Cell Biology

Circular RNA (circRNA) is an intriguing class of non-coding RNA that exists as a continuous closed loop. With the improvements in high throughput sequencing, biochemical analysis, and bioinformatic algorithms, studies on circRNA expression became abundant in recent years. However, functional studies of circRNA are still limited. Subcellular localization of circRNA may provide some clues in elucidating its biological functions by performing subcellular fractionation assay. Notably, circRNAs that are predominantly found in the cytoplasm are more likely to be involved in post-transcriptional gene regulation, e.g., acting as micoRNA sponge, whereas nuclear-retained circRNAs are predicted to play a role in transcriptional regulation. Subcellular fractionation could help researchers to narrow down and prioritize downstream experiments. The majority of the currently available protocols describe the steps for subcellular fractionation followed by western blot analysis for protein molecules. Here, we present a protocol for the subcellular fractionation of cells to detect circRNA via RT-qPCR with divergent primers. Moreover, detailed steps for the generation of specific circRNAs-enriched cDNA included in this protocol will enhance the amplification and detection of low-abundance circRNAs. This will be useful for researchers studying low-abundance circRNAs.

Key features

• This protocol builds upon the method developed by Gagnon et al. (2014) and extends its application to circRNA study.

• Protocol for amplification of low levels of circRNA expression.

• Analysis takes into consideration the ratio of cytoplasmic RNA concentration to nuclear RNA concentration.

Graphical overview

The subfractionation of the endoplasmic reticulum (ER) is a widely used technique in cell biology. However, current protocols present limitations such as low yield, the use of large number of dishes, and contamination with other organelles. Here, we describe an improved method for ER subfractionation that solves other reported methods' main limitations of being time consuming and requiring less starting material. Our protocol involves a combination of different centrifugations and special buffer incubations as well as a fine-tuned method for homogenization followed by western blotting to confirm the purity of the fractions. This protocol contains a method to extract clean ER samples from cells using only five (150 mm) dishes instead of over 50 plates needed in other protocols. In addition, in this article we not only propose a new cell fractionation approach but also an optimized method to isolate pure ER fractions from one mouse liver instead of three, which are commonly used in other protocols. The protocols described here are optimized for time efficiency and designed for seamless execution in any laboratory, eliminating the need for special/patented reagents.

Key features

• Subcellular fractionation from cells and mouse liver.

• Uses only five dishes (150 mm) or one mouse liver to extract highly enriched endoplasmic reticulum without mitochondrial-associated membrane contamination.

• These protocols require the use of ultracentrifuges, dounce homogenizers, and/or Teflon Potter Elvehjem.

• As a result, highly enriched/clean samples are obtained.

Graphical overview

Many postitive-stranded RNA viruses, such as Hepatitis C virus (HCV), highjack cellular membranes, including the Golgi, ER, mitchondria, lipid droplets, and utilize them for replication of their RNA genome or assembly of new virions. By investigating how viral proteins associate with cellular membranes we will better understand the roles of cellular membranes in the viral life cycle. Our lab has focused specifically on the role of lipid droplets and lipid-rich membranes in the life cycle of HCV. To analyze the role of lipid-rich membranes in HCV RNA replication, we utilized a membrane flotation assay based on an 10-20-30% iodixanol density gradient developed by Yeaman et al. (2001). This gradient results in a linear increase in density over almost the entire length of the gradient, and membrane particles are separated in the gradient based on their buoyant characteristics. To preserve membranes in the lysate, cells are broken mechanically in a buffer lacking detergent. The cell lysate is loaded on the bottom of the gradient, overlaid with the gradient, and membranes float up as the iodixanol gradient self-generates. The lipid content of membranes and the concentration of associated proteins will determine the separation of different membranes within the gradient. After centrifugation, fractions can be sampled from the top of the gradient and analyzed using standard SDS-PAGE and western blot analysis for proteins of interest.

It has recently been reported that tomato fruit chromoplasts can synthesize ATP de novo using an ATP synthase complex harboring an atypical γ-subunit which is also present in a variety of plant species. However many aspects related with the biochemical processes underlying this process remain largely unknown. Here we describe detailed protocols for the isolation of tomato fruit chromoplasts and the determination of ATP levels (end-point measurements) and ATP synthesis rates (kinetic measurements) in these organelles using bioluminescent luciferin/luciferase based assays.

Here, we introduce the protocol for small-scale and simple subcellular fractionation used in our recent publication (Taguchi et al., 2013), which uses homogenization by passing through needles and sucrose step-gradient.

Subcellular fractionation is a very useful technique but usually a large number of cells are required. Because we needed subcellular fractionation of transiently-transfected cells, we developed a protocol for smaller numbers of cells. Our protocol for the subcellular fractionation is based on the protocol published by de Araújo and Huber (de Araujo et al., 2007), although substantial modifications have been made according to our experiences and information from personal communications. As optimal conditions seem to vary between cell lines, we advise to further modify the protocol to optimize for individual experiments. Our method is simple but sufficient for analysis of integral membrane proteins or proteins anchored to organelles by glycosylphosphatidylinositol or other lipid anchors, e.g. prion protein. However, proteins non-covalently attached to membranes or membrane proteins of organelles seem to be more prone to dissociation from the organelles during preparation and, if these proteins are the object of study, further modifications might be necessary.

Unlike in a continuous gradient, where a protein of interest is scattered over a wide range, step-gradient fractionation is advantageous in detection of relatively small amounts of proteins from small-scale experiments, because it concentrates the protein of interest in one fraction, if an appropriate combination of sucrose concentrations is used.

Pseudomonas aeruginosa is a Gram negative bacterium. Separating the cell envelope compartments enables proteins to be localized to confirm where in the cell they function. Cell fractionation can also provide a first step in a protein purification strategy (Williams et al., 1998). This protocol has been designed to obtain the different fractions of P. aeruginosa, namely the inner membrane, outer membrane, cytoplasmic and periplasmic compartments. Specific detection of the arginine specific autotransporter (AaaA) (Luckett et al., 2012) in the outer membrane of P. aeruginosa has been performed using this protocol.

The understanding of the organization of postsynaptic signaling systems at excitatory synapses has been aided by the identification of proteins in the postsynaptic density (PSD) fraction, a subcellular fraction enriched in structures with the morphology of PSDs. Here we described an efficient way to isolate the crude synaptosome, presynaptic fraction, and PSD fraction. It helps to identify the location of synaptic protein and find the potential synaptic complex.

Growth cones are motile structures at the tips of growing neurites, which play an essential role in regulation of growth and navigation of growing axons and dendrites of neurons in the developing nervous system. This protocol describes isolation of growth cones from the brain tissue from young mice. Growth cones isolated using this protocol have been extensively characterized using electron microscopy (Pfenninger et al., 1983) and may be used for any kind of subsequent biochemical and/or functional analyses, including Western blot analysis of protein expression (Westphal et al., 2010), analysis of the activity of growth cone-accumulated enzymes (Leshchyns’ka et al., 2003; Li et al., 2013), and analysis of the endocytosis and exocytosis rates (Chernyshova et al., 2011).

This subcellular fractionation protocol is used for separation of cellular organelles based on their density. We have designed and optimized the protocol for separation of subcellular compartments of brain homogenates with focus on the localization and trafficking of transmembrane proteins, but we have also successfully used this protocol for fractionation of other types of tissue. The protocol has two major steps 1) preparation of homogenate from dissected tissue and 2) separation of organelles by centrifugation of homogenates using a continuous sucrose gradient.

Subcellular localization is crucial for the proper functioning of a protein. Deregulation of subcellular localization may lead to pathological consequences and result in diseases like cancer. Immuno-fluorescent staining and subcellular fractionation can be used to determine localization of a protein. Here we discuss a protocol to separate the nuclear, cytosolic, and membrane fractions of cultured human cell lines using a centrifuge and ultracentrifuge. The membrane fraction contains plasma membranes and ER-golgi membranes, but no mitochondria or nuclear structures. The fractions can be further analyzed using Western blotting. This protocol is based on that from Dr. Richard Patten at Abcam, and was modified and utilized in a publication by Huang et al. (2012).