发育生物学

Maintenance of DNA integrity is of pivotal importance for cells to circumvent detrimental processes that can ultimately lead to the development of various diseases. In the face of a plethora of endogenous and exogenous DNA damaging agents, cells have evolved a variety of DNA repair mechanisms that are responsible for safeguarding genetic integrity. Given the relevance of DNA damage and its repair for disease pathogenesis, measuring them is of considerable interest, and the comet assay is a widely used method for this. Cells treated with DNA damaging agents are embedded into a thin layer of agarose on top of a microscope slide. Subsequent lysis removes all protein and lipid components to leave ‘nucleoids’ consisting of naked DNA remaining in the agarose. These nucleoids are then subjected to electrophoresis, whereby the negatively charged DNA migrates towards the anode depending on its degree of fragmentation, creating shapes resembling comets, which can be visualized and analysed by fluorescence microscopy. The comet assay can be adapted to assess a wide variety of genotoxins and repair kinetics, and both DNA single-strand and double-strand breaks. In this protocol, we describe in detail how to perform the neutral comet assay to assess double-strand breaks and their repair using cultured human cell lines. We describe the workflow for assessing the amount of DNA damage generated by ionizing radiation or present endogenously in the cells, and how to assess the repair kinetics after such an insult. The procedure described herein is easy to follow and cost-effective.

An important but often overlooked aspect of gene regulation occurs at the level of protein translation. Many genes are regulated not only by transcription but by their propensity to be recruited to actively translating ribosomes (polysomes). Polysome profiling allows for the separation of unbound 40S and 60S subunits, 80S monosomes, and actively translating mRNA bound by two or more ribosomes. Thus, this technique allows for actively translated mRNA to be isolated. Transcript abundance can then be compared between actively translated mRNA and all mRNA present in a sample to identify instances of post-transcriptional regulation. Additionally, polysome profiling can be used as a readout of global translation rates by quantifying the proportion of actively translating ribosomes within a sample. Previously established protocols for polysome profiling rely on the absorbance of RNA to visualize the presence of polysomes within the fractions. However, with the advent of flow cells capable of detecting fluorescence, the association of fluorescently tagged proteins with polysomes can be detected and quantified in addition to the absorbance of RNA. This protocol provides detailed instructions on how to perform fluorescent polysome profiling in C. elegans to collect actively translated mRNA, to quantify changes in global translation, and to detect ribosomal binding partners.

Whole-lifespan single-cell analysis has greatly increased our understanding of fundamental cellular processes such as cellular aging. To observe individual cells across their entire lifespan, all progeny must be removed from the growth medium, typically via manual microdissection. However, manual microdissection is laborious, low-throughput, and incompatible with fluorescence microscopy. Here, we describe assembly and operation of the multiplexed-Fission Yeast Lifespan Microdissector (multFYLM), a high-throughput microfluidic device for rapidly acquiring single-cell whole-lifespan imaging. multFYLM captures approximately one thousand rod-shaped fission yeast cells from up to six different genetic backgrounds or treatment regimens. The immobilized cells are fluorescently imaged for over a week, while the progeny cells are removed from the device. The resulting datasets yield high-resolution multi-channel images that record each cell’s replicative lifespan. We anticipate that the multFYLM will be broadly applicable for single-cell whole-lifespan studies in the fission yeast (Schizosaccharomyces pombe) and other symmetrically-dividing unicellular organisms.

The Smurf Assay (SA) was initially developed in the model organism Drosophila melanogaster where a dramatic increase of intestinal permeability has been shown to occur during aging (Rera et al., 2011). We have since validated the protocol in multiple other model organisms (Dambroise et al., 2016) and have utilized the assay to further our understanding of aging (Tricoire and Rera, 2015; Rera et al., 2018). The SA has now also been used by other labs to assess intestinal barrier permeability (Clark et al., 2015; Katzenberger et al., 2015; Barekat et al., 2016; Chakrabarti et al., 2016; Gelino et al., 2016). The SA in itself is simple; however, numerous small details can have a considerable impact on its experimental validity and subsequent interpretation. Here, we provide a detailed update on the SA technique and explain how to catch a Smurf while avoiding the most common experimental fallacies.

Genomic sequencing efforts can implicate large numbers of genes and de novo mutations as potential disease risk factors. A high throughput in vivo model system to validate candidate gene association with pathology is therefore useful. We present such a system employing Drosophila to validate candidate congenital heart disease (CHD) genes. The protocols exploit comprehensive libraries of UAS-GeneX-RNAi fly strains that when crossed into a 4XHand-Gal4 genetic background afford highly efficient cardiac-specific knockdown of endogenous fly orthologs of human genes. A panel of quantitative assays evaluates phenotypic severity across multiple cardiac parameters. These include developmental lethality, larva and adult heart morphology, and adult longevity. These protocols were recently used to evaluate more than 100 candidate CHD genes implicated by patient whole-exome sequencing (Zhu et al., 2017).

The rate of oxygen consumption is a vital marker indicating cellular function during lifetime under normal or metabolically challenged conditions. It is used broadly to study mitochondrial function (Artal-Sanz and Tavernarakis, 2009; Palikaras et al., 2015; Ryu et al., 2016) or investigate factors mediating the switch from oxidative phosphorylation to aerobic glycolysis (Chen et al., 2015; Vander Heiden et al., 2009). In this protocol, we describe a method for the determination of oxygen consumption rates in the nematode Caenorhabditis elegans.

Eukaryotic cells heavily depend on adenosine triphosphate (ATP) generated by oxidative phosphorylation (OXPHOS) within mitochondria. ATP is the major energy currency molecule, which fuels cell to carry out numerous processes, including growth, differentiation, transportation and cell death among others (Khakh and Burnstock, 2009). Therefore, ATP levels can serve as a metabolic gauge for cellular homeostasis and survival (Artal-Sanz and Tavernarakis, 2009; Gomes et al., 2011; Palikaras et al., 2015). In this protocol, we describe a method for the determination of intracellular ATP levels using a bioluminescence approach in the nematode Caenorhabditis elegans.