Spatially resolved protein interaction maps

A functional understanding of complex biologicalprocesses requires spatially resolved quantification of protein functions, i.e. their activities or protein-protein interactions. Towards this goal microscopy-based approaches to acquire such information are employed, based on a variety of physical or optical principles. FCS and FCCS (fluorescence (cross) correlation spectroscopy, generally called fluorescence fluctuation spectroscopy) are confocal microscope-based methods that allow assessing transport and mobility properties as well as interactions of molecules (protein-protein, protein-nucleic acid, protein-compound, protein complex formation) in vitro and in living cells. FCS and FCCS measurements can be done using commercial equipment based on confocal microscopes. These permit single point measurements inside individual cells and enable hihgly sensitive maesuremnets of proteins expressed at endogenous levels (1). Using custom-bulidt microscopes FCS measuremntscanalsobe conductd in an imaging mode where diffusion and protien-proteininteractionscan be imaged with high spatial resolution (2). Besides our work on signlal transduction pathways, we also used FCCS in different other finished or ongoing projects, e.g. to study spindle checkpoint signaling (3) or the regulation of veiscle formation during endocytosis (4).

 

Imaging protein turnover

Cellular protein homeostasis ensures for each protein optimal levels at the location of its function within the organism. Protein homeostasis is governed by a series of feedback controlled regulatory circuits that encompass each level of the entire cellular machinery. While much of the regulatory networks underlying protein homeostasis have been deciphered, however, it becomes also more and more clear that the spatio- temporal organization of the cell is a major determinant of regulation of protein homeostasis. Nuclear organization, transport and localization of mRNA, protein translation and protein modification and degradation all constitute factors that regulate protein homeostasis in a manner influenced by the intracellular organization and the local context. Hence, one of the main current challenges is to move studies on protein homeostasis from a cell population level to the level of single cell studies with subcellular resolution. To achieve this, new methods are required that allow access to the parameter that determine protein homeostasis in a manner compatible with microscopy.

Another method we developed is based on a novel type of fluorescent timer protein that gives visual access to the age of a protein simultaneously with its abundance and localization (Khmelinskii et al., 2012). This reporter consists of a tandem fusion of a green and a red fluorescent protein that again can be fused to a protein of interest. The combination of a green fluorescent protein, which has a fast maturing chromophore with a red fluorescent protein with a slow maturing chromophore provides a mean to obtain not only the localization of a protein, but upon measurement of the red and green signal also information about the age of a pool of the protein. Therefore the tandem fluorescent protein timer reporter, termed tFT, constitutes a way to measure the dynamics of protein turnover in the cell, ideally with subcellular resolution. We validated the tFT approach using artificial substrates that target the reporter for degradation via the N-end rule pathway via the ubiquitin/proteasome system (UPS). Using a systemic assay with yeast synthetic gene arrays (SGA), we confirmed that ability of SGA to be used to map genome wide degradation pathways. We used quantitative microscopy to study the turnover of proteins with subcellular resolution and focused on the nuclear pore complex (NPC). This revealed a quantitative picture about NPC homeostasis and inheritance during mitosis. One of the outcomes of this study was the disproval of a hypothesis where retention of NPCs in mother cells was believed to underlie asymmetric segregation of a determinant of cellular senescence and aging. We found that NPCs are in fact partitioned during mitosis (Khmelinskii et al., 2012, and Khmelinskii et al., (2010) Nature 466:E1) and that alternative explanations can account for the retention of aging factors (Khmelinskii et al., 2011). Furthermore, tFT based in vivo measurements of GPCR signaling in migrating tissues have led to the validation of a new model for self-perpetuated morphogen gradients (collaboration with Darren Gilmour, EMBL) (Dona et al., 2013).

 

Functional analysis of protein degradation networks

Towards a comprehenisve analysis of protein degradation we have generated a tFT library where >4000 yeast protein are tagged C-terminally with the tFT tag. Thsi resources enables us tocreen for proteins that specifically changetheir turn over as a function of particular disturbance. Using this resource we identified (5) a degradation pathway that appears to be dedicated to remove proteins from the nucleus, either the inner nuclear membrane (INM) or the nuclear lumen, that accidentally entered this compartment. On one hand side that pathway is required to keep the nuclear lumen essentially free from the two transcription factors Stp1 and Stp2 in order to prevent unnecessary (and energy consuming) expression of amino acid transporters in situation where no external amino acids are present (6,7). The second function of the pathway appears to clear INM from proteins that accidentally entered this site of the cell, either because their are mistargeted, or because their are passively entering from the outer nuclear membrane/ER. The core component of this pathway is the Asi-complex, an E3 ubiquitin ligase composed from two membrane proteins Asi1 and Asi3 that each contain a C-terminal RING domain, and the auxiliary protein Asi2, another membrane protein. Using the tandem fluorescent timer (tFT) method (8,9) we identified a range of substrates for the Asi-complex and demonstrated that Asi-dependent degradation and the two classical ERAD pathways, the Doa10 and the Hrd1 dependent pathways that remove misfolded proteins from the ER are distinct pathways. Nevertheless, they share with the Asi-dependent pathway some of the auxiliary machinery, i.e. the E2 enzymes and components for membrane extraction (e.g. Cdc4810) and Ubx99 (unpublished results).

FigureAsi

Figure 1: The function of the Asi complex and other ER associated E3 ubiquitin ligases.(a) Three major E3-ubiquitin ligases do localize to membranes continuous with the ER membrane: Hrd1, Doa10 and Asi1/3. The Asi-complex seems to specifically function at the inner nuclear membrane (INM) where it removes mis-targeted proteins from the nucleoplasm and the INM.(b) Composition, localisation and degron-requirements for the three E3 ligases of (a). The two ERAD E3 ligases Doa10 and Hrd1 recognize substrates based on the localisation of the degradation mediating part of the protein (i.e. the degrons or the misfolded regions of the protein): cytoplasme, intramembrane or lumen, as indicated.

 

Methods in microscopy

Our work is highly demanding with respect to microscopic instrumentation. As it is often the case with advancedinstrumentation, the maintenance of optimally performing equipment is challenging and time consuming. Moreover, for microscopes, no simple method is available to assess the optimal performance of such instruments, and many labs miss the requiresd specialist knowledge and hence frequently run instruments that perform with less than 50% of the practically achievable performance of the system. Towards a solution of this problem, we developed a comprehensive software tool that guides non-specialist users rapidely and reproducibly through a comprehensive performance testing of a microscope. This software is published in Nature Methods (10) and is now used in many labs and microscope facilities thoughout the world and became a de facto standart to test microscope performance.

 

Future directions

Currently, the lab is working on a series of new projects that involve recently established technology (i.e. screening FCS, 2D-FCCS, tFT) to address important questions in the field of endocytosis, MAP kinase signaling, protein homeostasis (funded by SFP 1036), non-coding RNA biology and regulation of mitochondrial quality control in mammalian cells.

 

References

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  2. Capoulade J, Wachsmuth M, Hufnagel L, Knop M. Quantitative fluorescence imaging of protein diffusion and interaction in living cells. Nat Biotechnol 2011;29:835–9.
  3. Heinrich S, Geissen E-M, Kamenz J, Trautmann S, Widmer C, Drewe P, Knop M, Radde N, Hasenauer J, Hauf S. Determinants of robustness in spindle assembly checkpoint signalling. Nat Cell Biol 2013;15:1328–39.
  4. Boeke D, Trautmann S, Meurer M, Wachsmuth M, Godlee C, Knop M, kaksonen M. Quantification of cytosolic interactions identifies Ede1 oligomers as key organizers of endocytosis. Mol Syst Biol 2014;10:756.
  5. Khmelinskii A, Blaszczak E, Pantazopoulou M, Fischer B, Omnus DJ, Le Dez G, Brossard A, Gunnarsson A, Barry JD, Meurer M, Kirrmaier D, Boone C, et al. Protein quality control at the inner nuclear membrane. Nature 2014;516:410–3.
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  8. Khmelinskii A, Knop M. Analysis of protein dynamics with tandem fluorescent protein timers. Methods Mol Biol 2014;1174:195–210.
  9. Khmelinskii A, Keller PJ, Bartosik A, Meurer M, Barry JD, Mardin BR, Kaufmann A, Trautmann S, Wachsmuth M, Pereira G, Huber W, Schiebel E, et al. Tandem fluorescent protein timers for in vivo analysis of protein dynamics. Nat Biotechnol 2012;30:708–14.
  10. Theer P, Mongis C, Knop M. PSFj: know your fluorescence microscope. Nat Methods 2014;11:981–2.