Molecular Biology

Utilizingresources available from the mother's body to guarantee healthy offspring growth is the fundamental reproductive strategy. Recently, we showed that a class of the largest extracellular vesicles known as exophers, which are responsible for the removal of neurotoxic components from neurons (Melentijevic et al., 2017) and damaged mitochondria from cardiomyocytes (Nicolás-Ávila et al., 2020), are released by the Caenorhabditis elegans hermaphrodite body wall muscles (BWM), to support embryonic growth (Turek et al., 2021). Employing worms expressing fluorescent reporters in BWM cells, we found that exopher formation (exophergenesis) is sex-specific and fertility-dependent. Moreover, exophergenesis is regulated by the developing embryo in utero, and exophers serve as transporters for muscle-generated yolk proteins, which can be used to nourish the next generation. Given the specific regulation of muscular exophergenesis, and the fact that muscle-generated exophers are much larger than neuronal ones and have different targeting, their identification and quantification required a modified approach from that designed for neuronal-derived exophers (Arnold et al., 2020). Here, we present a methodology for assessing and quantifying muscle-derived exophers that can be easily extended to determine their function and regulation in various biological contexts.

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Advances in imaging technology offer new opportunities in developmental biology. To observe the development of internal structures, microtome cross-sectioning followed by H&E staining on glass slides is a common procedure; however, this technique can be destructive, and artifacts can be introduced during the process. In this protocol, we describe a less invasive procedure with which we can stain insect samples and obtain reconstructed three-dimensional images using micro-computed tomography, or micro-CT (µCT). Specifically, we utilize the fungus-farming ambrosia beetle species Euwallacea validus to observe the morphology of mycangia, a critical internal organ with which beetles transport fungal symbionts. Not only this protocol is ideal to observe mycangia, our staining/scanning procedure can also be applied to observe other delicate tissues and small organs in arthropods.

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RIBO-seq and proteogenomics have revealed that mammalian genomes harbor thousands of unannotated small and alternative open reading frames (smORFs, <100 amino acids, and alt-ORFs, >100 amino acids, respectively). Several dozen mammalian smORF-encoded proteins (SEPs) and alt-ORF-encoded proteins (alt-proteins) have been shown to play important biological roles, while the overwhelming majority of smORFs and alt-ORFs remain uncharacterized, particularly at the molecular level. Functional proteomics has the potential to reveal key properties of unannotated SEPs and alt-proteins in high throughput, and an approach to identify SEPs and alt-proteins undergoing regulated synthesis should be of broad utility. Here, we introduce a chemoproteomic pipeline based on bio-orthogonal non-canonical amino acid tagging (BONCAT) (Dieterich et al., 2006) to profile nascent SEPs and alt-proteins in human cells. This approach is able to identify cellular stress-induced and cell-cycle regulated SEPs and alt-proteins in cells.

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Schematic overview of BONCAT-based chemoproteomic profiling of nascent, unannotated small and alternative open reading frame-encoded proteins (SEPs and alt-proteins)

Sclerotinia sclerotiorum causes white mold, leading to substantial losses on a wide variety of hosts around the world. Many genes encoding effector proteins play important roles in the pathogenesis of S. sclerotiorum. Therefore, establishment of a transformation system for the exploration of gene function is necessarily significant. Here, we introduce a modified protocol to acquire protoplasts for transformation and generate knockout strains, which completements the transformation system of S. sclerotiorum.

MicroRNAs (miRNA) are small (21–24 nt) non-coding RNAs involved in many biological processes in both plants and animals. The biogenesis of plant miRNAs starts with the transcription of MIRNA (MIR) genes by RNA polymerase II; then, the primary miRNA transcripts are cleaved by Dicer-like proteins into mature miRNAs, which are then loaded into Argonaute (AGO) proteins to form the effector complex, the miRNA-induced silencing complex (miRISC). In Arabidopsis , some MIR genes are expressed in a tissue-specific manner; however, the spatial patterns of MIR gene expression may not be the same as the spatial distribution of miRISCs due to the non-cell autonomous nature of some miRNAs, making it challenging to characterize the spatial profiles of miRNAs. A previous study utilized protoplasting of green fluorescent protein (GFP) marker transgenic lines followed by fluorescence-activated cell sorting (FACS) to isolate cell-type-specific small RNAs. However, the invasiveness of this approach during the protoplasting and cell sorting may stimulate the expression of stress-related miRNAs. To non-invasively profile cell-type-specific miRNAs, we generated transgenic lines in which root cell layer-specific promoters drive the expression of AGO1 and performed immunoprecipitation to non-invasively isolate cell-layer-specific miRISCs. In this protocol, we provide a detailed description of immunoprecipitation of root cell layer-specific GFP-AGO1 using EN7::GFP-AGO1 and ACL5::GFP-AGO1 transgenic plants, followed by small RNA sequencing to profile single-cell-type-specific miRNAs. This protocol is also suitable to profile cell-type-specific miRISCs in other tissues or organs in plants, such as flowers or leaves.

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CRISPR/Cas9 screening has revolutionized functional genomics in biomedical research and is a widely used approach for the identification of genetic dependencies in cancer cells. Here, we present an efficient and versatile protocol for the cloning of guide RNAs (gRNA) into lentiviral vectors, the production of lentiviral supernatants, and the transduction of target cells in a 96-well format. To assess the effect of gene knockouts on cellular fitness, we describe a competition-based cell proliferation assay using flow cytometry, enabling the screening of many genes at the same time in a fast and reproducible manner. This readout can be extended to any parameter that is accessible to flow-based measurements, such as protein expression and stability, differentiation, cell death, and others. In summary, this protocol allows to functionally assess the effect of a set of 50–300 gene knockouts on various cellular parameters within eight weeks.

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Cloning systems like Gateway and Golden Gate/Braid are known because of their efficiency and accuracy. While the main drawback of Gateway is the expensive cost of the enzymes used in its two-step (LR and BP) reaction, Golden Gate requires non-reusable components due to their specific restriction sites. We present the Brick into the Gateway (BiG) protocol as a new cloning strategy, faster and more economic method that combines (i) reusable modules or bricks assembled by the GoldenBraid approach, and (ii) Gateway LR reactions [recombination of attachment sites: attL (L from left) and attR (R from right)] avoiding the BP reaction [recombination of attachment sites: attP (P from phage) and attB (B from bacteria)] usually necessary in the Gateway cloning. The starting point is to perform a PCR reaction to add type IIS restriction sites into DNA fragments generating specific fusion sites. Then, this PCR product is used to design GoldenBraid bricks, including the attL Gateway recombination sites. Using the Golden Gate method, these bricks are assembled to produce an attL1–gene of interest–attL2 fragment, which is integrated into a compatible vector producing a Gateway entry vector. Finally, the fragment containing the target gene is recombined by LR reaction into the Gateway destination vector.

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RNA is a vital component of the cell and is involved in a diverse range of cellular processes through a variety of functions. However, many of these functions cannot be performed without interactions with proteins. There are currently several techniques used to study protein–RNA interactions, such as electrophoretic mobility shift assay, fluorescence anisotropy, and filter binding. RNA-pulldown is a technique that uses biotinylated RNA probes to capture protein–RNA complexes of interest. First, the RNA probe and a recombinant protein are incubated to allow the in vitro interaction to occur. The fraction of bound protein is then captured by a biotin pull-down using streptavidin-agarose beads, followed by elution and immunoblotting for the recombinant protein with a His-tag–reactive probe. Overall, this method does not require specialized equipment outside what is typically found in a modern molecular laboratory and easily facilitates the maintenance of an RNase-free environment.

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Genetic transformation is a powerful method for the investigation of gene function and improvement of crop plants. The transgenes copy number in the transgenic line is involved in gene expression level and phenotypes. Additionally, identification of transgene zygosity is important for quantitative assessment of phenotype and for tracking the inheritance of transgenes in progeny generations. Several methods have been developed for estimating the transgene copy number, including southern blot assay and quantitative polymerase chain reaction (qPCR) experiments. Southern hybridization, although convincing and reliable, is a time-consuming method through which the examination of the copy number is challenging in species with large genomes like wheat plants. Although qPCR is potentially simpler to perform, its results lack accuracy and precision, especially to distinguish between one and two copy events in transgenic plants with large genomes. The droplet digital PCR (ddPCR)–based method for investigation of transgenes copy number has been widely used in an array of crops. In this method, the specific primers to amplify target transgenes and reference genes are used as a single duplexed reaction, which is divided into tens of thousands of nanodroplets. The copy number in independent transgenic lines is determined by detection and quantification of droplets using sequence-specific fluorescently labeled probes. This method offers superior accuracy and reliability with a low cost and scalability as other PCR techniques in the investigation of transgenes copy number.

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Flow chart for the ddPCR protocol

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.