been developed. However, analyzing molecular dynamics of fluorescent particles with intermittent excitation and low signal-to-noise ratio present at high densities has remained a challenge. We overcame this problem by applying spatio-temporal image correlation spectroscopy (STICS) analysis to photo-activated (PA) microscopy time series. In order to determine under which imaging conditions this approach is valid, we simulated PA images of diffusing particles in a homogeneous environment and varied photo-activation, reversible blinking and irreversible photo-bleaching rates. Further, we simulated data with high particle densities that populated mobile objects (such as adhesions and vesicles) that often interfere with STICS and fluctuation-based analysis. We demonstrated in experimental measurements that the diffusion coefficient of the epidermal growth factor receptor (EGFR) fused to PAGFP in live COS-7 cells can be determined in the plasma membrane and revealed differences in the time-dependent diffusion maps between wild-type and mutant Lck in activated T cells. In summary, we have developed a new analysis approach for live cell photo-activation microscopy data based on image correlation spectroscopy to quantify the spatio-temporal dynamics of single proteins.

Dynamic Proteomics: In Vivo Proteome-Wide Measurement of Protein Kinetics Using Metabolic Labeling.

Control of biosynthetic and catabolic rates of polymers, including proteins, stands at the center of phenotype, physiologic adaptation, and disease pathogenesis. Advances in stable isotope-labeling concepts and mass spectrometric instrumentation now allow accurate in vivo measurement of protein synthesis and turnover rates, both for targeted proteins and for unbiased screening across the proteome. We describe here the underlying principles and operational protocols for measuring protein dynamics, focusing on metabolic labeling with (2)H2O (heavy water) combined with tandem mass spectrometric analysis of mass isotopomer abundances in trypsin-generated peptides. The core principles of combinatorial analysis (mass isotopomer distribution analysis or MIDA) are reviewed in detail, including practical advantages, limitations, and technical procedures to ensure optimal kinetic results. Technical factors include heavy water labeling protocols, optimal duration of labeling, clean up and simplification of sample matrices, accurate quantitation of mass isotopomer abundances in peptides, criteria for adequacy of mass spectrometric abundance measurements, and calculation algorithms. Some applications are described, including the noninvasive "virtual biopsy" strategy for measuring molecular flux rates in tissues through measurements in body fluids. In addition, application of heavy water labeling to measure flux lipidomics is noted. In summary, the combination of stable isotope labeling, particularly from (2)H2O, with tandem mass spectrometric analysis of mass isotopomer abundances in peptides, provides a powerful approach for characterizing the dynamics of proteins across the global proteome. Many applications in research and clinical medicine have been achieved and many others can be envisioned.

Quantitative measurement of intracellular protein dynamics using photobleaching or photoactivation of fluorescent proteins.

Unlike in vitro protein dynamics, intracellular protein dynamics are intricately regulated by protein-protein interactions or interactions between proteins and other cellular components, including nucleic acids, the plasma membrane and the cytoskeleton. Alteration of these dynamics plays a crucial role in physiological phenomena such as gene expression and cell division. Live-cell imaging via microscopy with the inherent properties of fluorescent proteins, i.e. photobleaching and photoconversion, or fluorescence correlation spectroscopy, provides insight into the movement of proteins and their interactions with cellular components. This article reviews techniques based on photo-induced changes in the physicochemical properties of fluorescent proteins to measure protein dynamics inside living cells, and it also discusses the strengths and weaknesses of these techniques.

Measuring protein dynamics in live cells: protocols and practical considerations for fluorescence fluctuation microscopy.

Quantitative analysis of protein complex stoichiometries and mobilities are critical for elucidating the mechanisms that regulate cellular pathways. Fluorescence fluctuation spectroscopy (FFS) techniques can measure protein dynamics, such as diffusion coefficients and formation of complexes, with extraordinary precision and sensitivity. Complete calibration and characterization of the microscope instrument is necessary in order to avoid artifacts during data acquisition and to capitalize on the full capabilities of FFS techniques. We provide an overview of the theory behind FFS techniques, discuss calibration procedures, provide protocols, and give practical considerations for performing FFS experiments. One important parameter recovered from FFS measurements is the relative molecular brightness that can correlate with oligomerization. Three methods for measuring molecular brightness (fluorescence correlation spectroscopy, photon-counting histogram, and number and brightness analysis) recover similar values when measuring samples under ideal conditions in vitro. However, examples are given illustrating that these different methods used for calculating molecular brightness of fluorescent molecules in cells are not always equivalent. Methods relying on spot measurements are more prone to bleaching and movement artifacts that can lead to underestimation of brightness values. We advocate for the use of multiple FFS techniques to study molecular brightnesses to overcome and compliment limitations of individual techniques.

Photoswitching-free FRAP analysis with a genetically encoded fluorescent tag.

Fluorescence recovery after photobleaching (FRAP) is a widely used imaging technique for measuring protein dynamics in live cells that has provided many important biological insights. Although FRAP presumes that the conversion of a fluorophore from a bright to a dark state is irreversible, GFP as well as other genetically encoded fluorescent proteins now in common use can also exhibit a reversible conversion known as photoswitching. Various studies have shown how photoswitching can cause at least four different artifacts in FRAP, leading to false conclusions about various biological phenomena, including the erroneous identification of anomalous diffusion or the overestimation of the freely diffusible fraction of a cellular protein. Unfortunately, identifying and then correcting these artifacts is difficult. Here we report a new characteristic of an organic fluorophore tetramethylrhodamine bound to the HaloTag protein (TMR-HaloTag), which like GFP can be genetically encoded, but which directly and simply overcomes the artifacts caused by photoswitching in FRAP. We show that TMR exhibits virtually no photoswitching in live cells under typical imaging conditions for FRAP. We also demonstrate that TMR eliminates all of the four reported photoswitching artifacts in FRAP. Finally, we apply this photoswitching-free FRAP with TMR to show that the chromatin decondensation following UV irradiation does not involve loss of nucleosomes from the damaged DNA. In sum, we demonstrate that the TMR Halo label provides a genetically encoded fluorescent tag very well suited for accurate FRAP experiments.

The Phylogenetics of Dynamics in DNA Clamp Proteins

Using HXMS to measure protein dynamics, Fang and colleagues (in this issue of Structure) probe the dynamics of processivity clamp proteins from all kingdoms of life. The proteins have similar structures butdivergent sequences. Their results show conserved dynamics correlating with primary functions and divergent dynamics for divergent functions.

Amplitudes and time scales of picosecond-to-microsecond motion in proteins studied by solid-state NMR: a critical evaluation of experimental approaches and application to crystalline ubiquitin

Solid-state NMR provides insight into protein motion over time scales ranging from picoseconds to seconds. While in solution state the methodology to measure protein dynamics is well established, there is currently no such consensus protocol for measuring dynamics in solids. In this article, we perform a detailed investigation of measurement protocols for fast motions, i.e. motions ranging from picoseconds to a few microseconds, which is the range covered by dipolar coupling and relaxation experiments. We perform a detailed theoretical investigation how dipolar couplings and relaxation data can provide information about amplitudes and time scales of local motion. We show that the measurement of dipolar couplings is crucial for obtaining accurate motional parameters, while systematic errors are found when only relaxation data are used. Based on this realization, we investigate how the REDOR experiment can provide such data in a very accurate manner. We identify that with accurate rf calibration, and explicit consideration of rf field inhomogeneities, one can obtain highly accurate absolute order parameters. We then perform joint model-free analyses of 6 relaxation data sets and dipolar couplings, based on previously existing, as well as new data sets on microcrystalline ubiquitin. We show that nanosecond motion can be detected primarily in loop regions, and compare solid-state data to solution-state relaxation and RDC analyses. The protocols investigated here will serve as a useful basis towards the establishment of a routine protocol for the characterization of ps–μs motions in proteins by solid-state NMR.Electronic supplementary materialThe online version of this article (doi:10.1007/s10858-013-9787-x) contains supplementary material, which is available to authorized users.

Recombination-Induced Tag Exchange (RITE) Cassette Series to Monitor Protein Dynamics in Saccharomyces cerevisiae

Proteins are not static entities. They are highly mobile, and their steady-state levels are achieved by a balance between ongoing synthesis and degradation. The dynamic properties of a protein can have important consequences for its function. For example, when a protein is degraded and replaced by a newly synthesized one, posttranslational modifications are lost and need to be reincorporated in the new molecules. Protein stability and mobility are also relevant for the duplication of macromolecular structures or organelles, which involves coordination of protein inheritance with the synthesis and assembly of newly synthesized proteins. To measure protein dynamics, we recently developed a genetic pulse-chase assay called recombination-induced tag exchange (RITE). RITE has been successfully used in Saccharomyces cerevisiae to measure turnover and inheritance of histone proteins, to study changes in posttranslational modifications on aging proteins, and to visualize the spatiotemporal inheritance of protein complexes and organelles in dividing cells. Here we describe a series of successful RITE cassettes that are designed for biochemical analyses, genomics studies, as well as single cell fluorescence applications. Importantly, the genetic nature and the stability of the tag switch offer the unique possibility to combine RITE with high-throughput screening for protein dynamics mutants and mechanisms. The RITE cassettes are widely applicable, modular by design, and can therefore be easily adapted for use in other cell types or organisms.

The Molecular Basis for Cellular Contractility Driven Mechanosensing

Internal and external mechanical forces govern many cellular processes in normal and diseased states such as muscle contraction, tissue invasion and metastasis of tumor cells. Thus a cell's ability to sense and respond to mechanical stimuli is critical for cell behavior and survival. In Dictyostelium, the mechanoenzyme myosin II and the actin crosslinker cortexillin I work cooperatively to control cellular contractility in response to deformation. The scaffolding proteins IQGAP1 and IQGAP2, which can bind to cortexillin I, further regulate this mechanosensory system through feedback loops. The IQGAPs are not required for mechanosensing, but affect the stress-dependent myosin accumulation in the cortex. IQGAP1 inhibits mechanosensing, while IQGAP2 is required for suppressing this inhibition. A molecular understanding of how this mechanosensory system works to modulate contractility remains elusive. By measuring protein dynamics using fluorescence recovery after photobleaching (FRAP), we observed stress-dependent reduction in the mobility of many of the proteins in the mechanosensory system. Further, these dynamics are dependent on the integrity of the mechanosensory system, as mutant cells with disrupted mechanosensing typically show slower protein dynamics and increased cortical association of the proteins than in wild type cells. In addition, we are also characterizing the biochemical interactions important for mechanosensory response. These experiments will allow us to develop a molecular model to explain how mechanical forces are transmitted within the cell. The knowledge of the molecular underpinnings of this myosin II-based feedback control system will be applicable to other contractile systems such as those involved in development and morphogenesis of multicellular organisms.

Deep-UV Resonance Raman Analysis of the Rhodobacter capsulatus Cytochrome bc1 Complex Reveals a Potential Marker for the Transmembrane Peptide Backbone

Classical strategies for structure analysis of proteins interacting with a lipid phase typically correlate ensemble secondary structure content measurements with changes in the spectroscopic responses of localized aromatic residues or reporter molecules to map regional solvent environments. Deep-UV resonance Raman (DUVRR) spectroscopy probes the vibrational modes of the peptide backbone itself, is very sensitive to the ensemble secondary structures of a protein, and has been shown to be sensitive to the extent of solvent interaction with the peptide backbone [ Wang , Y. , Purrello , R. , Georgiou , S. , and Spiro , T. G. ( 1991 ) J. Am. Chem. Soc. 113 , 6368 - 6377 ]. Here we show that a large detergent solubilized membrane protein, the Rhodobacter capsulatus cytochrome bc(1) complex, has a distinct DUVRR spectrum versus that of an aqueous soluble protein with similar overall secondary structure content. Cross-section calculations of the amide vibrational modes indicate that the peptide backbone carbonyl stretching modes differ dramatically between these two proteins. Deuterium exchange experiments probing solvent accessibility confirm that the contribution of the backbone vibrational mode differences are derived from the lipid solubilized or transmembrane α-helical portion of the protein complex. These findings indicate that DUVRR is sensitive to both the hydration status of a protein's peptide backbone, regardless of primary sequence, and its secondary structure content. Therefore, DUVRR may be capable of simultaneously measuring protein dynamics and relative water/lipid solvation of the protein.

The Lives of Proteins

A new method to measure protein dynamics in living cells reveals how protein removal regulates cell physiology.

Small Cells—Big Future

Small Cells—Big Future

Measuring the turnover rates of Arabidopsis proteins using deuterium oxide: an auxin signaling case study

Rapid environmental responses in plants rely on endogenous signaling mechanisms, which in many cases are mediated by changes in protein turnover rates. It is therefore necessary to develop methods for measuring protein dynamics that monitor large sets of plant proteins to begin to apply a systems biology approach to the study of plant behavior. The use of stable isotope labeling strategies that are adaptable to proteomic methods is particularly attractive for this purpose. Here, we explore one example of such methods that is particularly suitable for plants at the seedling stage, where measurement of amino acid and protein turnover rates is accomplished using a heavy water labeling strategy. The method is backed by microarray evaluation to define its feasibility for specific experimental approaches, and the CULLIN-ASSOCIATED AND NEDDYLATION DISSOCIATED 1 (CAND1) and TRANSPORT INHIBITOR RESPONSE 1 (TIR1) proteins are used to illustrate the potential utility in understanding hormonal signaling regulation. These studies provide insight not only into the potential utility of the method, but also address possible areas of concern regarding the use of heavy water labeling during plant growth. These considerations suggest a prescription for specific experimental designs that minimize interference resulting from the induction of treatment-specific gene expression in the results obtained.

Photoactivatable GFP tagging cassettes for protein‐tracking studies in the budding yeast Saccharomyces cerevisiae

Yeast cell biologists use a variety of fluorescent protein tags for determining protein localization and for measuring protein dynamics using fluorescence recovery after photobleaching (FRAP). Although many modern fluorescent proteins, such as those with photoactivatable and photoconvertible characteristics, have been developed, none has been exploited for studies in budding yeast. We describe here the construction of yeast-tagging vectors containing photoactivatable green fluorescent protein (PA-GFP) for analysis of protein behaviour. We tagged two yeast proteins, Erg6p and Num1p, with PA-GFP and demonstrated specific photoactivation of the fusion proteins in live cells. Fluorescence intensity measurements showed that a short 5 s exposure to 413 nm light is sufficient to produce the maximum level of activated GFP fluorescence. Local photoactivation of cortical Num1p-PA-GFP showed movement of the marked proteins, providing new insights into the behaviour of Num1p at the cell cortex. Since photoactivation can be achieved using standard mercury arc illumination, the PA-GFP tag represents a convenient and economical way to determine protein dynamics in the cell. Thus, the tagging modules should facilitate protein-tracking studies in a wide variety of cell biological processes in yeast.