The Classic Labyrinth Test (CLT) is a simple way to evaluate behaviors in rodents such as learning ability, memory, and anxiety. The protocol presented here describes the procedure for use with rats, but the protocol can also be adapted for use in mice if a smaller device is used. In short, the CLT uses a square-shaped maze with a starting point and a stopping point. After the animal is trained, the animal is allowed to view and explore the labyrinth freely for 10 min. During this time, all of the animal's vertical and horizontal movements within the labyrinth are recorded. This is a very challenging task because it requires the animal to remember the quickest path between the starting points and the end. In cases where the labyrinth is designed so that the animal only needs to walk forward, it is quite easy for healthy rats, but for rats exposed to neuro-xenobiotics (drugs, pesticides) there will be disturbances in their path. Researchers use many different versions of this test and the procedure for each version can vary significantly. Here, we present a working protocol that enables the detection of traces of some toxic substances that may be exposed to individuals over a long period and in very small amounts under specific conditions such as drugs, medicines and pesticides.

Osteoblasts are bone marrow endosteum-lining niche cells playing important roles in the regulation of hematopoietic stem cells by secreting factors and cell adhesion molecules. Characterization of primary osteoblasts has been achieved through culture of outgrowth of collagenase treated bone. Immunophenotyping and flow-based analysis of long bone osteoblasts offer a simplified and rapid approach to characterize osteoblasts. We describe a modified procedure of isolating mouse bone marrow osteoblastic cells based on cell surface immunophenotyping. The chemokine CXCL12 (also known as stromal-derived factor, SDF-1) together with its receptor CXCR4 are expressed by osteoblasts and bone marrow stroma cells. The CXCL12-CXCR4 axis is important for hematopoietic stem cell retention to their niches (Sugiyama et al., 2006) and for supporting leukemia initiating cell activity (Pitt et al., 2015). Here we describe the procedure of intracellular staining of CXCL12.

Non-muscle myosin II (NMII) form bipolar filaments, which bind F-actin to exert cellular contractility during physiological processes (Vicente-Manzanares et al., 2009). Using a combinatorial approach to fluorescently label both N- and C-termini of the NMII heavy chain, recent works have demonstrated the ability to visualize NMII bipolar filaments at various subcellular localizations (Ebrahim et al., 2013; Beach et al., 2014). At the zonula adherens (ZA) of epithelia, NMII minifilaments bind the circumferential actin bundles in a pseudo-sarcomeric manner (Ebrahim et al., 2013), a conformation required to maintain junctional tension and tissue integrity (Ratheesh et al., 2012). By expressing green fluorescent protein (GFP)-NMIIA heavy chain and immunolabel it using a NMIIA C-terminus specific antibody, we were able to visualize the NMII minifilaments bound to F-actin bundles in Caco-2 cells (Michael et al., 2016), as previously reported (Ebrahim et al., 2013; Beach et al., 2014). In addition, we designed an FIJI/MATLAB analysis module to quantify the size, distance and alignment of these minifilaments with respect to junctional F-actin at the ZA. Measurements of the dispersion of minifilaments angles were proven to be a useful parameter that closely correlated to the extent of contractility at junctions (Michael et al., 2016).

Macrophages represent a widely distributed and functionally diverse population of innate myeloid cells involved in inflammatory response to pathogens, tissue homeostasis and tissue repair (Murray and Wynn, 2011). Macrophages can be broadly grouped into two subpopulations with opposing activites: M1 or pro-inflammatory macrophages that promote T-helper type 1 (Th1) cell immunity and tissue damage, and M2 or anti-inflammatory/alternatively activated macrophages implicated in Th2 response and resolution of inflammation. Here we describe a rapid assay we used previously to monitor changes in pro-inflammatory and anti-inflammatory cytokine production by lipopolysaccharide (LPS)-activated macrophages in response to therapeutic paracrine factors produced by adult stem cells (Bartosh et al., 2010; Ylostalo et al., 2012; Bartosh et al., 2013). The assay can be adapted appropriately to test macrophage response to other agents as well that will be referred to herein as ‘test reagents’ or ‘test compounds’.

In this protocol, the mouse macrophage cell line J774A.1 is expanded as an adherent monolayer on petri dishes allowing for the cells to be harvested easily without enzymes or cell scrapers that can damage the cells. The macropahges are then stimulated in suspension with LPS and seeded into 12-well cell culture plates containing the test reagents. After 16-18 h, the medium conditioned by the macrophages is harvested and the cytokine profile in the medium determined with enzyme-linked immunosorbent assays (ELISA). We routinely measure levels of the pro-inflammtory cytokine TNF-alpha and the anti-inflammatory cytokine interleukin-10 (IL-10).

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.

In this protocol, engineered Cas9-ribonucleoprotein (Cas9 protein and sgRNA, together called Cas9-RNP) and gold nanoparticles are used to make nanoassemblies that are employed to deliver Cas9-RNP into cell cytoplasm and nucleus. Cas9 protein is engineered with an N-terminus glutamic acid tag (E-tag or En, where n = the number of glutamic acid in an E-tag and usually n = 15 or 20), C-terminus nuclear localizing signal (NLS), and a C-terminus 6xHis-tag. [Cas9En hereafter]

To use this protocol, the first step is to generate the required materials (gold nanoparticles, recombinant Cas9En, and sgRNA). Laboratory-synthesis of gold nanoparticles can take up to a few weeks, but can be synthesized in large batches that can be used for many years without compromising the quality. Cas9En can be cloned from a regular SpCas9 gene (Addgene plasmid id = 47327), and expressed and purified using standard laboratory procedures which are not a part of this protocol. Similarly, sgRNA can be laboratory-synthesized using in vitro transcription from a template gene (Addgene plasmid id = 51765) or can be purchased from various sources.

Once these materials are ready, it takes about ~30 min to make the Cas9En-RNP complex and 10 min to make the Cas9En-RNP/nanoparticles nanoassemblies, which are immediately used for delivery (Figure 1). Complete delivery (90-95% cytoplasmic and nuclear delivery) is achieved in less than 3 h. Follow-up editing experiments require additional time based on users’ need.

Synthesis of arginine functionalized gold nanoparticles (ArgNPs) (Yang et al., 2011), expression of recombinant Cas9En, and in vitro synthesis of sgRNA is reported elsewhere (Mout et al., 2017). We report here only the generation of the delivery vehicle i.e., the fabrication of Cas9En-RNP/ArgNPs nanoassembly.

CD38 is a multifunctional enzyme involved in calcium signaling and Nicotinamide Adenine Dinucleotide (NAD⁺) metabolism. Through its major activity, the hydrolysis of NAD⁺, CD38 helps maintain the appropriate levels of this molecule for all NAD⁺-dependent metabolic processes to occur. Due to current advances and studies relating NAD⁺ decline and the development of multiple age-related conditions and diseases, CD38 gained importance in both basic science and clinical settings. The discovery and development of strategies to modulate its function and, possibly, treat diseases and improve health span put CD38 under the spotlights. Therefore, a consistent and reliable method to measure its activity and explore its use in medicine is required. We describe here the methods how our group measures both the hydrolase and cyclase activity of CD38, utilizing a fluorescence-based enzymatic assay performed in a plate reader using 1,N⁶-Ethenonicotinamide Adenine Dinucleotide (ε-NAD) and Nicotinamide Guanine Dinucleotide (NGD) as substrates, respectively.

Connexins (Cxs) are integral membrane proteins of vertebrates that associate to form hexameric transmembrane channels, named hemichannels. Twenty-one Cx types have been described, which are named according to their molecular weight. Cxs are expressed in many cell types, e.g. epithelial cells, astrocytes and immune cells. Hemichannels allow the passage of molecules of up to 1-2 kDa along the concentration gradient. When surface-exposed, hemichannels mediate the exchange of molecules between the cytosol and the extracellular space. Hemichannels are closed by default, but several cues inducing their opening have been described, e.g. a drop in the extracellular Ca²⁺ concentration (Evans et al., 2006) or infection with enteric pathogens (Puhar et al., 2013; Tran Van Nhieu et al., 2003). Hemichannel opening can be measured by electrophysiology, by quantifying the release of a hemichannel-permeable molecule into the extracellular medium or by quantifying the uptake of a hemichannel-permeable, plasma membrane-impermeant molecule. As the extent of uptake of a molecule is proportional to its concentration, exposure time, temperature (these parameters are controlled) and, importantly, to the number of active hemichannels on the cell surface, uptake assays are routinely used to assess hemichannel opening. This protocol for the uptake of the fluorescent dye ethidium bromide was used with Hela cells that were stably transfected with Cx26 or Cx43 (Paemeleire et al., 2000). Nevertheless, it could likely be used with other Cx-expressing cell types.