Biochemistry

Membraneless organelles, such as germ granules and stress granules, are liquid-like condensates formed by phase transition. Recently, we and others have adopted proximity-based labeling methods to determine the composition of these membraneless compartments. Here, we describe the use of TurboID—an engineered promiscuous biotin ligase—to label and purify proteins localizing to Caenorhabditis elegans germ granules, known as P granules. We provide a detailed protocol for visualization of the subcellular localization of biotinylated proteins from dissected gonads, assessment of TurboID enrichment using streptavidin blots, and enrichment of biotinylated proteins under stringent conditions. Altogether, this protocol provides a workflow to unravel the proteome of C. elegans germ granules. Importantly, the assays described here can be applied to interrogate many membraneless organelles, in a diversity of living multicellular organisms.

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Site-directed spin labeling in conjunction with electron paramagnetic resonance (EPR) is an attractive approach to measure residue specific dynamics and point-to-point distance distributions in a biomolecule. Here, we focus on the labeling of proteins with a Cu(II)-nitrilotriacetic acid (NTA) complex, by exploiting two strategically placed histidine residues (called the dHis motif). This labeling strategy has emerged as a means to overcome key limitations of many spin labels. Through utilizing the dHis motif, Cu(II)NTA rigidly binds to a protein without depending on cysteine residues. This protocol outlines three major points: the synthesis of the Cu(II)NTA complex; the measurement of continuous wave and pulsed EPR spectra, to verify a successful synthesis, as well as successful protein labeling; and utilizing Cu(II)NTA labeled proteins, to measure distance constraints and backbone dynamics. In doing so, EPR measurements are less influenced by sidechain motion, which influences the breadth of the measured distance distributions between two spins, as well as the measured residue-specific dynamics. More broadly, such EPR-based distance measurements provide unique structural constraints for integrative structural biophysics and complement traditional biophysical techniques, such as NMR, cryo-EM, FRET, and crystallography.

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Monitoring the success of Cu(II)NTA labeling.

Inteins garner significant interest from both basic and applied researchers due to their unique catalytic abilities. Herein, we describe a protocol for accurately monitoring protein splicing without purification using in-gel fluorescence immediately following Tris-Glycine SDS-PAGE. Following expression in Escherichia coli, cells are lysed by sonication, cell supernatants are separated using Tris-Glycine SDS-PAGE, and superfolder GFP (sfGFP) fluorescence is directly visualized within gels. This method is rapid, with sfGFP immediately imaged following SDS-PAGE without staining. Further, sfGFP can be specifically detected in complex samples such as E. coli cell supernatants, proteins run at expected masses, and all steps can be performed at ambient temperature. This strategy is broadly applicable beyond the study of protein splicing and can be used for sensitive and specific visualization of superfolder sfGFP-tagged proteins in-gel.

Proximity-based protein labeling has been developed to identify protein-nucleic acid interactions. We have reported a novel method termed CRUIS (CRISPR-based RNA-United Interacting System), which captures RNA-protein interactions in living cells by combining the RNA-binding capacity of CRISPR/Cas13 and the proximity-tagging activity of PUP-IT. Enzymatically deactivated Cas13a (dCas13a) is fused to the proximity labeling enzyme PafA. In the presence of a guide RNA, dCas13a binds specific target RNA region, while the fused PafA mediates the labeling of biotin-tagged Pup on proximal proteins. The labeled proteins can be enriched by streptavidin pull-down and identified by mass spectrometry. Here we describe the general procedure for capturing RNA-protein interactions using this method.

The acrosome reaction is a highly regulated exocytotic event that primes spermatozoa for successful fertilization. Upon induction, acrosomal exocytosis proceeds via a wave of vesiculation that radiates across the sperm head, destabilizing the acrosomal vesicle and resulting in the release of the acrosomal contents. Having shed their acrosome, spermatozoa are then capable of penetrating the outer vestments of the oocyte and initiating fertilization. Accordingly, the failure of spermatozoa to complete an acrosome reaction represents a relatively common etiology in male infertility patients, and the ability to induce acrosomal exocytosis has found clinical utility in the evaluation of sperm fertilizing capacity. Here, we firstly describe protocols for driving the capacitation of human spermatozoa in vitro using chemically defined media in order to prime the cells for completion of acrosomal exocytosis. We then describe methodology routinely used for the induction of acrosomal exocytosis incorporating either a physiological agonist (i.e., the steroidal hormone, progesterone) or pharmacological reagent (i.e., the divalent cation ionophore, A23187). Finally, we describe the application of histochemical and immunofluorescence techniques that can be applied to study the completion of the acrosome reaction. Such protocols have important diagnostic utility for sperm function testing in both clinical and andrological research laboratories.

Advances in fluorescence microscopy (FM), electron microscopy (EM), and correlative light and EM (CLEM) offer unprecedented opportunities for studying diverse proteins and nanostructures involved in fundamental cell biology. It is now possible to visualize and quantify the spatial organization of cellular proteins and other macromolecules by FM, EM, and CLEM. However, tagging and tracking cellular proteins across size scales is restricted by the scarcity of methods for attaching appropriate reporter chemistries to target proteins. Namely, there are few genetic tags compatible with EM. To overcome these issues we developed Versatile Interacting Peptide (VIP) tags, genetically-encoded peptide tags that can be used to image proteins by fluorescence and EM. VIPER, a VIP tag, can be used to label cellular proteins with bright, photo-stable fluorophores for FM or electron-dense nanoparticles for EM. In this Bio-Protocol, we provide an instructional guide for implementing VIPER for imaging a cell-surface receptor by CLEM. This protocol is complemented by two other Bio-Protocols outlining the use of VIPER (Doh et al., 2019a and 2019b).

Genetically-encoded tags are useful tools for multicolor and multi-scale cellular imaging. Versatile Interacting Peptide (VIP) tags, such as VIPER, are new genetically-encoded tags that can be used in various imaging applications. VIP tags consist of a coiled-coil heterodimer, with one peptide serving as the genetic tag and the other (“probe peptide”) delivering a reporter compatible with imaging. Heterodimer formation is rapid and specific, allowing proteins to be selectively labeled for live-cell and fixed-cell imaging. In this Bio-Protocol, we include a detailed guide for implementing the VIPER technology for imaging receptors on live cells and intracellular targets in fixed cells. This protocol is complemented by two other Bio-Protocols outlining the use of VIPER (Doh et al., 2019a and 2019b).

Versatile Interacting Peptide (VIP) tags are a new class of genetically-encoded tag designed for imaging cellular proteins by fluorescence and electron microscopy. In 2018, we reported the VIPER tag (Doh et al., 2018), which contains two elements: a genetically-encoded peptide tag (i.e., CoilE) and a probe peptide (i.e., CoilR). These two peptides deliver contrast to a protein of interest by forming a specific, high-affinity heterodimer. The probe peptide was designed with a single cysteine residue for site-specific modification via thiol-maleimide chemistry. This feature can be used to attach a variety of biophysical reporters to the peptide, including bright fluorophores for fluorescence microscopy or electron-dense nanoparticles for electron microscopy. In this Bio-Protocol, we describe our methods for expressing and purifying recombinant CoilR. Additionally, we describe protocols for making fluorescent or biotinylated probe peptides for labeling CoilE-tagged cellular proteins. This protocol is complemented by two other Bio-Protocols outlining the use of VIPER (Doh et al., 2019a and 2019b).

The architecture of quinone/inhibitor-access channel in proton-translocating NADH-quinone oxidoreductase (respiratory complex I) was modeled by X-ray crystallography and cryo-EM, however, it remains debatable whether the channel model reflects the physiologically relevant state present throughout the catalytic cycle. Using photoreactive [¹²⁵I]amilorides, we demonstrated that amiloride-type inhibitors bind to the interfacial region of multiple subunits (49-kDa, ND1, PSST, and 39-kDa subunits), which is difficult to reconcile with the current channel model. This report describes the procedures for photoaffinity labeling of bovine submitochondrial particles by photoreactive [¹²⁵I]amilorides. The protocol could be widely applicable for the characterization of various biologically active compounds, whose target protein remains to be identified or characterized.

Copper-catalyzed azide-alkyne-cycloaddition (CuAAC), also known as ‘click chemistry’ serves as a technique for bio-orthogonal, that is, bio-compatible labeling of macromolecules including proteins or lipids. Click chemistry has been widely used to covalently, selectively, and efficiently attach probes such as fluorophores or biotin to small bio-orthogonal chemical reporter groups introduced into macromolecules. In bio-orthogonal non-canonical amino acid tagging (BONCAT) and fluorescent non-canonical amino acid tagging (FUNCAT) proteins are metabolically labeled with a non-canonical, azide-bearing amino acid and subsequently CuAAC-clicked either to an alkyne-bearing biotin (BONCAT) for protein purification, Western blot, or mass spectrometry analyses or to an alkyne-bearing fluorophore (FUNCAT) for immunohistochemistry. In combination with mass spectrometry, these kinds of labeling and tagging strategies are a suitable option to identify and characterize specific proteomes in living organisms without the need of prior cell sorting. Here, we provide detailed protocols for FUNCAT and BONCAT click chemistry and the detection of tagged de novo synthesized proteins in Drosophila melanogaster.