The timing of spikes dictates a neuron’s impact on downstream circuits and behavior, and spike timing is determined by the membrane potential (Vm). However, due to technical challenges, it has been impossible to analyze the relative timing of Vm dynamics between neurons during behavior. Using large scale membrane voltage imaging, we simultaneously recorded Vm from many individual hippocampal neurons in animals engaged in a virtual spatial task. We found that relative phase of Vm theta oscillations across neurons exhibit gradual or discrete shifts depending on spatial position. This finding extends beyond previous studies showing Vm dynamics in single neurons or spiking activity in multiple neurons, revealing previously unknown evidence for consistent coding of space by spike-independent relative phase of Vm theta dynamics between neurons.

Different spatial or temporal protein populations, such as cell-surface/intracellular or pre-existing/nascent subpopulations, determine the basal and activity-induced functions of individual synapses within a neuron in vivo. Here, we developed a simple and generalizable platform to image different spatial and temporal subpopulations of endogenous proteins at thousands of synapses in single neurons in the mammalian brain. The platform is based on the development, improvement and integration of CRISPR-Cas9-mediated protein labeling methods, chemical tag labeling techniques, and a semi-automatic analytical pipeline. The combined platform enables whole-cell mapping of total, cell-surface, intracellular, pre-existing, nascent or nascent-and-surface populations of endogenous proteins, such as receptor, scaffold and signaling proteins, at thousands of synapses in individual neurons in living or fixed mouse brain. Our single-cell "synaptome" mapping of endogenous protein subpopulations comprehensively visualizes the spatial representation of synapse diversity in protein localization, trafficking and turnover, providing valuable insights into single-cell organization and computations in the brain.

Enzyme-based self-labeling tags enable the covalent attachment of synthetic molecules to proteins inside living cells. A frontier of this field is designing cell-permeable multifunctional ligands that contain fluorophores in combination with affinity tags or pharmacological agents. This is challenging since attachment of additional chemical moieties onto fluorescent ligands can adversely affect membrane permeability. To address this problem, we examined the chemical properties of rhodamine-based self-labeling tag ligands through the lens of medicinal chemistry. We found that the lactone-zwitterion equilibrium constant () of rhodamines inversely correlates with their distribution coefficients (log), suggesting that ligands based on dyes exhibiting low and high log values, such as Si-rhodamines, would efficiently enter cells. We designed cell-permeable multifunctional HaloTag ligands with a biotin moiety to purify mitochondria or a JQ1 appendage to translocate BRD4 within the nucleus. We found that translocation of BRD4 to constitutive heterochromatin in cells leads to apparent increases in transcriptional activity. These fluorescent reagents enable affinity capture and translocation of intracellular proteins in living cells, and our general design concepts will facilitate the design of multifunctional chemical tools for biology.

Preprint: https://doi.org/10.1101/2022.07.02.498544 
Preprint: https://doi.org/10.32388/0xcyuc 

Differential Scanning Fluorimetry (DSF) is a biophysical assay that is used to estimate protein stability in vitro. In a DSF experiment, the increased fluorescence of a solvatochromatic dye, such as Sypro Orange, is used to detect the unfolding of a protein during heating. However, Sypro Orange is only compatible with a minority of proteins (< 30%), limiting the scope of this method. We recently reported that protein-adaptive DSF (paDSF) can partially solve this problem, wherein the protein is initially pre-screened against ∼300 chemically diverse dyes, termed the Aurora collection. While this approach significantly improves the number of targets amenable to DSF, it still fails to produce protein-dye pairs for some proteins. Here, we report the expansion of the dye collection to Aurora 2.0, which includes a total of 517 structurally diverse molecules and multiple new chemotypes. To assess performance, these dyes were screened against a panel of ∼100 proteins, which were selected, in part, to represent the most challenging targets (e.g. small size). From this effort, Aurora 2.0 achieved an impressive success rate of 94%, including producing dyes for some targets that were not matched in the original collection. These findings support the idea that larger, more chemically diverse libraries improve the likelihood of detecting melting transitions across a wider range of proteins. We propose that Aurora 2.0 makes paDSF an increasingly powerful method for studying protein stability, ligand binding and other biophysical properties in high throughput.

Fluorescent proteins have transformed biological imaging, yet their limited photostability and brightness restrict their applications. We used deep learning-based de novo protein design methods to design binders to three bright, stable and cell-permeable dyes spanning the imaging spectrum: JF657 (far red), JF596 (orange-red) and JF494 (green). We obtain highly specific dye-binding proteins with low nanomolar affinities for the intended target; a crystal structure of one binder confirms close resemblance to the design model. Simultaneous labeling of mammalian cells expressing three dye-specific binders at different subcellular compartments demonstrates the utility in multiplex imaging. We further expand the functionality of the binder by incorporating an active site that carries out nucleophilic aromatic substitution to form a covalent linkage with the dye, and develop split versions which reconstitute fluorescence at subcellular locations where both halves are present towards monitoring in-cell protein interactions and chemically induced dimerization. Our designed high affinity and specificity dye binders open up new opportunities for multiplexed biological imaging.

Neurochemical signals like dopamine (DA) play a crucial role in a variety of brain functions through intricate interactions with other neuromodulators and intracellular signaling pathways. However, studying these complex networks has been hindered by the challenge of detecting multiple neurochemicals in vivo simultaneously. To overcome this limitation, we developed a single-protein chemigenetic DA sensor, HaloDA1.0, which combines a cpHaloTag-chemical dye approach with the G protein-coupled receptor activation-based (GRAB) strategy, providing high sensitivity for DA, sub-second response kinetics, and an extensive spectral range from far-red to near-infrared. When used together with existing green and red fluorescent neuromodulator sensors, Ca2+ indicators, cAMP sensors, and optogenetic tools, HaloDA1.0 provides high versatility for multiplex imaging in cultured neurons, brain slices, and behaving animals, facilitating in-depth studies of dynamic neurochemical networks.

Voltage imaging is a promising technique for high-speed recording of neuronal population activity. However, tissue scattering severely limits its application in dense neuronal populations. Here, we adopted the principle of localization microscopy, a technique that enables super-resolution imaging of single-molecules, to resolve dense neuronal activities in vivo. Leveraging the sparse activation of neurons during action potentials (APs), we precisely localize the fluorescence change associated with each AP, creating a super-resolution image of neuronal activities. This approach, termed Activity Localization Imaging (ALI), identifies overlapping neurons and separates their activities with over 10-fold greater precision than what tissue scattering permits. Using ALI, we simultaneously recorded over a hundred densely-labeled CA1 neurons, creating a map of hippocampal theta oscillation at single-cell and single-cycle resolution.

Preprint: https://doi.org/10.1101/2023.12.03.56840

Intermediate filaments (IFs) play key roles in cellular mechanics, signaling, and organization, but tools for their rapid, selective disassembly remain limited. Here, we introduce FilaBuster, a photochemical approach for efficient and spatiotemporally controlled IF disassembly in living cells. FilaBuster uses a three-step strategy: (1) targeting HaloTag to IFs, (2) labeling with a covalent photosensitizer ligand, and (3) light-induced generation of localized reactive oxygen species to trigger filament disassembly. This modular strategy applies broadly across IF subtypes—including vimentin, GFAP, desmin, peripherin, and keratin 18—and is compatible with diverse dyes and imaging platforms. Using vimentin IFs as a model system, we establish a baseline implementation in which vimentin-HaloTag labeled with a photosensitizer HaloTag ligand triggers rapid and specific IF disassembly upon light activation. We then refine this approach by (i) expanding targeting strategies to include a vimentin nanobody-HaloTag fusion, (ii) broadening the range of effective photosensitizers, and (iii) optimizing irradiation parameters to enable precise spatial control over filament disassembly. Together, these findings position FilaBuster as a robust platform for acute, selective, and spatiotemporally precise disassembly of IF networks, enabling new investigations into their structural and functional roles in cell physiology and disease.

Synaptic plasticity alters neuronal connections in response to experience, which is thought to underlie learning and memory. However, the loci of learning-related synaptic plasticity, and the degree to which plasticity is localized or distributed, remain largely unknown. Here we describe a new method, DELTA, for mapping brain-wide changes in synaptic protein turnover with single-synapse resolution, based on Janelia Fluor dyes and HaloTag knock-in mice. During associative learning, the turnover of the ionotropic glutamate receptor subunit GluA2, an indicator of synaptic plasticity, was enhanced in several brain regions, most markedly hippocampal area CA1. More broadly distributed increases in the turnover of synaptic proteins were observed in response to environmental enrichment. In CA1, GluA2 stability was regulated in an input-specific manner, with more turnover in layers containing input from CA3 compared to entorhinal cortex. DELTA will facilitate exploration of the molecular and circuit basis of learning and memory and other forms of plasticity at scales ranging from single synapses to the entire brain.

A tool to map changes in synaptic strength during a defined time window could provide powerful insights into the mechanisms of learning and memory. Here we developed a technique, Extracellular Protein Surface Labeling in Neurons (EPSILON), to map α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR) exocytosis in vivo by sequential pulse-chase labeling of surface AMPARs with membrane-impermeable dyes. This approach yields synaptic-resolution maps of AMPAR exocytosis, a proxy for synaptic potentiation, in genetically targeted neurons during memory formation. In mice undergoing contextual fear conditioning, we investigated the relationship between synapse-level AMPAR exocytosis in CA1 pyramidal neurons and cell-level expression of the immediate early gene product cFos, a frequently used marker of engram neurons. We observed a strong correlation between AMPAR exocytosis and cFos expression, suggesting a synaptic mechanism for the association of cFos expression with memory engrams. The EPSILON technique is a useful tool for mapping synaptic plasticity and may be extended to investigate trafficking of other transmembrane proteins.