Developmental Biology

This protocol describes the generation of chimeric mice in which the Y chromosome is deleted from a proportion of blood cells. This model recapitulates the phenomenon of hematopoietic mosaic loss of Y chromosome (mLOY), which is frequently observed in the blood of aged men. To construct mice with hematopoietic Y chromosome loss, lineage-negative cells are isolated from the bone marrow of ROSA26-Cas9 knock-in mice. These cells are transduced with a lentivirus vector encoding a guide RNA (gRNA) that targets multiple repeats of the Y chromosome centromere, effectively removing the Y chromosome. These cells are then transplanted into lethally irradiated wildtype C57BL6 mice. Control gRNAs are designed to target either no specific region or the fourth intron of Actin gene. Transduced cells are tracked by measuring the fraction of blood cells expressing the virally encoded reporter gene tRFP. This model represents a clinically relevant model of hematopoietic mosaic loss of Y chromosome, which can be used to study the impact of mLOY on various age-related diseases.

Graphical overview

For several decades, aging in Saccharomyces cerevisiae has been studied in hopes of understanding its causes and identifying conserved pathways that also drive aging in multicellular eukaryotes. While the short lifespan and unicellular nature of budding yeast has allowed its aging process to be observed by dissecting mother cells away from daughter cells under a microscope, this technique does not allow continuous, high-resolution, and high-throughput studies to be performed. Here, we present a protocol for constructing microfluidic devices for studying yeast aging that are free from these limitations. Our approach uses multilayer photolithography and soft lithography with polydimethylsiloxane (PDMS) to construct microfluidic devices with distinct single-cell trapping regions as well as channels for supplying media and removing recently born daughter cells. By doing so, aging yeast cells can be imaged at scale for the entirety of their lifespans, and the dynamics of molecular processes within single cells can be simultaneously tracked using fluorescence microscopy.

Key features

• This protocol requires access to a photolithography lab in a cleanroom facility.

• Photolithography process for patterning photoresist on silicon wafers with multiple different feature heights.

• Soft lithography process for making PDMS microfluidic devices from silicon wafer templates.

Drosophila melanogaster is a classic model organism to study gene function as well as toxicological effects. To study gene function, the expression of a particular gene of interest is disrupted by using the widely explorable Drosophila genetic toolkit, whereas to study toxicological effects the flies are exposed to a particular toxicant through diet. These experiments often require the quantification of lethality from embryonic to adult stages, as well as the assessment of the life span in order to check the role of the gene/toxicant of interest in Drosophila. Here, we propose an experimental protocol that enables a consistent and rigorous assessment of lethality and life span of cadmium chloride (CdCl₂)–exposed or genetically perturbed flies [downregulation and overexpression of the cytosolic Cu, Zn superoxide dismutase (SOD1) gene], consecutively. The protocol insists upon the requirement of one single experimental setup that is unique, distinctive, and cost-effective as it engages minimal laboratory equipment and resources. The described methods lead to the smooth observation of the embryos, their successive stagewise transition, and life span of the adult flies post eclosion. Additionally, these methods also facilitate the assessment of crawling and climbing behavioral parameters of the larvae and adults, respectively, and allow the calculation of lethal concentration (LC₅₀) for the mentioned toxicant as well as median survival of the flies, which can be a determining factor in proceeding with further stages of experiments.

Graphical abstract

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.