Monitoring Keap1-Nrf2 interactions in single live cells.

The transcription factor NF-E2 p45-related factor 2 (Nrf2) and its negative regulator Kelch-like ECH associated protein 1 (Keap1) control the expression of nearly 500 genes with diverse cytoprotective functions. Keap1, a substrate adaptor protein for Cullin3/Rbx1 ubiquitin ligase, normally continuously targets Nrf2 for degradation, but loses this ability in response to electrophiles and oxidants (termed inducers). Consequently, Nrf2 accumulates and activates transcription of its downstream target genes. Many inducers are phytochemicals, and cruciferous vegetables represent one of the richest sources of inducer activity among the most commonly used edible plants. Here we summarize the discovery of the isothiocyanate sulforaphane as a potent inducer which reacts with cysteine sensors of Keap1, leading to activation of Nrf2. We then describe the development of a quantitative Förster resonance energy transfer (FRET)-based methodology combined with multiphoton fluorescence lifetime imaging microscopy (FLIM) to investigate the interactions between Keap1 and Nrf2 in single live cells, and the effect of sulforaphane, and other cysteine-reactive inducers, on the dynamics of the Keap1-Nrf2 protein complex. We present the experimental evidence for the "cyclic sequential attachment and regeneration" or "conformation cycling" model of Keap1-mediated Nrf2 degradation. Finally, we discuss the implications of this mode of regulation of Nrf2 for achieving a fine balance under normal physiological conditions, and the consequences and mechanisms of disrupting this balance for tumor biology.

Regulatory flexibility in the Nrf2-mediated stress response is conferred by conformational cycling of the Keap1-Nrf2 protein complex.

The transcription factor NF-E2 p45-related factor 2 (Nrf2), a master regulator of cytoprotective genes, is controlled by dimeric Kelch-like ECH associated protein 1 (Keap1), a substrate adaptor protein for Cullin3/RING-box protein 1 ubiquitin ligase, which normally targets Nrf2 for ubiquitination and degradation but loses this ability in response to electrophiles and oxidants (inducers). By using recombinant proteins and populations of cells, some of the general features of the regulation of Nrf2 by Keap1 have been outlined. However, how the two proteins interact at a single-cell level is presently unknown. We now report the development of a quantitative Förster resonance energy transfer-based system using multiphoton fluorescence lifetime imaging microscopy and its application for investigating the interaction between Nrf2 and Keap1 in single live cells. By using this approach, we found that under homeostatic conditions, the interaction between Keap1 and Nrf2 follows a cycle in which the complex sequentially adopts two distinct conformations: "open," in which Nrf2 interacts with a single molecule of Keap1, followed by "closed," in which Nrf2 binds to both members of the Keap1 dimer. Inducers disrupt this cycle by causing accumulation of the complex in the closed conformation without release of Nrf2. As a consequence, free Keap1 is not regenerated, and newly synthesized Nrf2 is stabilized. On the basis of these findings, we propose a model we have named the "cyclic sequential attachment and regeneration model of Keap1-mediated degradation of Nrf2." This previously unanticipated dynamism allows rapid transcriptional responses to environmental changes and can accommodate multiple modes of regulation.

Sds22 regulates aurora B activity and microtubule-kinetochore interactions at mitosis.

We have studied Sds22, a conserved regulator of protein phosphatase 1 (PP1) activity, and determined its role in modulating the activity of aurora B kinase and kinetochore-microtubule interactions. Sds22 is required for proper progression through mitosis and localization of PP1 to mitotic kinetochores. Depletion of Sds22 increases aurora B T-loop phosphorylation and the rate of recovery from monastrol arrest. Phospho-aurora B accumulates at kinetochores in Sds22-depleted cells juxtaposed to critical kinetochore substrates. Sds22 modulates sister kinetochore distance and the interaction between Hec1 and the microtubule lattice and, thus, the activation of the spindle assembly checkpoint. These results demonstrate that Sds22 specifically defines PP1 function and localization in mitosis. Sds22 regulates PP1 targeting to the kinetochore, accumulation of phospho-aurora B, and force generation at the kinetochore-microtubule interface.

Quantitative analysis of chromatin compaction in living cells using FLIM-FRET.

We present a quantitative Förster resonance energy transfer (FRET)-based assay using multiphoton fluorescence lifetime imaging microscopy (FLIM) to measure chromatin compaction at the scale of nucleosomal arrays in live cells. The assay uses a human cell line coexpressing histone H2B tagged to either enhanced green fluorescent protein (FP) or mCherry FPs (HeLa(H2B-2FP)). FRET occurs between FP-tagged histones on separate nucleosomes and is increased when chromatin compacts. Interphase cells consistently show three populations of chromatin with low, medium, or high FRET efficiency, reflecting spatially distinct regions with different levels of chromatin compaction. Treatment with inhibitors that either increase chromatin compaction (i.e., depletion of adenosine triphosphate) or decrease chromosome compaction (trichostatin A) results in a parallel increase or decrease in the FLIM-FRET signal. In mitosis, the assay showed variation in compaction level, as reflected by different FRET efficiency populations, throughout the length of all chromosomes, increasing to a maximum in late anaphase. These data are consistent with extensive higher order folding of chromatin fibers taking place during anaphase.

Identifying specific protein interaction partners using quantitative mass spectrometry and bead proteomes.

The identification of interaction partners in protein complexes is a major goal in cell biology. Here we present a reliable affinity purification strategy to identify specific interactors that combines quantitative SILAC-based mass spectrometry with characterization of common contaminants binding to affinity matrices (bead proteomes). This strategy can be applied to affinity purification of either tagged fusion protein complexes or endogenous protein complexes, illustrated here using the well-characterized SMN complex as a model. GFP is used as the tag of choice because it shows minimal nonspecific binding to mammalian cell proteins, can be quantitatively depleted from cell extracts, and allows the integration of biochemical protein interaction data with in vivo measurements using fluorescence microscopy. Proteins binding nonspecifically to the most commonly used affinity matrices were determined using quantitative mass spectrometry, revealing important differences that affect experimental design. These data provide a specificity filter to distinguish specific protein binding partners in both quantitative and nonquantitative pull-down and immunoprecipitation experiments.

Visualization of intracellular PP1 targeting through transiently and stably expressed fluorescent protein fusions.

Protein phosphatase 1 (PP1) is a ubiquitous serine/threonine phosphatase that regulates many cellular processes, including cell division, signaling, differentiation, and metabolism. It is expressed in mammalian cells as three closely related isoforms: alpha, beta/delta, and gamma1. These isoforms differ in their relative affinities for proteins, termed targeting subunits, that mediate their intracellular localization and substrate specificity. Because of the dynamic nature of these interactions, it is important to find experimental approaches that permit direct analyses of PP1 localization and PP1-targeting subunit interactions in live cells. When transiently or stably expressed as fluorescent protein (FP) fusions, the three isoforms are active phosphatases with distinct localization patterns and can interact with both endogenous and exogenous targeting subunits. Their changing spatio-temporal distributions can be monitored both throughout the cell cycle and following cellular perturbations by time-lapse fluorescence microscopy, and turnover rates of intracellular pools of the protein calculated by fluorescence recovery after photobleaching (FRAP). Interactions with targeting subunits can be visualized in vivo by fluorescence resonance energy transfer (FRET), using techniques such as sensitized emission, acceptor photobleaching, or fluorescence lifetime imaging.