分子生物学

Understanding protein–protein interactions is crucial for unravelling subcellular protein distribution, contributing to our understanding of cellular organisation. Moreover, interaction studies can reveal insights into the mechanisms that cover protein trafficking within cells. Although various techniques such as Förster resonance energy transfer (FRET), co-immunoprecipitation, and fluorescence microscopy are commonly employed to detect protein interactions, their limitations have led to more advanced techniques such as the in situ proximity ligation assay (PLA) for spatial co-localisation analysis. The PLA technique, specifically employed in fixed cells and tissues, utilises species-specific secondary PLA probes linked to DNA oligonucleotides. When proteins are within 40 nm of each other, the DNA oligonucleotides on the probes interact, facilitating circular DNA formation through ligation. Rolling-circle amplification then produces DNA circles linked to the PLA probe. Fluorescently labelled oligonucleotides hybridise to the circles, generating detectable signals for precise co-localisation analysis. We employed PLA to examine the co-localisation of dynein with the Kv7.4 channel protein in isolated vascular smooth muscle cells from rat mesenteric arteries. This method enabled us to investigate whether Kv7.4 channels interact with dynein, thereby providing evidence of their retrograde transport by the microtubule network. Our findings illustrate that PLA is a valuable tool for studying potential novel protein interactions with dynein, and the quantifiable approach offers insights into whether these interactions are changed in disease.

Signaling pathways are involved in key cellular functions from embryonic development to pathological conditions, with a pivotal role in tissue homeostasis and transformation. Although most signaling pathways have been intensively examined, most studies have been carried out in murine models or simple cell culture. We describe the dissection of the TGF-β signaling pathway in human tissue using CRISPR-Cas9 genetically engineered human keratinocytes (N/TERT-1) in a 3D organotypic skin model combined with quantitative proteomics and phosphoproteomics mass spectrometry. The use of human 3D organotypic cultures and genetic engineering combined with quantitative proteomics and phosphoproteomics is a powerful tool providing insight into signaling pathways in a human setting. The methods are applicable to other gene targets and 3D cell and tissue models.

Key features

• 3D organotypic models with genetically engineered human cells.

• In-depth quantitative proteomics and phosphoproteomics in 2D cell culture.

• Careful handling of cell cultures is critical for the successful formation of theorganotypic cultures.

• For complete details on the use of this protocol, please refer to Ye et al. 2022.

As the most energy- and metabolite-consuming process, protein synthesis is under the control of several intrinsic and extrinsic factors that determine its fine-tuning to the cellular microenvironment. Consequently, variations in protein synthesis rates occur under various physiological and pathological conditions, enabling an adaptive response by the ce•ll. For example, global protein synthesis increases upon mitogenic factors to support biomass generation and cell proliferation, while exposure to low concentrations of oxygen or nutrients require translational repression and reprogramming to avoid energy depletion and cell death. To assess fluctuations in protein synthesis rates, radioactive isotopes or radiolabeled amino acids are often used. Although highly sensitive, these techniques involve the use of potentially toxic radioactive compounds and require specific materials and processes for the use and disposal of these molecules. The development of alternative, non-radioactive methods that can be easily and safely implemented in laboratories has therefore been encouraged to avoid handling radioactivity. In this context, the SUrface SEnsing of Translation (SUnSET) method, based on the classical western blot technique, was developed by Schmidt et al. in 2009. The SUnSET is nowadays recognized as a simple alternative to radioactive methods assessing protein synthesis rates.

Key features

• As a structural analogue of aminoacyl-transfer RNA, puromycin incorporates into the elongating peptide chain.

• Detection of puromycin-labeled peptides by western blotting reflects translation rates without the need for radioactive isotopes.

• The protocol described here for in vitro applications is derived from the SUnSET method originally published by Schmidt et al. (2009).

The auxin-inducible degron (AID) system is a versatile tool in cell biology and genetics, enabling conditional protein regulation through auxin-induced degradation. Integrating CRISPR/Cas9 with AID expedites tagging and depletion of a required protein in human and mouse cells. The mechanism of AID involves interactions between receptors like TIR1 and the AID tag fused to the target protein. The presence of auxin triggers protein ubiquitination, leading to proteasome-mediated degradation. We have used AID to explore the mitotic functions of the replication licensing protein CDT1. Swift CDT1 degradation via AID upon auxin addition achieves precise mitotic inhibition, revealing defects in mitotic spindle structure and chromosome misalignment. Using live imaging, we found that mitosis-specific degradation of CDT1 delayed progression and chromosome mis-segregation. AID-mediated CDT1 inhibition surpasses siRNA-based methods, offering a robust approach to probe CDT1’s mitotic roles. The advantages of AID include targeted degradation and temporal control, facilitating rapid induction and reversal of degradation—contrasting siRNA’s delayed RNA degradation and protein turnover. In summary, the AID technique enhances precision, control, and efficiency in studying protein function and regulation across diverse cellular contexts. In this article, we provide a step-by-step methodology for generating an efficient AID-tagging system, keeping in mind the important considerations that need to be adopted to use it for investigating or characterizing protein function in a temporally controlled manner.

Key features

• The auxin-inducible degron (AID) system serves as a versatile tool, enabling conditional protein regulation through auxin-induced degradation in cell biology and genetics.

• Integration of CRISPR/Cas9 knock-in technology with AID expedites the tagging and depletion of essential proteins in mammalian cells.

• AID’s application extends to exploring the mitotic functions of the replication licensing protein CDT1, achieving precise mitotic inhibition and revealing spindle defects and chromosome misalignment.

• The AID system and its diverse applications advance the understanding of protein function and cellular processes, contributing to the study of protein regulation and function.

Graphical overview

Cdt1–auxin-inducible degron (AID) tagging workflow. (A) Schematic of the cloned Cdt1 gRNA vector and the repair template generated to endogenously tag the Cdt1 genomic locus with YFP and AID at the C-terminal using CRISPR/CAS9-based genome editing. The two plasmids are transfected into DLD1-TIR1 stable cells, followed by sorting and scaling up of YFP-positive single cells. (B) The molecular mechanism of auxin-induced proteasome-mediated degradation of the target protein (CDT1) shown at the bottom of the figure is well worked out.

Human mitochondrial DNA (mtDNA) encodes several components of oxidative phosphorylation responsible for the bulk of cellular energy production. The mtDNA is transcribed by a dedicated human mitochondrial RNA polymerase (POLRMT) that is structurally distinct from its nuclear counterparts, instead closely resembling the single-subunit viral RNA polymerases (e.g., T7 RNA polymerase). The initiation of transcription by POLRMT is aided by two initiation factors: transcription factor A, mitochondrial (TFAM), and transcription factor B2, mitochondrial (TFB2M). Although many details of human mitochondrial transcription initiation have been elucidated with in vitro biochemical and structural studies, much remains to be addressed relating to the mechanism and regulation of transcription. Studies of such mechanisms require reliable, high-yield, and high-purity methods for protein production, and this protocol provides the level of detail and troubleshooting tips that are necessary for a novice to generate meaningful amounts of proteins for experimental work. The current protocol describes how to purify recombinant POLRMT, TFAM, and TFB2M from Escherichia coli using techniques such as affinity column chromatography (Ni²⁺ and heparin), how to remove the solubility tags with TEV protease and recover untagged proteins of interest, and how to overcome commonly encountered challenges in obtaining high yield of each protein.

Key features

• This protocol builds upon purification methods developed by Patel lab (Ramachandran et al., 2017) and others with greater detail than previously published works.

• The protocol requires several days to complete as various steps are designed to be performed overnight.

• The recombinantly purified proteins have been successfully used for in vitro transcription experiments, allowing for finer control of experimental components in a minimalistic system.

Biological processes are dependent on protein concentration and there is an inherent variability among cells even in environment-controlled conditions. Determining the amount of protein of interest in a cell is relevant to quantitatively relate it with the cells (patho)physiology. Previous studies used either western blot to determine the average amount of protein per cell in a population or fluorescence intensity to provide a relative amount of protein. This method combines both techniques. First, the protein of interest is purified, and its concentration determined. Next, cells containing the protein of interest with a fluorescent tag are sorted into different levels of intensity using fluorescence-activated cell sorting, and the amount of protein for each intensity category is calculated using the purified protein as calibration. Lastly, a calibration curve allows the direct relation of the amount of protein to the intensity levels determined with any instrument able to measure intensity levels. Once a fluorescence-based instrument is calibrated, it is possible to determine protein concentrations based on intensity.

Key features

• This method allows the evaluation and comparison of protein concentration in cells based on fluorescence intensity.

• Requires protein purification and fluorescence-activated cell sorting.

• Once calibrated for one protein, it allows determination of the levels of this protein using any fluorescence-based instrument.

• Allows to determine subcellular local protein concentration based on combining volumetric and intensity measurements.

Graphical overview

Protein level quantification across fluorescence-based platforms