Cancer Biology

Many protein families consist of multiple highly homologous proteins, whether they are encoded by different genes or originating from the same genomic location. Predominance of certain isoforms has been linked to various pathological conditions, such as cancer. Detection and relative quantification of protein isoforms in research are commonly done via immunoblotting, immunohistochemistry, or immunofluorescence, where antibodies against an isoform-specific epitope of particular family members are used. However, isoform-specific antibodies are not always available, making it impossible to decipher isoform-specific protein expression patterns. Here, we describe the insertion of the versatile 11 amino acid HiBiT tag into the genomic location of the protein of interest. This tag was developed and is distributed by Promega (Fitchburg, WI, USA). This protocol describes precise and specific protein expression analysis of highly homologous proteins through expression of the HiBiT tag, enabling protein expression quantification when specific antibodies are missing. Protein expression can be analyzed through traditional methods such as western blotting or immunofluorescence, and also in a luciferase binary reporter system, allowing for reliable and fast relative expression quantification using a plate reader.

Graphical overview

N⁶-methyladenosine (m⁶A) is the most prevalent internal modification of eukaryotic messenger RNAs (mRNAs), affecting their fold, stability, degradation, and cellular interaction(s) and implicating them in processes such as splicing, translation, export, and decay. The m⁶A modification is also extensively present in non-coding RNAs, including microRNAs (miRNAs), ribosomal RNAs (rRNAs), and transfer RNAs (tRNAs). Common m6A methylation detection techniques play an important role in understanding the biological function and potential mechanism of m⁶A, mainly including the quantification and specific localization of m⁶A modification sites. Here, we describe in detail the dot blotting method for detecting m⁶A levels in RNA (mRNA as an example), including total RNA extraction, mRNA purification, dot blotting, and data analysis. This protocol can also be used to enrich specific RNAs (such as tRNA, rRNA, or miRNA) by isolation technology to detect the m⁶A level of single RNA species, so as to facilitate further studies of the role of m6A in biological processes.

The core planar cell polarity (PCP) protein Vang/Vangl, including Vangl1 and Vangl2 in vertebrates, is indispensable during development. Our previous studies showed that the activity of Vangl is tightly controlled by two important posttranslational modifications, ubiquitination and phosphorylation. Vangl is ubiquitinated through an endoplasmic reticulum-associated degradation (ERAD) pathway and is phosphorylated by casein kinase 1 (CK1) in response to Wnt. Here, we present step-by-step procedures to analyze Vangl ubiquitination and phosphorylation, including cell culture, transfection, sample preparation, and signal detection, as well as the use of newly available phospho-specific antibodies to detect Wnt-induced Vangl2 phosphorylation. The protocol described here can be applicable to the analysis of posttranslational modifications of other membrane proteins.

RNA binding proteins (RBPs) are critical regulators of cellular phenotypes, and dysregulated RBP expression is implicated in various diseases including cancer. A single RBP can bind to and regulate the expression of many RNA molecules via a variety of mechanisms, including translational suppression, prevention of RNA degradation, and alteration in subcellular localization. To elucidate the role of a specific RBP within a given cellular context, it is essential to first identify the group of RNA molecules to which it binds. This has traditionally been achieved using cross-linking-based assays in which cells are first exposed to agents that cross-link RBPs to nucleic acids and then lysed to extract and purify the RBP-nucleic acid complexes. The nucleic acids within the mixture are then released and analyzed via conventional means (e.g., microarray analysis, qRT-PCR, RNA sequencing, or Northern blot). While cross-linking-based ribonucleoprotein immunoprecipitation (RIP) has proven its utility within some contexts, it is technically challenging, inefficient, and suboptimal given the amount of time and resources (e.g., cells and antibodies) required. Additionally, these types of studies often require the use of over-expressed versions of proteins, which can introduce artifacts. Here, we describe a streamlined version of RIP that utilizes exclusion-based purification technologies. This approach requires significantly less starting material and resources compared to traditional RIP approaches, takes less time, which is tantamount given the labile nature of RNA, and can be used with endogenously expressed proteins. The method described here can be used to study RNA-protein interactions in a variety of cellular contexts.

Graphical abstract:

DNA and RNA nucleases are wide-ranging enzymes, taking part in broad cellular processes from DNA repair to immune response control. Growing interest in the mechanisms and activities of newly discovered nucleases inspired us to share the detailed protocol of our nuclease assay (Sheppard et al., 2019). This easy and inexpensive method can provide data that enables understanding of the molecular mechanism for novel or tested nucleases, from substrate preference and cofactors involved to catalytic rate of reaction.

Visualization of RNA molecules in situ helps to better understand the functions of expressed genes. Currently, most conventional in situ hybridization methods for visualization of individual RNAs are based on fluorescence detection. Herein we present a chromogenic in situ hybridization protocol for visualization of single RNA molecules in fixed cells and tissues. The protocol is based on padlock probing and rolling circle amplification to generate detectable chromogenic signal from single RNA molecules. Chromogenic signal can avoid background autofluorescence and can be preserved for a longer period than fluorescence signal.

Heparanase, an endo-β-D-glucuronidase, cleaves cell surface and extracellular matrix heparan sulfate (HS) chains at distinct sites and plays important biological roles including modulation of cell growth and metastasis. Although a number of different types of heparanase assays have been reported to date, most are labor intensive, complex and/or expensive to carry out. We reasoned that a simpler heparanase assay could be developed using heparin labeled with Dabcyl and EDANS as donor and acceptor fluorophores so as to generate a FRET signal. Our results show that a more robust heparanase assay could be developed based on the principle studied herein and more homogeneous preparation of heparin. Yet, the assay in its current form could be used for routine screening of potential inhibitors in a high-throughput manner as well as for studying heparanase activity expressed in tumors as well as biological fluids like plasma.

Protein synthesis is one of the most fundamental biological processes to maintain cellular proteostasis. Azidohomoalaine (AHA) is a non-radioactive and “clickable” amino acid analog of methionine which can be incorporated into newly synthesized proteins. Thus, AHA-labeled nascent proteins can be detected and quantified through fluorescent labeling by "click" chemistry. Here we describe a protocol to measure protein synthesis by AHA labeling and flow cytometry. Taking advantage of gating different cell populations, we provide a typical example of the flow cytometric-based analysis of protein synthesis during the cell cycle. While we used mouse B cells in this protocol this method can be readily applied to any cell types and organisms.

The glycosaminoglycan hyaluronan (HA) is a key component of the extracellular matrix. The molecular weight of HA is heterogeneous and can reach from several million to several hundred daltons. The effect of HA on cell behavior is size dependent; fragmented HA acts as a danger signal, stimulates cell migration and proliferation and is proinflammatory, native high molecular weight HA suppresses inflammation. Therefore, it is important to analyze HA size distribution when studying the role of HA in tissue homeostasis and pathology. This protocol describes isolation of HA from mouse mammary glands but can also be applied to other tissues. The quality of the isolated HA is sufficient to analyze size distribution by gel electrophoresis (Calabro et al., 2000).

Specialized secretory cells known as goblet cells in the intestine and respiratory epithelium are responsible for the secretion of mucins. Mucins are large heavily glycosylated proteins and typically have a molecular mass higher than 10⁶ Da. These large proteins are densely substituted with short glycan chains, which have many important functional roles including determining the hydration and viscoelastic properties of the mucus gel that lines and protects the intestinal epithelium. In this protocol, we comprehensively describe the method for extraction of murine mucus and its analysis by agarose gel electrophoresis. Additionally we describe the use of High Iron Diamine-Alcian Blue, Periodic Acid Schiff’s-Alcian Blue and immune–staining methods to identify and differentiate between the different states of glycosylation on these mucin glycoproteins, in particular with a focus on sulphation and sialylation.