Abstract: Purified RNA may need to be concentrated by precipitation for downstream applications. Precipitation of RNA with ethanol (or isopropanol) is the standard method to recover RNA from aqueous solutions.

Abstract: This protocol differs from other transformation procedures in that the bacterial culture is grown at 18°C rather than the conventional 37°C. Otherwise, the protocol is unremarkable and follows a fairly standard course. Under standard laboratory conditions, efficiencies of 1 × 108 to 3 × 108 transformed colonies/µg of plasmid DNA are typical.

Abstract: Determining the concentration of protein samples generally is accomplished either by measuring the UV absorbance at 280 nm or by reacting the protein quantitatively with dyes and/or metal ions (Bradford, Lowry, or BCA assays). For purified proteins, UV absorbance remains the most popular method because it is fast, convenient, and reproducible; it does not consume the protein; and it requires no additional reagents, standards, or incubations. No method of protein concentration determination is perfect because each is subject to a different set of constraints such as interference of buffer components and contaminating proteins in direct UV determination (A 280) or reactivity of individual proteins and buffer components with the detecting reagents in colorimetric assays. In cases in which protein concentration is critical (e.g., determination of catalytic rate constants for an enzyme), it may be advisable to compare the results of several assays.

Abstract: Cross-linked chains of polyacrylamide can be used as electrically neutral gels to separate double-stranded DNA fragments according to size and single-stranded DNAs according to size and conformation. Polyacrylamide gels have the following three major advantages over agarose gels: (1) Their resolving power is so great that they can separate molecules of DNA whose lengths differ by as little as 0.1% (i.e., 1 bp in 1000 bp). (2) They can accommodate much larger quantities of DNA than agarose gels. Up to 10 µg of DNA can be applied to a single slot (1 cm × 1 mm) of a typical polyacrylamide gel without significant loss of resolution. (3) DNA recovered from polyacrylamide gels is extremely pure and can be used for the most demanding purposes (e.g., microinjection of mouse embryos). However, polyacrylamide gels have the disadvantage of being more difficult to prepare and handle than agarose gels. Methods are presented here for preparing and running nondenaturing polyacrylamide gels and for detection of DNA in these gels by staining.

Abstract: Preparing electrocompetent bacteria is considerably easier than preparing cells for transformation by chemical methods. Bacteria are simply grown to mid-log phase, chilled, centrifuged, washed extensively with ice-cold buffer or H2O to reduce the ionic strength of the cell suspension, and then suspended in an ice-cold buffer containing 10% glycerol. DNA may be introduced immediately into the bacteria by exposing them to a short high-voltage electrical discharge. Alternatively, the cell suspension may be snap-frozen and stored at -70°C for up to 6 mo before electroporation, without loss of transforming efficiency.

Abstract: Immobilized metal affinity chromatography (IMAC) is based on the affinity of polyhistidine tracts for divalent metal cations (usually Ni2+) immobilized as transition metal chelate complexes on a chromatography resin. The main protocol here is optimized for use of Ni2+-NTA resin to purify soluble 6xHis-tagged proteins by a straightforward batch method during the binding step, followed by gravity flow for washes and elution. This protocol does not require any specialized equipment other than a simple glass or plastic column. IMAC resins can be used in multiple formats, including batch, gravity flow, centrifuge columns, and fast performance liquid chromatography (FPLC) systems. FPLC systems are designed specifically for the chromatographic separations of proteins and other biomolecules. These systems typically contain multiple pumps, an in-line UV absorption monitor, conductivity meter, pH meter, fraction collector, and other options that allow for the simultaneous purification, analysis, and fractionation of the sample. When linked with the appropriate instruments, an FPLC can become a high-precision, automated instrument that separates proteins at a high resolution. An alternative protocol is included here that describes 6xHis-tagged protein purification using FPLC. Procedures for the cleaning and regeneration of the IMAC resin for reuse are also described, and, finally, considerations for storing purified proteins are discussed.

Abstract: Mice, rats, or hamsters are immunized by giving biweekly injections of a purified antigen, cultured cells, or cDNA. For mice, if a pure, soluble protein antigen is being used and is abundant, a dose of 50-100 µg in adjuvant at each immunization is a sensible general recommendation; for rats and hamsters, a dose of 100-200 µg is sufficient. Lower doses can be used for antigens with higher immunogenicity. Adjuvants (Freund's, Ribi, Hunter's TiterMax, ImmunEasy, or Alum) should be mixed with the immunizing antigen for the first two immunizations only; Complete Freund's adjuvant is only used with the first immunization. Subsequent immunizations are performed in phosphate-buffered saline (PBS) or normal saline, with or without Incomplete Freund's adjuvant. The choice of adjuvant is dependent on the subclass of immunoglobulin required. Over the course of the 6-wk immunization schedule, each animal usually receives a total of six injections (three subcutaneous and three intraperitoneal). Once a good titer has developed against the antigen of interest, regular boosts and bleeds are performed to collect the maximum amount of serum. For rats and hamsters, boosts should be spaced every 2-3 wk, and serum samples of 400-500 µL should be collected 10-12 d after each boost. For mice, boosts should be spaced every 2-3 wk, and serum samples of 200-300 µL should be collected 10-12 d after each boost.

Abstract: Rabbits can be immunized by administering biweekly injections of a purified antigen, cultured cells, or cDNA. The minimum amount of antigen capable of inducing a response will depend on the nature of the antigen and on the host, but for rabbits, the minimum dose will be in the range of 10 μg per injection, although 100 μg per injection will be used more commonly. If a pure, soluble protein antigen is being used and is abundant, then a dose of 0.5-1 mg in adjuvant at each immunization is a sensible general recommendation. The injection sites on the rabbit are shaved and disinfected before immunization. Adjuvants are mixed with the immunizing antigen for the first two immunizations only, and Complete Freund's adjuvant is only used with the first immunization; subsequent immunizations are performed in phosphate-buffered saline (PBS) or normal saline, with or without Incomplete Freund's adjuvant. Once a good titer has developed against the antigen of interest, regular boosts and bleeds are performed to collect the maximum amount of serum. For rabbits, boosts should be spaced every 6 wk, and serum samples of 20-40 mL should be collected ∼10-12 d after each boost; typically, a single sample bleed from a rabbit will yield 25 mL of serum.

Abstract: AAV virions are built from three major capsid proteins, VP1, VP2, and VP3, at a ratio of 1:1:18. On a silver-stained SDS-polyacrylamide gel, VP1, VP2, and VP3 should be the only visible bands in a highly purified recombinant adeno-associated virus (rAAV) preparation, migrating at approximately 87, 73, and 62 kDa, respectively. This protocol describes how SDS-PAGE and silver staining can be used to determine the purity of an rAAV preparation. In addition, using a highly purified rAAV preparation whose particle titer is known, this assay can be used to derive a semiquantitative estimate of the particle concentration of a test vector.

Abstract: The amphibian Xenopus constitutes a powerful, versatile, and cost-effective nonmammalian model with which to investigate important contemporary issues of immunity relevant to human health such as ontogeny of immunity, self-tolerance, wound healing, autoimmunity, cancer immunity, immunotoxicology, and adaptation of host immune defenses to emerging pathogens. This model system presents several attractive features: an external developmental environment free of maternal influence that allows for easy experimental access from early life stages; an immune system that is remarkably similar to that of mammals; the availability of large-scale genetic and genomic resources; invaluable major histocompatibility complex (MHC)-defined inbred strains of frogs; and useful tools such as lymphoid tumor cell lines, monoclonal antibodies, and MHC tetramers. Modern reverse genetic loss-of-function and genome-editing technologies applied to immune function further empower this model. Finally, the evolutionary distance between Xenopus and mammals permits distinguishing species-specific adaptation from more conserved features of the immune system. In this introduction, the advantages and features of Xenopus for immunological research are outlined, as are existing tools, resources, and methods for using this model system.

Abstract: The Xenopus oocyte is a unique model system, allowing both the study of complex biological processes within a cellular context through expression of exogenous mRNAs and proteins, and the study of the cell, molecular, and developmental biology of the oocyte itself. During oogenesis, Xenopus oocytes grow dramatically in size, with a mature oocyte having a diameter of ∼1.3 mm, and become highly polarized, localizing many mRNAs and proteins. Thus, the mature oocyte is a repository of maternal mRNAs and proteins that will direct early embryogenesis prior to zygotic genome transcription. Importantly, the Xenopus oocyte also has the capacity to translate exogenous microinjected RNAs, which has enabled breakthroughs in a wide range of areas including cell biology, developmental biology, molecular biology, and physiology. This introduction outlines how Xenopus oocytes can be used to study a variety of important biological questions.

Abstract: Modular recirculating animal aquaculture systems incorporate UV sterilization and biological, mechanical, and activated carbon filtration, creating a nearly self-contained stable housing environment for Xenopus tropicalis Nonetheless, minimal water exchange is necessary to mitigate accumulation of metabolic waste, and regular weekly, monthly, and yearly maintenance is needed to ensure accurate and efficient operation. This protocol describes the methods for establishing a new recirculating system and the necessary maintenance, as well as water quality parameters, required for keeping Xenopus tropicalis.

Abstract: Fusion proteins that contain a glutathione S-transferase (GST) moiety can be purified to near homogeneity by affinity chromatography on glutathione-linked resins. Glutathione immobilized on a chromatography matrix, such as agarose or Sepharose, acts as a substrate for the GST moiety of fusion proteins. Contaminating proteins are washed away, and the bound GST fusion proteins are then readily displaced from the resin by elution with buffers containing free glutathione.

Abstract: An excellent source of antigens is to overexpress cloned genes in bacteria. A wide variety of coding regions can be expressed in bacteria either on their own or as fusion proteins. Indeed, it is often convenient to use vectors that express the antigen fused to an affinity tag, such as glutathione-S-transferase (GST), maltose-binding protein (MBP), or poly-histidine (e.g., 6×His). Each of these tags reliably binds to a specific affinity column enabling the antigen to be conveniently eluted under appropriate conditions. It should be noted that GST adds ∼25 kDa and MBP adds 40 kDa to the expressed protein. Because bacteria will reliably express proteins that are <80 kDa, the mass of the affinity tag should be included when designing your recombinant protein construct.

Abstract: Cells for staining are usually prepared from one of three sources: adherent cells, suspension cells, or whole tissues. Antibodies generally are applied directly to the area of the cells or tissues that is being studied. The antibodies can be labeled directly or they can be detected by using a labeled secondary reagent that will bind specifically to the primary antibody. Detection reagents for cell staining can be labeled with fluorochromes, enzymes, gold, or iodine.

Abstract: Negative staining is a simple and rapid method for studying the morphology and ultrastructure of small particulate specimens (e.g., viruses, bacteria, cell fragments, and isolated macromolecules such as proteins and nucleic acids). The technique described in this protocol involves allowing particles or fragments of cells to settle onto a support film, then applying a drop of metal salt solution to the adherent particulate specimen. The stain penetrates the interstices of the particles to bring out detail. In this situation, the preparation dries rapidly. The dissolved substance precipitates out of solution in an amorphous condition at the 0.1-nm level, and it is deposited over the support film and exposed surface of the specimen. The theoretical requirements of a good negative staining are a substance (1) of high density to provide high contrast, (2) at high solubility so that the stain does not come out of solution prematurely but does so only at the final stage of drying, (3) of high melting point and boiling point so that the material does not evaporate at high temperatures induced by the electron beam, and (4) in which the precipitate should be essentially amorphous down to the limit of resolution.

Abstract: Yeast cells display cell walls that must first be broken before the addition of detergents for lysis. This method describes the use of glass beads in combination with a mechanical bead beater to disrupt cell walls of both Saccharomyces cerevisiae or Schizosaccharomyces pombe directly in a nonionic detergent Lysis buffer containing 0.1% Nonidet P-40. Alternatively, this protocol can be applied for the lysis of yeast cells in Lysis buffer without detergent; upon completion of the bead beating, Triton X-100 is added to complete lysis. Yeast cells are cultured and collected while in log phase before being washed once and mixed together with glass beads in a tube. The applied shaking process facilitates disruption of the cell walls, upon which separation of yeast and glass beads is accomplished by forcing lysed cells through a hole created in the bottom of the tube during the centrifugation process. An alternative bead-beating protocol details the use of Lysis Buffer 2, which does not contain detergents and calls for the addition of Triton X-100 after cell lysis in the presence of glass beads. Use of Lysis Buffer 2 without detergent may avoid bubble and foam formation during the bead-beating process that could potentially denature proteins.

Abstract: In this protocol, yeast DNA is prepared by digestion of the cell wall and lysis of the resulting spheroplasts with SDS. This method reproducibly yields several micrograms of yeast DNA that can be efficiently cleaved by restriction enzymes and used as a template in polymerase chain reaction (PCR). Note that yeast colonies can also be used directly in PCR, without purifying yeast DNA.

Abstract: In two-step bacterial artificial chromosome (BAC) engineering, a single plasmid is introduced into the BAC-carrying cell lines. The shuttle vector pLD53.SCAB (or pLD53.SCAEB) carries the recA gene and the R6Kγ origin, which requires the π protein to replicate. PIR2 cells, expressing π, are typically used for the amplification of the vector and maintain about 15 copies/cell of the donor vector, which is relatively stable in this host.

Abstract: Bacterial artificial chromosome (BAC) clones are rendered electrocompetent and transformed with the recombinant shuttle vector, pLD53SCAB/AB-box. Cointegrates are selected by growth on chloramphenicol and ampicillin to ensure recombination of the shuttle vector into the BAC.

Abstract: Over many years, the Xenopus laevis embryo has provided a powerful model system to investigate how mechanical forces regulate cellular function. Here, we describe a system to apply reproducible tensile and compressive force to X. laevis animal cap tissue explants and to simultaneously assess cellular behavior using live confocal imaging.

Abstract: Sensory systems detect environmental stimuli and transform them into electrical activity patterns interpretable by the central nervous system. En route to higher brain centers, the initial sensory input is successively transformed by interposed secondary processing centers. Mapping the neuronal activity patterns at all of those stages is essential to understand sensory information processing. Larval Xenopus laevis is very well-suited for whole-brain imaging of neuronal activity. This is mainly due to its small size, transparency, and the accessibility of both peripheral and central parts of sensory systems. Here we describe a protocol for calcium imaging at several levels of the olfactory system using focal injection of chemical calcium indicator dyes or a Xenopus transgenic line with neuronal GCaMP6s expression. In combination with fast volumetric multiphoton microscopy, the calcium imaging methods described can provide detailed insight into spatiotemporal activity of entire brain regions at different stages of sensory information processing. Although the methods are broadly applicable to the central nervous system, in this work we focus on protocols for calcium imaging of glomeruli in the olfactory bulb and odor-responsive neurons in the olfactory amygdala.

Abstract: In this method, E. coli DNA Pol I binds to a nick or short gap in duplex DNA. The 5' → 3' exonuclease activity of Pol I then removes nucleotides from one strand of the DNA, creating a template for the synthesis of DNA by the 5' → 3' polymerase activity of Pol I. The simultaneous elimination of nucleotides from the 5' side and the addition of nucleotides to the 3' side result in movement of the nick (nick translation) along the DNA, which becomes labeled to high specific activity.

Abstract: Quantifying RNA is an important and necessary step before most RNA analysis techniques. Methods for quantifying RNA can be classified into two categories: ultraviolet (UV) spectrophotometric methods, which are based on the absorption spectra of the purine and pyrimidine bases; and fluorescent dye-based methods, which measure the fluorescence intensity of dyes that selectively fluoresce when bound to nucleic acids. If the RNA sample is pure (i.e., without significant amounts of contaminants such as proteins, phenol, agarose, or other nucleic acids), UV spectrophotometric measurement of the amount of UV irradiation absorbed by the bases is simple and accurate. However, if the sample contains significant quantities of impurities or if the concentration of RNA is very low, it is better to use fluorescent dye-based methods. An overview of spectrophotometric and fluorescent dye-based RNA quantification methods is given here, as are several options for storing purified RNA preparations. Proper storage of RNA samples is important; it can help minimize RNase contamination and consequent sample degradation.

Abstract: Escherichia coli DNA Pol I can carry out three enzymatic reactions: It possesses 5' → 3' DNA polymerase activity and 3' → 5' and 5' → 3' exonuclease activity. Pol I can be cleaved by mild treatment with subtilisin into two fragments; the larger fragment is known as the Klenow fragment. The Klenow fragment retains the polymerizing activity and the 3' → 5' exonuclease of the holo-enzyme but lacks its powerful 5' → 3' exonuclease activity. These enzymes and their applications in molecular cloning are introduced here.

Abstract: Differential detergent fractionation of cells is a rapid method for extraction of cytoplasmic and nuclear proteins in preparation of an immunoprecipitation. This method can be applied for use of adherent or suspension cells and can significantly reduce nonspecific background in an immunoprecipitation by separation of cellular compartments into individual fractions. The lysis of cells by differential detergents permits the rapid extraction of proteins from the cytoplasm (digitonin), the cytoplasmic membranes, and organelles (Triton X-100), and nucleoplasm (Tween/DOC), facilitated through the use of distinct extraction buffers. Cytoplasmic and nuclear matrix proteins as well as DNA are left behind during the detergent-based extraction.

Abstract: This protocol describes procedures for cloning blunt-ended DNA fragments into linearized plasmid vectors. To obtain the maximum number of "correct" ligation products when cloning blunt-ended target fragments, the two components of DNA in the ligation reaction must be present at an appropriate ratio. If the molar ratio of plasmid vector to target DNA is too high, then the ligation reaction may generate an undesirable number of circular empty plasmids, both monomeric and polymeric; if too low, the ligation reaction may generate an excess of linear and circular homopolymers and heteropolymers of varying sizes, orientations, and compositions. For this reason, the orientation of the foreign DNA and the number of inserts in each recombinant clone must always be validated by restriction endonuclease mapping or some other means.

Abstract: In the one-step approach to bacterial artificial chromosome (BAC) modification, two plasmids are introduced into the BAC host cells. The shuttle pLD53.SC2, carrying the EFGP reporter sequence and requiring the π protein to replicate, must be grown in PIR1- or PIR2-competent Escherichia coli Our preference for these vectors is PIR1, because these cells are able to maintain about 250 copies of the donor vector. This small-sized vector is stable in PIR1. The RecA plasmid pSV1.RecA has a temperature-sensitive origin of replication and can be grown in most competent bacteria at 30°C; here we use DH5α competent cells. This protocol describes preparation of the vector DNAs. The shuttle-reporter vector DNA is subsequently digested for introduction of one homology arm (typically the A-box).

Abstract: Many of the commonly used techniques in molecular cloning depend on methods to map accurately the distribution of radioactive atoms on two-dimensional (2D) surfaces. Without this ability, methods such as Southern blotting, northern hybridizations, radiolabeled DNA sequencing, and library screening would not have been possible. In the 1970s and 1980s-the pioneering days of molecular cloning-imaging of 2D surfaces was obtained using autoradiography. In this technique, β-particles emitted by radioactive specimens were recorded on X-ray film, producing a latent image that can be converted to a true image by developing and fixing the film. Autoradiography was a lot of fun, but it was also messy. In the impatient excitement of wanting to see how an experiment had turned out, people used to hold the newly developed X-ray films in their metal frames up to the darkroom light. Drips of the final wash would run down their arms, clothes would be stained, and shoes ruined. It is hardly surprising that autoradiography was quickly abandoned when sensitive phosphorimagers came onto the market at the end of the 1990s.