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The only way to solubilize many antigens for immunoprecipitation is by denaturation. This cell lysis protocol is ideally suited for this purpose to release proteins from complex structures or reveal antibody epitopes hidden within native proteins. Short linear epitopes may not be accessible to antibodies within the native tertiary and quaternary protein structures, but they become exposed upon the unraveling of proteins, exposing their secondary structure. Antibodies otherwise not suitable for the immunoprecipitation of proteins prepared under nondenaturing conditions are now able to bind these antigens of interest in cell lysates prepared under denaturing conditions. These antibodies may also work well for immunoblotting purposes when the protein target is completely denatured. Harvested cells in this protocol are washed in tris-buffered saline (TBS) before lysis in 2% sodium dodecyl sulfate (SDS)-containing Lysis buffer for 10 min at 100°C. The resulting sample is diluted 20-fold in TBS to reduce the SDS concentration to ≤0.1% before the addition of an antibody for immunoprecipitation. Addition of 2% bovine serum albumin (BSA) or 0.1% Nonidet P-40 to the TBS before an immunoprecipitation, respectively, ensures either removal of SDS from the target protein or retaining denatured proteins in solution.

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.

This rapid and efficient method to prepare highly purified recombinant adeno-associated viruses (rAAVs) is based on binding of negatively charged rAAV capsids to an anion-exchange resin that is pH dependent.

This is a simple method for rapid preparation of recombinant adeno-associated virus (rAAV) stocks, which can be used for in vivo gene delivery. The purity of these vectors is considerably lower than that obtained by either CsCl gradient centrifugation or by combination of iodixanol gradient ultracentrifugation followed by column chromatography.

The most commonly used method for production of recombinant adeno-associated virus (rAAVs) in research laboratories is by transient triple transfection of 293 cells with AAV cis and trans plasmids and an adenovirus helper plasmid. This protocol describes the processes required to prepare the transfected cell suspension for virus purification.

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.

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.

In this protocol, DNA fragments are separated according to size by electrophoresis through low-melting-temperature agarose, and then recovered by melting the agarose and extracting with phenol:chloroform. The protocol works best for DNA fragments ranging in size from 0.5 to 5.0 kb. Yields of DNA fragments outside this range are usually lower, but often are sufficient for many purposes.

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.

For most immunoblots developed with chemiluminescence or with fluorochrome-based detection systems, it is possible to remove the primary and secondary antibodies from the membrane without affecting the bound antigen. This allows you to reuse the membrane for detection of another protein antigen. The blots developed with chromogenic substrates can also be stripped of antibodies and reprobed, but the bands detected in the first round of immunoblotting will remain unaffected. Stripping and reprobing of the membrane are particularly useful when the amount of sample is limited or when it is important to accurately compare the signal between two different protein antigens in the same sample. Examples of such experiments include determining the levels of a protein antigen in a series of samples relative to the loading control and comparison of the phosphorylated form to the total levels of the protein in the sample.

Before probing blots for the presence of an antigen, the total composition of the transferred proteins can be determined by staining the nitrocellulose or polyvinylidene fluoride (PVDF) membrane. Staining for proteins is useful to determine the position of the non-prestained molecular weight markers or individual lanes on the gel and to ensure that efficient transfer has occurred. It can be also used to verify equal loading of the samples in the gel when a comparison of the protein of interest between the different samples is important. The conventional procedures such as Coomassie Blue and silver staining methods used for staining polyacrylamide gels are incompatible with immunoblotting. Ponceau S is the more common staining method in immunoblotting protocols because it is compatible with antibody–antigen binding, is cost efficient, and provides a good contrast between the stained bands and background. In this protocol, nitrocellulose or PVDF membrane is rinsed with ultrapure H₂O after the transfer of proteins. Ponceau S dye is applied as an acidic aqueous solution, and the proteins on the membrane are stained with red color. The membrane is briefly destained with water and can be photographed or scanned to obtain the image of the total protein staining. Individual lane positions or the molecular weight standards can be marked with a pencil, if required.

The Bradford assay is a quick and fairly sensitive method for measuring the concentrations of proteins. It is based on the shift in absorbance maximum of Coomassie Brilliant Blue G-250 dye from 465 to 595 nm following binding to denatured proteins in solution.

Colloidal gold–antibody conjugates are easy to prepare and are an excellent choice for microscopic applications. Colloidal gold is an aqueous suspension of nanometer-sized particles of gold. Typically, chloroauric acid, HAuCl₄, is reduced with dilute solutions of sodium citrate, as described here. This will cause the gold to form small aggregates that will associate with proteins. Gold particles of specific sizes can be isolated and differentiated microscopically, allowing these particles to be used for multiple-label experiments. Colloidal gold-labeled antibodies are widely used in electron microscopy (EM), and can be used for light microscopy but require additional steps (silver enhancement).

There are many uses for antibodies labeled with metal ions. Most of these methods involve first attaching a metal chelator to the antibody molecule. This is achieved using standard cross-linking chemistry and then adding the desired metal at appropriate concentration and pH. The method described here outlines a basic procedure for creating a lanthanide conjugate. Lanthanide conjugates are used for proximity assays, as MRI contrast agents, or for mass cytometry experiments. Different metals and chelators can be substituted, but the basic procedures are similar.

Successful modification of the bacterial artificial chromosome (BAC) after two-step BAC engineering is confirmed in two separate polymerase chain reactions (PCRs). The first reaction (5' co-integrate PCR) uses a forward 5' co-integrate primer (a sequence located upstream of the 5' end of the A-box) and a reverse 3' primer on the vector (175PA+50AT) or within the reporter sequence or mutated region as appropriate. The second reaction (3' co-integrate PCR) uses a forward 5' primer on the recA gene (RecA1300S) and a reverse 3' co-integrate primer (a sequence located downstream from the 3' end of the B-box). Those colonies shown to be positive in PCR analysis are further tested for sensitivity to UV light. After the resolution, colonies that have lost the excised recombination vector including sacB and recA genes become UV light sensitive.

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.

Plasmid DNA is prepared from the recombinant shuttle vector pLD53.SCAB/A-B created by cloning of the A and B homology arms for two-step bacterial artificial chromosome (BAC) engineering. To confirm that the A-box and B-box arms have been successfully incorporated into pLD53.SCAB, the pattern of enzyme digestion of the modified plasmid is compared with that of the unmodified pLD53.SCAB. Once the shuttle vector is shown to carry the proper sequences, it is ready for transfer into the BAC host.

This protocol describes the preparation of the shuttle vector before its introduction into bacterial artificial chromosome (BAC) host cells for BAC two-step engineering. The homology arm sequences, prepared previously, are introduced by ligation into the digested shuttle vector DNA to provide sites for recombination within the BAC clone. Crude lysates of individual bacterial transformants serve as templates in polymerase chain reaction (PCR) analysis to confirm the presence of the homology arms in the recombinant shuttle vector.

The 700-bp A homology arm (A-box) and the 700-bp B homology arm (B-box) are amplified by polymerase chain reaction (PCR) using purified bacterial artificial chromosome (BAC) DNA as template for two-step BAC engineering. The resulting A-box PCR product contains an AscI site at its 5' end (the 5' primer incorporates an AscI site, and the 3' primer does not incorporate any restriction sites). The B-box PCR product contains an XmaI site at its 3' end (the 5' primer does not incorporate any restriction sites, and the 3' primer incorporates an XmaI site). The amplification products are then digested with the appropriate restriction endonucleases to render them suitable for cloning into the shuttle vector.

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 R6K origin, which requires the protein to replicate. PIR2 cells, expressing , 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.

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