Protein Folding Landscapes in the Living Cell - The Journal of

PERSPECTIVE pubs.acs.org/JPCL

Protein Folding Landscapes in the Living Cell Simon Ebbinghaus*,†,‡,§ and Martin Gruebele*,†,‡,z †

Department of Chemistry, ‡Center for the Physics of Living Cells, and zDepartment of Physics, and Center for Biophysics and Computational Biology, University of Illinois, Urbana, Illinois 61801, United States

ABSTRACT This Perspective highlights studies of protein structure, stability, and folding kinetics that can now be carried out in prokaryotic and eukaryotic cells. The general features of cooperative folding transitions and kinetics remain the same as those in vitro, but the folding free-energy landscape and local viscosity vary according to microenvironments in the cellular landscape. Experiments in vitro provide a basis for comparing crowding and chemical effects with experiments inside of the cell. Perhaps cells even evolved to post-translationally modulate protein folding and binding through chemical patterns or by crowding.

that we recently developed. Fast relaxation imaging (FReI) has allowed us to peer into cells and monitor protein stability, folding kinetics, and interactions from the subcellular level up to whole cell populations.8,10 Different environments in the cell and different cells within a population differ remarkably from one another and from buffer solutions. Protein Folding Landscapes in the Cell. Proteins fold on a multidimensional free-energy landscape.11 The population distribution on that landscape is exponentially sensitive to the free energy; therefore, a single pathway usually carries most of the folding population.12 As a result, one-dimensional simplifications such as two-state folding or sequential intermediates often adequately describe the folding process. However, protein folding is a very fast chemical reaction, occurring in microseconds to hours at room temperature; the ups and downs of the free energy landscape are very small compared to most chemical reactions. Thus, a small modulation of a protein's energy landscape (just a few kBT) could switch from one dominant path to another. The measured effects of solvation and crowding on folding thermodynamics and kinetics in vitro are large indeed and have been studied systematically.13-15 Much less is known about how protein stability is altered in the natural folding environment, the densely packed cell (see Gershenson and Gierasch16 for a recent review of the field). Even less is known about in vivo folding kinetics. After the initial folding event following ribosomal synthesis,17,18 proteins unfold and refold numerous times because protein stability is relatively low to allow for protein flexibility and function. A typical protein may fold and unfold thousands of times within a cell before its lifespan is up. Thus, in-cell folding studies are a timely extension of in vitro folding experiments. Challenges for Measuring In-Cell Protein Structure, Stability, and Dynamics. Major challenges of in-cell experiments include

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he inside of the cell is a complicated environment for a protein to navigate compared to the homogeneous solvent conditions of in vitro experiments. The cell is crowded by up to 300 mg of macromolecules per mL,1 creating a matrix with variable viscosity and microstructure. Crowding reduces the conformational entropy of proteins and can speed up the conformational search for the folded state2 despite increased viscosity in the cell. Small solutes, pH variations, or macromolecules perturb protein folding (in analogy to osmolytes and denaturants in vitro).3 Protein-protein binding also modulates a protein's folding landscape strongly.4 For example, chaperones like heat shock protein 70 (Hsp70) bind to proteins that are denatured by cellular stress and assist the refolding process.5

Different environments in the cell and different cells within a population differ remarkably from one another and from buffer solutions. Most biophysical techniques are restricted to in vitro environments ranging from the gas phase to buffer solutions to crystals at cryogenic temperature. A number of techniques, such as NMR and optical spectroscopy,6-8 are now being adapted to study protein structure, stability, and folding directly inside of living cells, while techniques such as mass spectrometry provide a snapshot of protein stability within cells through destructive measurement.9 New tools and methods are required to quantify the effects of the cell on proteins, building on the extensive insights gained from in vitro studies. Our goal is to provide a perspective on advances in four closely linked areas, in-cell protein structure, protein stability, folding kinetics, and future functional dynamics studies, with a particular highlight on a technique

r 2011 American Chemical Society

Received Date: December 23, 2010 Accepted Date: January 20, 2011 Published on Web Date: February 01, 2011

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DOI: 10.1021/jz101729z |J. Phys. Chem. Lett. 2011, 2, 314–319


λ repressor within E. coli greatly increases within bacteria grown in a hyperosmotic environment. Ignatova et al. have overcome the sample destruction problem. They used urea titration to measure the stability of cellular retinoic acid-binding protein I (CRABP I) directly in bacterial cells.7,23 The authors incorporated a tetra-cystein motif and measured FIAsH fluorescence, reporting on the fold and aggregation of the protein. Protein Kinetics in the Cell. Diffusion of whole proteins within the cytoplasm of a cell has been studied for many years by techniques such as photobleaching recovery or fluorescence anisotropy.29-31 Metabolic enzymes in the cell can have diffusion coefficients reduced by a factor greater than 10 compared to aqueous solution, while extraneous proteins such as green fluorescent protein (GFP) migrate much faster.31 These techniques measure translation of the whole protein on a micrometer scale within the cell or rotational diffusion of proteins. In contrast, the polypeptide chain motions that occur during protein folding could not be detected inside of living cells until recently. To enable such experiments, we developed fast relaxation imaging (FReI) to spatiotemporally resolve protein folding kinetics inside of a single living cell8 or to compare folding from cell to cell.10 FReI subjects a cell containing a FRET-labeled target protein to a small step upward or downward in temperature (Figure 1). As the protein unfolds or refolds toward a new equilibrium, its FRET efficiency is imaged throughout the cell as a function of time, yielding a movie of protein folding kinetics inside of the cell. We studied the structure, stability, and folding kinetics of a fluorescent phosphoglycerate kinase (PGK) fusion protein in mammalian cells, in aqueous buffer solution, and in an artificially crowded environment.32 PGK is a multistep folder, making it an ideal candidate for modulation of the various folding steps by the cell. We chose genetically encoded AcGFP1 (green fluorescence) and mCherry (red fluorescence) FRET labels and express the PGK fusion construct in U2OS bone tissue cancer cells (Figure 1), HeLa, and E. coli cells. Within the limitations of our current millisecond instrument response, upward and downward temperature steps, or any desired temperature profile, can be achieved with a programmable infrared laser pulse (λ = 2200 nm).8 Temperature modulation has also recently been used by Schoen et al. to study nucleic acid dynamics in live cells.33 Cellular water is heated efficiently and uniformly by mid-infrared wavelengths, without phototoxicity.8,34 The intracellular temperature profile can be monitored directly by exciting only the acceptor chromophore with a green laser. By measuring FRET efficiency at constant temperature, we can obtain low-resolution structural information about the protein. By collecting data at a sequence of temperatures, thermal protein denaturation curves can be obtained within the cell.

molecular specificity and adequate sensitivity. Until label-free techniques such as infrared or nonlinear Raman spectroscopy (currently applied to the quantitative analysis of tissues19,20) are able to distinguish a single protein type within one cell, labeling techniques remain the method of choice. Fluorescent labels have been widely used to detect proorster teins in living cells.21 Experiments based on FRET (F€ resonance energy transfer) can be utilized to study proteinprotein interactions as well as intramolecular protein dynamics. In FRET, a donor absorbs light and either transfers the energy to an acceptor yielding red-shifted fluorescence or emits directly if the acceptor is too far away. The FRET donor-acceptor length scale of a few nanometers is wellsuited for such experiments.22 In addition to chemically attached dyes and fluorescent non-natural amino acids, genetically encoded fluorescent proteins are particularly advantageous for in-cell experiments. They can be produced directly by the cell itself along with the target protein.8 On the downside, these probes modify the protein or its local environment, and attachment sites are limited. Another alternative consists of amino acid motifs that form fluorescent complexes with a probe molecule.23 The other challenge of in-cell experiments is the small number of proteins available within a single cell compared to bulk experiments. There is very little signal available per unit time and per cell volume (e.g., 50 μm 50 μm 5 μm for a typical mammalian cell from a HeLa cell line). This is a particular problem for NMR experiments, although labeling can help here also; overexpressed cellular proteins grown on isotopically labeled media allow experiments to be conducted, at least on cell ensembles.24-27 Fluorescence-based experiments have been able to reach volumes of a few cubic micrometers within the cytoplasm of a cell to study protein folding of as few as ∼1000 proteins.8 The development of better red and near-infrared fluorescent labels will reduce competition from cell autofluorescence and push this limit further toward single proteins. Protein Structure in the Cell. In-cell NMR techniques have been developed to directly calibrate the cellular protein structure to in vitro measurements. Cell-penetrating peptides are used to deliver isotope-labeled proteins to the cell and to detect proteins in mammalian cells.28 Increased hydrogen exchanges rates for ubiquitin have been detected in the intracellular environment, plausibly due to increased conformational heterogeneity.28 Fluorine-labeling NMR was further used to measure rotational diffusion of proteins inside of cells.26 Such measurements complement in vitro techniques including crystallography, small-angle X-ray scattering, or cryoelectronmicrography. Protein Stability in the Cell. The SUPREX method (stability of unpurified proteins from rates of H/D exchange) is one example of how a bioincompatible method has been adapted to provide information of a protein in its native environment. SUPREX monitors the distribution of folded and unfolded proteins within a cell at the instant the cell is deuteriumlabeled. The measurement is carried out by mass spectrometry after cell lysis to analyze cell contents with high sensitivity and specificity.9 Experiments by Ghememaghami et al. showed that the thermodynamic stability of monomeric


FReI yields a movie of protein folding kinetics inside of the cell.

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Figure 1. FReI experiment: At time zero, a programmable infrared laser steps the temperature of the cell and its surrounding aqueous medium uniformly (∼0.3 mm heating spot size, red schematic trace). The folding equilibrium of green donor-red acceptor labeled PGK in the cell suddenly shifts. The return to equilibrium is imaged on a fluorescence microscope in green (D = donor) and red (A = acceptor) fluorescence channels with millisecond time resolution (illustrated by a folded PGK protein at t = 0 s, unfolded at t = 15 s). A folding trace can be obtained from the D/A ratio or the difference of the D and A channels scaled for quantum yield (shown schematically by the blue “Protein Folding” trace). For example, during unfolding, the FRETefficiency decreases; therefore, green D fluorescence increases, and red A fluorescence decreases, as shown in the images of an actual cell during the temperature jump data collection. FRETstructural experiment: The orange double arrow schematically illustrates the hinge motion that brings the two halves of the enzymatic active site together.

By collecting data after a temperature jump, folding kinetics can be measured for each pixel (currently diffraction-limited resolution) imaged within the cell.8 A number of interesting features of in-cell folding have already emerged from our initial studies of PGK.8,10 We consider in turn protein structure, stability, and folding kinetics. PGK structure is affected by the cell. We find that the donorto-acceptor fluorescence ratio (D/A) is typically only ∼3 for the fusion construct in cells under folding conditions ( 0.45 s-1 and red < 0.4 s-1. on the right). Folding microenvironments within the cytoplasm have different folding rates. These microenvironments often have a filamentous


Figure 2. Modulation of the free energy ΔG (blue to red color code) and effective free energy of activation ΔG† (vertical grid, includes modulation of the viscosity-dependent prefactor of the rate coefficient) of PGK fusion protein within a U2OS cell. The freeenergy parameters are defined in the yellow box, showing a one-dimensional projection of the energy landscape. The measurement shown here is similar to the subpixel resolution measurement shown in Figure 5 of ref 8.

shape, although we cannot presently assign them to specific ultrastructure within the cytoplasm. There is also evidence that the functional form of the kinetics differs in different microenvironments (see supplement to ref 10). The origin could be homogeneous or inhomogeneous: the signal could be averaged over an ensemble of proteins that differ in their relaxation rate but not in mechanism, or proteins in different microenvironments might follow different folding paths and sample different intermediates. It will be very interesting to examine this problem with diffusion-kinetic network models and to study simple two-state folders to see whether they produce nonexponential kinetics inside of the cell. FReI also could be used to map out the time evolution of folding microenvironments. Figure 2 summarizes stability and rate variations of protein folding in a live cell.

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Not all cellular effects are dramatic. We screened a population of 30 mammalian cells by FReI and found that folding kinetics on average was slowed down only by a factor of 2 in the cells (4.1 s on average) with respect to that in the in vitro measurements.10 Compared to PGK diffusion,31 the average effect of the cell on folding rates is rather small. The diffusion coefficient on the nanometer scale where protein folding occurs is less affected by macromolecular crowding than the diffusion coefficient on the multimicrometer length scale of translational diffusion. From Folding to Functional or Dysfunctional Dynamics. Folding microenvironments could correspond to functional microenvironments. Ouyang et al.36 used intramolecular FRET to map the activity of the tyrosine kinase Src and of the GTPase Rac during directional cell migration and measured an activity gradient across the cytoplasm; Rac is highly active near the leading edge, whereas the Src activity is uniform throughout the cell. Functional variations likely exist on smaller length scales also. The most significant changes of protein structure and dynamics are expected for intrinsically disordered proteins (IDPs), which are often involved in signaling and binding.37 These proteins, like R-synuclein (involved in Parkinson's disease) or Huntingtin (involved in Huntington's disease), have a low content of secondary or tertiary structure when probed in vitro. Some IDPs may only be disordered when probed in vitro and gain structure in the cells or in highly crowded environments.25 In-cell methods such as FReI could shine more light on the extent to which such proteins must bind to fold in the cellular environment as opposed to simply being folded due to crowding and chemical interactions with the cellular matrix. The observed heterogeneity of protein folding kinetics and stability across a cell population and within cells shows that protein homeostasis encounters challenges within cells. Unfolded of misfolded proteins are the most extreme result of such challenges. It is now thought that misfolded proteins could become cytotoxic even at a soluble oligomer stage before fibrils or plaques are formed in the final stage of many protein misfolding diseases. To avoid misfolding and facilitate refolding, cells have evolved at least two mechanisms, one nonspecific and one specific. We found that PGK refolds with nearly 100% efficiency inside of a living cell following a short temperature step, unlike the partial irreversibility observed in simple in vitro crowding matrixes.8,32 It appears that cytoplasmic composition has evolved to provide a matrix conducive for protein refolding. Such a nonspecific capability of the cytosol takes a large load off the chaperone network.

Figure 3. Reversible binding of Hsp70 to unfolded PGK substrate in the presence of ATP monitored by the temperature-dependent equilibrium D/A fluorescence ratio (scaled to 1 at 22 °C). Significant chaperone response begins above 38 °C, tracking PGK unfolding (Tm(PGK mutant) ≈ 40 °C in ref 8). The inset schematically illustrates the onset of FRETas protein-protein interaction occurs.

be crucial for the onset of degenerative diseases in cells, especially neurons.39 In particular, heat shock proteins (HSPs) respond to increased amounts of misfolded protein. It remains unknown how the heat shock response cascade is initially triggered. In-cell thermodynamics and kinetics can be used to resolve the sequence of events as HSPs recognize and bind misfolded cellular proteins. Figure 3 shows the donoracceptor ratio for Hsp70 labeled with mCherry binding to PGK labeled with AcGFP1 as a function of cell temperature. Unfolding of the PGK construct between 38 and 45 °C for the mutant of PGK8 used for Figure 3 induces cellular chaperone binding and causes the D/A FRET ratio to decrease. Ignatova et al. have studied irreversible aggregation in the cell. The authors found that aggregation of a Huntingtin chimera proceeds via a more direct pathway toward amyloid formation than that in vitro.40 By providing temperature steps of varying length and intensity, FReI could be used to measure how reversibility of aggregation depends on the length and intensity of the stress. In summary, a whole complement of techniques is coming on line for the physicochemical study of protein structure, stability, folding, and function inside of cells. Cellular proteins have coevolved with the cells that contain them, and a comparison of in vitro and in-cell results will show to what extent living organisms have exploited the modulation of protein free-energy landscapes by the cell.

AUTHOR INFORMATION

It appears that cytoplasmic composition has evolved to provide a matrix conducive for protein refolding.

Corresponding Author: *To whom correspondence should be addressed. E-mail: mgruebel@ illinois.edu.

Notes The chaperone network provides a specific mechanism for protein recovery.38 Failure of the chaperone machinery could


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E-mail: [email protected].



Biographies

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Simon Ebbinghaus (www.rub.de/pc2/ebbinghaus) worked as a von Humboldt and CPLC postdoctoral fellow with Martin Gruebele to develop the FReI technique. He started his own lab at the RuhrUniversity Bochum (Germany) in 2010, funded by the Ministry of Innovation, Science, Research and Technology NRW, and was elected as a junior fellow of the Academy of Science and Arts NRW. Martin Gruebele (www.scs.illinois.edu/mgweb) studied ions of astrochemical interest as a graduate student, did his postdoc in femtochemistry, and is currently James R. Eiszner Professor of Chemistry, Professor of Physics, and Professor of Biophysics and Chemical Biology at the University of Illinois. His research interests include biomolecule dynamics in vivo and in vitro, molecular energy flow, and nanostructure dynamics.

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ACKNOWLEDGMENT We acknowledge funding from the

National Science Foundation (MCB 1019958). S.E. was also supported by the Alexander von Humboldt Foundation and by the NSF Center for Physics of Living Cells (CPLC, Illinois Physics Department). Assistance with the experiments was provided by Apratim Dhar and Steffen B€ uning.

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Protein Folding Landscapes in the Living Cell - The Journal of

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