Protein Purification and Analysis

Introduction

Information about the regulation of protein expression, protein modification, protein:protein interactions and protein function during different stages of cell development is needed to understand the development and physiology of organisms. This complex analysis of protein function is a major task facing scientists today. Although the field of proteomics was first described only as the study of proteins encoded by the genome, it has now expanded to include the function of all expressed proteins. Thus it is not just the study of all proteins expressed in a cell but also all protein isoforms and modifications, interactions, structure and high-order complexes (Tyers and Mann, 2003).

A fundamental step for studying individual proteins is the purification of the protein of interest. A variety of strategies have been developed for purifying proteins. These strategies address different requirements of downstream applications including scale and throughput. There are four basic steps required for protein purification: 1) cell lysis; 2) binding to a matrix; 3) washing; and 4) elution. Cell lysis can be accomplished a number of ways, including nonenzymatic methods (e.g., sonication or French press) or use of hydrolytic enzymes such as lysozyme or a detergent reagent such as FastBreak™ Cell Lysis Reagent (Cat.# V8571). FastBreak™ Cell Lysis Reagent offers a convenient format for the in-media lysis of E. coli cells expressing recombinant proteins without interfering with downstream purification of tagged proteins (Stevens and Kobs, 2004). In addition, the FastBreak™ Reagent requires only minor modifications to be used with mammalian and insect cell lines (Betz, 2004).

Affinity purification tags can be fused to any recombinant protein of interest, allowing fast, easy purification using the affinity properties of the tag (Nilsson et al. 1997). Certain tags are used because they encode an epitope that can be purified or detected by a specific antibody or because they enable simplified purification of a desired protein.

Since protein is directly involved in biological function, a great deal of emphasis has been placed on developing new tools for proteomic studies (Zhu et al. 2003). A number of methods are available for functional protein interaction studies. These include protein pull-down assays, yeast two-hybrid systems (Fields and Song, 1989; Chien et al. 1991) and mammalian two-hybrid systems (Giniger et al. 1985; Lin et al. 1988) to identify protein:protein interactions, as well as protein-chip technology, mass spectrometry, traditional one- or two-dimensional gel electrophoresis and ELISA for protein identification.

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Affinity Tags

Researchers often need to purify a single protein for further study. One method for isolating a specific protein is the use of affinity tags. Affinity purification tags can be fused to any recombinant protein of interest, allowing fast and easy purification following a procedure that is based on the affinity properties of the tag (Nilsson et al. 1997). Many different affinity tags have been developed to simplify protein purification (Terpe, 2002). Fusion tags are polypeptides, small proteins or enzymes added to the N- or C-terminus of a recombinant protein. The biochemical features of different tags influence the stability, solubility and expression of proteins to which they are attached (Stevens et al. 2001). Using expression vectors that include a fusion tag facilitates recombinant protein purification.

Polyhistidine

The most commonly used tag to purify and detect recombinant expressed proteins is the polyhistidine tag (Yip et al. 1989). Protein purification using polyhistidine tags relies on the affinity of histidine residues for immobilized metal such as nickel, which allows selective protein purification (Yip et al. 1989; Hutchens and Yip, 1990). This affinity interaction is believed to be a result of coordination of a nitrogen on the imidazole moiety of polyhistidine with a vacant coordination site on the metal. The metal is immobilized to a support through complex formation with a chelate that is covalently attached to the support.

Polyhistidine tags offer several advantages for protein purification. The small size of the polyhistidine tag renders it less immunogenic than other larger tags. Therefore, the tag usually does not need to be removed for downstream applications following purification. A large number of commercial expression vectors that contain polyhistidine are available. The polyhistidine tag may be placed on either the N- or C-terminus of the protein of interest. And finally, the interaction of the polyhistidine tag with the metal does not depend on the tertiary structure of the tag, making it possible to purify otherwise insoluble proteins using denaturing conditions.

Glutathione-S-Transferase

The use of the affinity tag glutathione-S-transferase (GST) is based on the strong affinity of GST for immobilized glutathione-covered matrices (Smith and Johnson, 1988). Glutathione-S-transferases are a family of multifunctional cytosolic proteins that are present in eukaryotic organisms (Mannervik and Danielson, 1988; Armstrong, 1997). GST isoforms are not normally found in bacteria; thus endogenous bacterial proteins don’t compete with the GST-fusion proteins for binding to purification resin. The 26kDa GST affinity tag enhances the solubility of many eukaryotic proteins expressed in bacteria.

HaloTag^® Protein Tag

Often times protein fusion tags are used to aid in expression of suitable levels of soluble protein as well as for purification. A unique protein tag, the HaloTag^® protein is engineered to enhance expression and solubility of recombinant proteins in E. coli. The HaloTag^® protein tag is a 34kDa, monomeric protein tag modified from Rhodococcus rhodochrous dehalogenase. The HaloTag^® protein was designed to bind rapidly and covalently with a unique synthetic linker to achieve an irreversible attachment. The synthetic linker may be attached to a variety of entities such as fluorescent dyes and solid supports, permitting labeling of fusion proteins in cell lysates for expression screening, and efficient capture of fusion proteins onto a purification resin (Figure 11.1).

For protein purification, the HaloTag^® Technology is compatible with many protein expression systems and can be applied to proteins expressed in E. coli, mammalian cells and cell-free systems. The lack of an endogenous equivalent of the HaloTag^® protein in mammalian cells minimizes the chances of detecting false positives or nonspecific interactions. The combination of covalent capture with rapid binding kinetics overcomes the equilibrium-based limitations associated with traditional affinity tags and enables efficient capture even at low expression levels. In addition, the highly stable HaloTag^® protein-ligand interaction permits boiling the protein complex in SDS sample buffer followed by SDS-PAGE analysis.

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Choosing the Right Protein Purification Strategy

Information about the regulation of protein expression, protein modification, protein:protein interactions and protein function during different stages of cell development is needed to understand the development and physiology of organisms. Thus, developing tools for studying proteins is critical. Protein purification is a fundamental step for studying individual proteins and protein complexes, and a variety of strategies for purifying proteins have been developed to address desired scale, throughput and downstream applications.

Protein purification can describe several different approaches with many purposes. The most obvious is to isolate pure proteins, which is the first step for determining protein activity or structure. A second approach is purifying proteins to study their interactions with other proteins, DNA or RNA. This approach can be used to test whether two proteins interact with each other, or to screen for a collection of proteins that interacts with a target protein. These approaches have important uses, but may require different techniques to achieve the best results.

Protein Purification

Proteins are an important class of biological macromolecules that maintain the structural and functional integrity of the cell. Many diseases are associated with protein malfunction. Researchers need to isolate pure proteins to be able to delve into the mechanistic aspects of protein function and design diagnostic and therapeutic tests and agents.

Cultured mammalian cells offer an environment well suited for producing properly folded and functional mammalian proteins with appropriate post-translational modifications (Geisse, 1996; Wum, 2004) . However, the low expression levels of recombinant proteins in cultured mammalian cells presents a challenge for their purification. As a result, attaining satisfactory yield and purity depends on highly selective and efficient capture of these proteins from the crude cell lysate.

Protein Complex Isolation

One challenge the field of proteomics faces is the ability to elucidate the function of proteins and to determine how they assemble into the complex networks that are responsible for key cellular processes. Surface-based proteomics requires general and facile methods for immobilizing proteins on solid surfaces in known orientations without disrupting protein structure or function. This immobilization must exhibit high binding capacity and minimal nonspecific adsorption. In addition, protein complex isolation without immobilization is necessary for a variety of downstream applications such as mass spectroscopy for the identification of protein partners and complementary labeling.

Analysis of protein:nucleic acid interactions reveals information that can be important for understanding mRNA regulation, chromosomal remodeling and transcription. Nucleic acid binding proteins are required for many processes in living organisms. Transcription factors play an important role in regulating transcription of DNA by binding to specific recognition sites on the chromosome, often at the gene’s promoter and by interacting with other proteins in the nucleus. This regulation is required for cell viability, differentiation and growth (Mankan, 2009; Gosh, 1998). The ability to detect and confirm the interaction of such proteins with various nucleic acid targets provides valuable information about the cell signaling cascades that govern the ability of a cell to divide, migrate, interact with its neighbors, develop and maintain specialized functions, and undergo apoptosis at the appropriate time.

Two common techniques used to detect the interaction of nucleic acid binding proteins with nucleic acids are the electrophoretic mobility shift assay (EMSA) and fluorescence anisotropy assay (Lane, 1992; LiCata, 2008). EMSA involves binding protein to a radiolabeled DNA probe followed by resolution on a polyacrylamide gel. Due to the increase in mass, protein:probe complexes migrate slower than free probe, allowing comparison of free versus bound probe. The specificity of such complexes is determined using competition experiments with unlabeled specific and nonspecific oligos. This method works best with purified protein and can be quite labor intensive, particularly when numerous samples are being processed. In fluorescence anisotropy, a DNA binding protein incubated with a fluorophore-labeled DNA substrate. The sample is excited with polarized light and the emitted light from the fluorophore is measured. Because a DNA:protein complex tumbles in solution more slowly than the unbound DNA sustrate, there is more emitted polarized light. This method also works best with purified protein and requires specialized equipment.

Protein chips have emerged as an approach for identification of protein:protein and protein:nucleic acid interactions (Hall, 2004; Hall, 2007; Hudson, 2006). Functional protein microarrays normally contain full-length functional proteins or protein domains bound to a solid surface. Fluorescently labeled DNA is used to probe the array and identify proteins that bind to that specific probe. Protein microarrays provide a method for high-throughput identification of DNA:protein interactions. Immobilized proteins can be used in protein pull-down assays to isolate protein binding partners in vivo (mammalian cells) or in vitro, or they can be evaluated for their enzymatic activity.

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Purification of Polyhistidine-Tagged Proteins

Rapid Purification of Polyhistidine-Tagged Proteins Using Magnetic Resins

There is a growing need for high-throughput protein purification methods. Magnetic resins enable affinity-tagged protein purification without the need for multiple centrifugation steps and sequential transfer of samples to multiple tubes. There are several criteria that define a good protein purification resin: minimal nonspecific protein binding, high binding capacity for the fusion protein and efficient recovery of the fusion protein. The MagneHis™ Protein Purification System meets these criteria, enabling purification of proteins with a broad range of molecular weights and different expression levels. The magnetic nature of the binding particles allows purification from crude lysates to be performed in a single tube. In addition, the system can be used on automated liquid-handling platforms for high-throughput applications.

MagneHis™ Protein Purification System

The MagneHis™ Protein Purification System (Cat.# V8500, V8550) uses paramagnetic precharged nickel particles (MagneHis™ Ni-Particles) to isolate polyhistidine-tagged protein directly from a crude cell lysate. Figure 11.2 shows a schematic diagram of the MagneHis™ Protein Purification System protocol. Using a tube format, polyhistidine-tagged protein can be purified on a small scale using less than 1ml of culture or on a large scale using more than 1 liter of culture. Samples can be processed in a high-throughput manner using a robotic platform such as the Beckman Coulter Biomek^® 2000 or Biomek^® FX or Tecan Freedom EVO^® instrument. Polyhistidine-tagged proteins can be purified under native or denaturing (2–8M urea or guanidine-HCl) conditions. The presence of serum in mammalian and insect cell culture medium does not interfere with purification. For more information and a detailed protocol, see Technical Manual #TM060.

Example Protocol for the MagneHis™ Protein Purification System for Bacterial Expression

Materials Required:
(see Composition of Solutions section)

• MagneHis™ Protein Purification System (Cat.# V8500, V8550) and protocol
• 37°C incubator for flasks/tubes
• shaker
• magnetic separation stand
• 1M imidazole solution (pH 8.0; for insect or mammalian cells or culture medium)
• additional binding/wash buffer (may be required if processing numerous insect cell, mammalian cell or culture medium samples)
• solid NaCl (for insect or mammalian cells or culture medium)

1. Add 110μl FastBreak™ Cell Lysis Reagent, 10X, to 1ml of fresh bacterial culture.
   
   
2. Resuspend DNase I as indicated on the vial. Add 1μl to the lysed culture.
   
   
3. Incubate with shaking for 10–20 minutes at room temperature.
   
   
4. Vortex the MagneHis™ Ni-Particles to a uniform suspension.
   
   
5. Add 30μl MagneHis™ Ni-Particles to 1.1ml of cell lysate.
   
   
6. Pipet to mix, and incubate for 2 minutes at room temperature.
   
   
7. Place the tube in the appropriate magnetic stand for approximately 30 seconds to capture the MagneHis™ Ni-Particles. Carefully remove supernatant.
   
   
8. Remove the tube from the magnet. Add 150μl of MagneHis™ Binding/Wash Buffer to the MagneHis™ Ni-Particles, and pipet to mix. Make sure that particles are resuspended well.
   
   
9. Place the tube in the appropriate magnetic stand for approximately 30 seconds to capture the MagneHis™ Ni-Particles. Carefully remove supernatant. Repeat Steps 8 and 9 for a total of three washes.
   
   
10. Add 100μl of MagneHis™ Elution Buffer, and pipet to mix. Incubate for 1–2 minutes at room temperature. Place the tube in the appropriate magnetic stand, and capture the particles. Carefully remove the supernatant, which now contains the fusion protein.
    
    

Purification using Denaturing Conditions. Proteins expressed in bacterial cells may be present in insoluble inclusion bodies. To determine if your protein is located in an inclusion body, perform the lysis step using FastBreak™ Cell Lysis Reagent, 10X, as described in Technical Manual #TM060. Pellet cellular debris by centrifugation, and check the supernatant and pellet for the polyhistidine-tagged protein by gel analysis. Insoluble proteins need to be purified under denaturing conditions. Since the interaction of polyhistidine-tagged fusion proteins and MagneHis™ Ni-Particles does not depend on tertiary structure, fusion proteins can be captured and purified using denaturing conditions by adding a strong denaturant such as 2–8M guanidine hydrochloride or urea to the cells. Denaturing conditions need to be used throughout the procedure; otherwise the proteins may aggregate. We recommend preparing denaturing buffers by adding solid guanidine-HCl or urea directly to the MagneHis™ Binding/Wash and Elution Buffers. For more detail, see Technical Manual #TM060.

Note: Do not combine FastBreak™ Cell Lysis Reagent and denaturants. Cells can be lysed directly using denaturants such as urea or guanidine-HCl.

Purification from Insect and Mammalian Cells. Process cells at a cell density of 2 × 10⁶ cells/ml of culture. Adherent cells may be removed from the tissue culture vessel by scraping and resuspending in culture medium to this density. Cells may be processed in culture medium containing up to 10% serum. Processing more than the indicated number of cells per milliliter of sample may result in reduced protein yield and increased nonspecific binding. For proteins that are secreted into the cell culture medium, cells should be removed from the medium prior to purification. For more detail, see Technical Manual #TM060.

MagZ™ Protein Purification System for Purification of Proteins Expressed in Rabbit Reticulocyte Lysate

Purification of a polyhistidine-tagged protein that has been expressed in rabbit reticulocyte lysate is complicated by hemoglobin in the lysate copurifying with the protein of interest. Hemoglobin copurification limits downstream applications (e.g., fluorescence-based functional assays, protein:protein interaction studies) and reduces the amount of protein purified. The MagZ™ Protein Purification System provides a simple, rapid and reliable method to purify expressed polyhistidine-tagged protein from rabbit reticulocyte lysate. Paramagnetic precharged particles can be used to isolate polyhistidine-tagged protein from 50–500μl of TNT^® Rabbit Reticulocyte Lysate with minimal copurification of hemoglobin. These polyhistidine-tagged proteins are 99% free of contaminating hemoglobin.

The MagZ™ System is flexible enough to be used with different labeling and detection methods. Polyhistidine-tagged proteins expressed in rabbit reticulocyte lysate can be labeled with [³⁵S]methionine or the FluoroTect™ Green_Lys in vitro Translation Labeling System. FluoroTect™-labeled polyhistidine-tagged proteins can be visualized by gel analysis and analyzed using a FluorImager^® instrument. Figure 11.3 shows a schematic diagram of the MagZ™ Protein Purification System protocol. For more detail, see Technical Bulletin #TB336.

Materials Required:
(see Composition of Solutions section)

• MagZ™ Protein Purification System (Cat.# V8830) and protocol
• platform shaker or rocker, rotary platform or rotator
• MagneSphere^® Technology Magnetic Separation Stand (Cat.# Z5331, Z5332, Z5341, Z5342)

Medium- to Large-Scale Purification of Polyhistidine-Tagged Proteins In Column or Batch Formats

The two most common support materials for resin-based, affinity-tagged protein purification are agarose and silica gel. As a chromatographic support, silica is advantageous because it has a rigid mechanical structure that is not vulnerable to swelling and can withstand large changes in pressure and flow rate without disintegrating or deforming. Silica is available in a wide range of pore and particle sizes including macroporous silica, providing a higher capacity for large biomolecules such as proteins. However, two of the drawbacks of silica as a solid support for affinity purification are the limited reagent chemistry that is available and the relatively low efficiency of surface modification.

The HisLink™ Protein Purification Resin (Cat.# V8821) and HisLink™ 96 Protein Purification System (Cat.# V3680, V3681) overcome these limitations using a new modification process for silica surfaces that provides a tetradentate metal-chelated solid support with a high binding capacity and concomitantly eliminates the nonspecific binding that is characteristic of unmodified silica. HisLink™ Resin is a macroporous silica resin modified to contain a high level of tetradentate-chelated nickel (>20mmol Ni/ml settled resin). Figure 11.4 show a schematic diagram of HisLink™ Resin and polyhistidine-tag interaction. The HisLink™ Resin has a pore size that results in binding capacities as high as 35mg of polyhistidine-tagged protein per milliliter of resin.

The HisLink™ Resin enables efficient capture and purification of bacterially expressed polyhistidine-tagged proteins. This resin may also be used for general applications that require an immobilized metal affinity chromatography (IMAC) matrix (Porath et al. 1975; Lonnerdal and Keen, 1982). HisLink™ Resin may be used in either column or batch purification formats. For a detailed protocol see Technical Bulletin #TB327.

Column-Based Purification using HisLink™ Resin

The HisLink™ Resin provides a conventional means to purify polyhistidine-tagged proteins and requires only a column that can be packed to the appropriate bed volume. If packed to 1ml under gravity-driven flow, HisLink™ Resin shows an average flow rate of approximately 1ml/minute. In general a flow rate of 1–2ml/minute per milliliter of resin is optimal for efficient capture of polyhistidine-tagged protein. Gravity flow of a cleared lysate over a HisLink™ column will result in complete capture and efficient elution of polyhistidine-tagged proteins; however, the resin may also be used with vacuum filtration devices (e.g., Vac-Man^® Vacuum Manifold, Cat.# A7231) to allow simultaneous processing of multiple columns. HisLink™ Resin is also an excellent choice for affinity purification using low- to medium-pressure liquid chromatography systems such as fast performance liquid chromatography (FPLC).

Example Protocol Using the HisLink™ Resin to Purify Proteins from Cleared Lysate by Gravity-Flow Column Chromatography

Materials Required:
(see Composition of Solutions section)

• HisLink™ Protein Purification Resin (Cat.# V8821) and protocol
• HEPES buffer (pH 7.5)
• imidazole
• binding buffer
• wash buffer
• elution buffer
• column [e.g., Fisher PrepSep Extraction Column (Cat.# P446) or Bio-Rad Poly-Prep^® Chromatography Column (Cat.# 731-1550)]

Cell Lysis: Cells may be lysed using any number of methods including sonication, French press, bead milling, treatment with lytic enzymes (e.g., lysozyme) or use of a commercially available cell lysis reagent such as the FastBreak™ Cell Lysis Reagent (Cat.# V8571). If lysozyme is used to prepare a lysate, add salt (>300mM NaCl) to the binding and wash buffers to prevent the lysozyme binding to the resin. Finally, adding protease inhibitors such as 1mM PMSF to cell lysates does not inhibit binding or elution of polyhistidine-tagged proteins with the HisLink™ Resin and is highly recommended to prevent degradation of protein of interest by endogenous proteases. When preparing cell lysates from high-density cultures, adding DNase and RNase (concentrations up to 20μg/ml) will reduce the lysate viscosity and aid in purification.

Example Protocol

1. Prepare the binding, wash and elution buffers.
   
   
2. Determine the column volume required to purify the protein of interest. In most cases 1ml of settled resin is sufficient to purify the amount of protein typically found in up to 1 liter of culture (cell density of O.D.₆₀₀ < 6.0). In cases of very high expression levels (e.g., 50mg protein/liter), up to 2ml of resin per liter of culture may be needed.
   
   
3. Once you have determined the volume of settled resin required, precalibrate this amount directly in the column by pipetting the equivalent volume of water into the column and marking the column to indicate the top of the water. This mark indicates the top of the settled resin bed. Remove the water before adding resin to the column.
   
   
4. Make sure that the resin is fully suspended; fill the column with resin to the line marked on the column by transferring the resin with a pipette. Allow the resin to settle, and adjust the level of the resin by adding or removing resin as necessary.

   Note: If the resin cannot be pipetted within 10–15 seconds of mixing, significant settling will occur, and the resin will need to be resuspended. Alternatively, a magnetic stir bar may be used to keep the resin in suspension during transfer. To avoid fracturing the resin, do not leave the resin stirring any longer than the time required to pipet and transfer the resin.

   
   
   
5. Allow the column to drain, and equilibrate the resin with five column volumes of binding buffer, allowing the buffer to completely enter the resin bed.
   
   
6. Gently add the cleared lysate to the resin until the lysate has completely entered the column. The rate of flow through the column should not exceed 1–2ml/minute for every 1ml of column volume. Under normal gravity flow conditions the rate is typically about 1ml/minute. The actual flow rate will depend on the type of column used and the extent to which the lysate was cleared and filtered. Do not let the resin dry out after you have applied the lysate to the column.
   
   
7. Wash unbound proteins from the resin using at least 10–20 column volumes of wash buffer. Divide the total volume of wash buffer into two or three aliquots, and allow each aliquot to completely enter the resin bed before adding the next aliquot.
   
   
8. Once the wash buffer has completely entered the resin bed, add elution buffer and begin collecting fractions (0.5–5ml fractions). Elution profiles are protein-dependent, but polyhistidine-tagged proteins will generally elute in the first 1ml. Elution is usually complete after 3–5ml of buffer have been collected per 1.0ml of settled resin, provided the imidazole concentration is high enough to efficiently elute the protein of interest.
   
   

Batch Purification Using HisLink™ Resin

One of the primary advantages of the HisLink™ Resin is its use in batch purification. In batch mode, the protein of interest is bound to the resin by mixing lysate with the resin for approximately 30 minutes at a temperature range of 4–22°C. Once bound with protein, the resin is allowed to settle to the bottom of the container, and the spent lysate is poured off. Washing only requires resuspension of the resin in an appropriate wash buffer followed by a brief period to allow the resin to settle. The wash buffer is then carefully poured off. This process is repeated as many times as desired. Final elution is best achieved by transferring the HisLink™ Resin to a column to elute the protein in fractions. The advantages of batch purification are: 1) less time is required to perform the purification; 2) large amounts of lysate can be processed; and 3) clearing the lysate prior to purification is not required.

Purification of Polyhistidine-Tagged Proteins by FPLC

The rigid particle structure of the silica base used in the HisLink™ Resin make this material an excellent choice for applications that require applied pressure to load the lysate, wash or elute protein from the resin. These applications involve both manual and automated systems that operate under positive or negative pressure (e.g., FPLC and vacuum systems, respectively). To demonstrate the use of HisLink™ Resin on an automated platform we used an AKTA explorer from GE Healthcare. Milligram quantities of polyhistidine-tagged protein were purified from one liter of culture. The culture was lysed in 20ml of binding/wash buffer and loaded onto a column containing 1ml of HisLink™ Resin. We estimate the total amount of protein recovered to be 75–90% of the protein expressed in the original lysate.

Purification under denaturing conditions: Proteins that are expressed as an inclusion body and have been solubilized with chaotrophic agents such as guanidine-HCl or urea can be purified by modifying the protocols to include the appropriate amount of denaturant (up to 6M guanidine-HCl or up to 8M urea) in the binding, wash and elution buffers.

Adjuncts for lysis or purification: The materials shown in Table 11.1 may be used without adversely affecting the ability of HisLink™ Resin to bind and elute polyhistidine-tagged proteins.

Table 11.1. Additives That Will Not Affect Binding or Elution of Polyhistidine-Tagged Proteins Using HisLink™ Resin.
Additive	Concentration
HEPES, Tris or sodium phosphate buffers	≤100mM
NaCl	≤1M
β-mercaptoethanol	≤100mM
DTT	≤10mM
Triton® X-100	≤2%
Tween®	≤2%
glycerol	≤20%
guanidine-HCl	≤6M
urea	≤8M
RQ1 RNase-Free DNase	≤5μl/1ml original culture

96-Well Format For Purification of Polyhistidine-Tagged Proteins

The HisLink™ 96 Protein Purification System (Cat.# V3680, V3681) uses a vacuum-based method to purify polyhistidine-tagged expressed proteins directly from E. coli cultures grown in deep-well, 96-well plates. The HisLink™ 96 System is amenable to manual or automated methods for high-throughput applications. In preparation for protein purification, bacterial cells expressing a polyhistidine-tagged protein are lysed directly in culture using the provided FastBreak™ Cell Lysis Reagent. The HisLink™ Resin is added directly to the lysate and mixed, and the polyhistidine-tagged proteins bind within 30 minutes. The samples are then transferred to a filtration plate. Unbound proteins are washed away, and the target protein is recovered by elution. Figure 11.5 describes protein purification using the HisLink™ 96 System. This system requires the use of the Vac-Man^® 96 Vacuum Manifold (Cat.# A2291, Figure 11.6) or a compatible vacuum manifold. For more detailed protocol information, see Technical Bulletin #TB342.

Manual Protocol

Materials Required:
(see Composition of Solutions section)

• HisLink™ 96 Protein Purification System (Cat.# V3680, V3681) and protocol
• Nuclease-Free Water (Cat.# P1195)
• Vac-Man^® 96 Vacuum Manifold (Cat.# A2291)
• plate shaker (manual) or multichannel pipette
• wide-bore tips (Racked, Sterile, Yellow Lift Top Racks; E&K Scientific Cat.# 3502-R96S)
• 96-well, deep-well plates (e.g., ABgene 2.2ml storage plate, Marsh Bio Products Cat.# AB-0932)
• 96-well sealing mats (Phenix Research Products Cat.# M-0662)
• 96-well plate adhesive sealers
• reservoir boats (Diversified Biotech Cat.# RESE-3000)

Automated Purification

The manual protocol described in Section III.C can be used as a guide to develop protocols for automated workstations. The protocol may require optimization, depending on the instrument used.

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Purification of GST-Tagged Proteins

Rapid Purification of GST-Tagged Proteins Using Magnetic Resins

There is a growing need for protein purification methods that are amenable to high-throughput screening. Magnetic resins enable affinity-tagged protein purification without the need for multiple centrifugation steps and transfer of samples to multiple tubes. There are several criteria that define a good protein purification resin: minimal nonspecific protein binding, high binding capacity for the fusion protein and efficient recovery of the fusion protein. The MagneGST™ Protein Purification System (Cat.# V8600, V8603) meets these criteria, enabling purification of proteins with a broad range of molecular weights and different expression levels. The magnetic nature of the binding particles allows purification from a crude lysate in a single tube. In addition, the system can be used with automated liquid-handling platforms for high-throughput applications.

MagneGST™ Protein Purification System for Purification of GST-Tagged Proteins

The MagneGST™ Protein Purification System provides a simple, rapid and reliable method to purify glutathione-S-transferase (GST) fusion proteins. Glutathione immobilized on paramagnetic particles (MagneGST™ Glutathione Particles; Cat.# V8611, V8612) is used to isolate GST-fusion proteins directly from a crude cell lysate using a manual or automated procedure. The use of paramagnetic particles eliminates several centrifugation steps and the need for multiple tubes. It also minimizes the loss of sample material. Although the MagneGST™ System is designed for manual applications, samples can also be processed using a robotic platform, such as the Beckman Coulter Biomek^® 2000 or Biomek^® FX workstation, for high-throughput applications. Visit the Promega web site for more information about using the MagneGST™ System in an automated format.

Bacterial cells containing a GST-fusion protein are lysed using the provided MagneGST™ Cell Lysis Reagent or an alternative lysis method, and the MagneGST™ Particles are added directly to the crude lysate. GST-fusion proteins bind to the MagneGST™ Particles. Unbound proteins are washed away, and the GST-fusion target protein is recovered by elution with 50mM glutathione. Figure 11.7 shows a schematic diagram of the MagneGST™ Protein Purification System protocol. For more detailed information about the protocol, see Technical Manual #TM240.

Additionally, we have used the MagneGST™ Particles to purify GST-fusion protein generated in vitro using the E. coli S30 Extract System for Circular DNA (Cat.# L1020). When eluted protein was analyzed by SDS polyacrylamide gel electrophoresis, no major contaminating proteins were found to copurify with the GST-fusion proteins.

Example Protocol for the MagneGST™ Protein Purification System

Materials Required:
(see Composition of Solutions section)

• MagneGST™ Protein Purification System (Cat.# V8600, V8603) and protocol
• 1.5ml microcentrifuge tubes for small-scale protein purifications or 15ml or 50ml conical tubes for large-scale protein purifications
• magnetic separation stand
• RQ-1 RNase-Free DNase (Cat.# M6101)
• shaker or rotating platform
• centrifuge

Cell Lysis

1. Prepare cell pellets from 1ml of bacterial culture.
   
   
2. Add 200μl of MagneGST™ Cell Lysis Reagent to each fresh or frozen cell pellet. Resuspend the cell pellet at room temperature (20–25°C) by pipetting or gentle mixing.
   
   
3. Add 2μl of RQ1 RNase-Free DNase.
   
   
4. Incubate the cell suspension at room temperature for 20–30 minutes on a rotating platform or shaker.
   
   

Equilibrate Particles

1. Thoroughly resuspend the MagneGST™ Particles by inverting the bottle to obtain a uniform suspension.
   
   
2. Pipet 100μl of MagneGST™ Particles into a 1.5ml tube.
   
   
3. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Magnetic capture of the particles will typically occur within a few seconds.
   
   
4. Carefully remove and discard the supernatant.
   
   
5. Remove the tube from the magnetic stand. Add 250μl of MagneGST™ Binding/Wash Buffer to the particles, and resuspend by pipetting or inverting.
   
   
6. Repeat Particle Equilibration Steps 3–5 twice for a total of three washes.
   
   

Bind Proteins

1. After the final wash, gently resuspend the particles in 100μl of MagneGST™ Binding/Wash Buffer.
   
   
2. Add 200μl of cell lysate, prepared as described above, to the particles.
   
   
3. Mix gently by pipetting or inverting. If the combined volume of cell lysate and MagneGST™ Particles is less than 300μl, add additional MagneGST™ Binding/Wash Buffer so that the final volume is 300μl.
   
   
4. Incubate with gentle mixing on a rotating platform or shaker for 30 minutes at 4°C or room temperature.
   
   

Wash

1. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet.
   
   
2. Carefully remove the supernatant. Save the supernatant (flowthrough) for SDS-PAGE analysis, if desired.
   
   
3. Remove the tube from the magnetic stand. Add 250μl of MagneGST™ Binding/Wash Buffer to the particles, and mix gently by pipetting or inverting. Incubate at room temperature or 4°C for 5 minutes. Occasionally mix by inverting the tube.
   
   
4. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet.
   
   
5. Carefully remove the supernatant. Save the supernatant if analysis of wash solution is desired.
   
   
6. Remove the tube from the magnetic stand. Add 250μl of MagneGST™ Binding/Wash Buffer to the particles, and mix gently by pipetting or inverting. Incubation is not necessary at this step.
   
   
7. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet.
   
   
8. Carefully remove the supernatant. Save the supernatant if analysis of wash solution is desired.
   
   
9. Repeat Wash Steps 6–8 once for a total of three washes.
   
   

Elution

1. After the final wash, add 200μl of elution buffer.
   
   
2. Incubate at room temperature or 4°C for 15 minutes with gentle mixing.
   
   
3. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet.
   
   
4. Carefully remove the supernatant, and transfer it to a clean tube. The supernatant contains the eluted GST-fusion protein.
   
   
5. If a second elution is desired, repeat Elution Steps 1–4.
   
   

Compatibility with Common Buffer Components: The MagneGST™ Particles have been shown to be compatible with many common buffer components (Table 11.2).

Table 11.2. Buffer Components Compatible with the MagneGST™ Particles.
Buffer Component	Concentration
DTT	≤10mM
NaCl	≤0.64M
Tris, HEPES, sodium phosphate, potassium phosphate	≤100mM
Triton® X-100	≤1%
Tween®	≤1%
MAZU	≤1%
cetyltrimethylammonium bromide (CTAB)	≤1%
ethanol	20%
protease inhibitor cocktail (Roche Molecular Systems, Inc. Cat.# 1836170)	1X

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Purification of HaloTag^® Fusion Proteins

HaloTag^® Protein Purification from Mammalian Cells

Cultured mammalian cells offer an environment well suited for producing properly folded and functional mammalian proteins with appropriate post translational modifications. However, the low expression levels of recombinant proteins in cultured mammalian cells presents a challenge for their purification. As a result, attaining satisfactory yield and purity depends on selective and efficient capture of these proteins from the crude cell lysate. The equilibrium-based binding of most affinity tag protein purification methods means that the protein is constantly being exchanged between the bound (to the resin) and unbound state. This equilibrium depends on the protein concentration and the binding affinity of the tag. As a result, binding efficiency may be reduced at low expression levels, leading to low recovery of the fusion protein.

The HaloTag^® Mammalian Protein Detection and Purification Systems (Cat.# G6795) utilize the HaloTag^® protein tag, which can be genetically fused to any protein and transiently or stably expressed in mammalian cells. Following cell lysis, the HaloTag^® fusion protein is covalently captured on the HaloLink™ Resin, and nonspecific proteins are washed away. The protein of interest is then released by a specific proteolytic cleavage at an optimized TEV recognition site contained within the amino acid linker sequence that connects the HaloTag^® protein tag and the protein of interest. The use of a TEV protease fused to HaloTag^® (HaloTEV; Cat.# G6601), which is covalently captured on the HaloLink™ Resin, eliminates the need for a secondary step to remove the protease, resulting in a streamlined purification process. This straightforward purification uses a single, mild physiological buffer throughout the entire process with no need for buffer exchange (Figure 11.8).

HaloTag^® Protein Purification from E. Coli

The HaloTag^® Protein Purification System (Cat.# G6280) allows covalent, efficient and specific capture of the proteins expressed in E. coli as N-terminal HaloTag^® fusion proteins. Many of the same characteristics that make the Halotag^® protein well-suited for purifying proteins from mammalian cells also make it a good choice for purifying proteins from E. coli cells. The choice of Flexi^® expression vectors is more limited for E. coli expression, with the appropriate vectors that encodes the HaloTag^® protein and expresses protein optimally in E. coli being the pFN18A HaloTag^® T7 Flexi^® Vector (Cat.# G2751) or the pFN18K HaloTag^® T7 Flexi^® Vector (Cat.# G2681). Alternatively, non-Flexi^® system vectors are available with dual tags of HaloTag^® protein and Polyhistidine. These vectors, pH6HTN His₆HaloTag^® T7 Vector (Cat.# G7971) or the pH6HTC His₆HaloTag^® T7 Vector (Cat.# G8031), allow traditional cloning using the multiple cloning site. These dual-tagged vectors enable purification of HaloTag^®-fused proteins that still retain the covalent coupling ability of the HaloTag^® protein. With the HaloTag^® Protein Purification System, it is easy to perform in-gel detection and quantification of protein expression levels using fluorescent HaloTag^® Ligands.

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Purification of Biotinylated Proteins

PinPoint™ Xa System and SoftLink™ Resin for Purification of Biotinylated Protein

Biotinylated fusion proteins such as those produced with the PinPoint™ Xa Protein Purification System (Cat.# V2020) can be affinity-purified using the SoftLink™ Soft Release Avidin Resin (Cat.# V2011). This proprietary resin allows elution of a fusion protein under native conditions by adding exogenous biotin.

The PinPoint™ Xa Protein Purification System is designed to produce and purify fusion proteins that are biotinylated in vivo. The biotinylation reaction in E. coli is catalyzed by biotin ligase holoenzyme and results in a fusion purification tag that carries a single biotin specifically on one lysine residue (Wilson et al. 1992; Xu and Beckett, 1994; Cronan, 1990). The biotin moiety is accessible to avidin or streptavidin, as demonstrated by binding to resins containing either molecule, and serves as a tag for detection and purification. E. coli produces a single endogenous biotinylated protein that, in its native conformation, does not bind to avidin, rendering the affinity purification highly specific for the recombinant fusion protein.

The system contains vectors in all possible reading frames, an avidin-conjugated resin, Streptavidin-Alkaline Phosphatase, a purification column and biotin. The PinPoint™ Xa Control Vector contains the chloramphenicol acetyltransferase (CAT) gene and is provided as a means of monitoring protein expression, purification and processing conditions. The PinPoint™ Vectors feature the encoded endoproteinase Factor Xa proteolytic site that provides a way to separate the purification tag from the native protein. These vectors also carry a convenient multiple cloning region for ease in construction of fusion proteins.

Biotinylated proteins synthesized using the PinPoint™ Xa System can be affinity-purified using the SoftLink™ Soft Release Avidin Resin. Avidin-biotin interactions are so strong that elution of biotin-tagged proteins from avidin-conjugated resins usually requires denaturing conditions. In contrast, the SoftLink™ Soft Release Avidin Resin, which uses monomeric avidin, allows the protein to be eluted with a nondenaturing 5mM biotin solution. The rate of dissociation of the monomeric avidin-biotin complex is sufficiently fast to effectively allow recovery of all bound protein in neutral pH and low salt conditions. The diagram in Figure 11.9 outlines the expression and purification system procedure.

The SoftLink™ Soft Release Avidin Resin is highly resistant to many chemical reagents (e.g., 0.1N NaOH, 50mM acetic acid and nonionic detergents), permitting stringent wash conditions.

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Protein:Protein Interaction Analysis: In Vivo and In Vitro Methods

Determining the protein:protein interaction map (“interactome”) of the whole proteome is one major focus of functional proteomics (Li et al. 2004; Huzbun et al. 2003). Various methods have been used for studying protein:protein interactions, including yeast, bacterial and mammalian two- and three-hybrid systems, immunoaffinity purifications, affinity tag-based methods and mass spectrometry (reviewed in Li et al. 2004; Huzbun et al. 2003; Zhu et al. 2003). Moreover, in vitro pull-down-based techniques such as tandem affinity purification (TAP) are being widely used for isolating protein complexes (Forler et al. 2003).

In vitro protein pull-down assays can be performed using cell lysates, cell-free lysates, tissue samples, etc. These options are not possible with two-hybrid approaches. There are several reports describing the use of in vitro pull-down assays for analyzing protein:protein interactions using proteins translated in vitro using cell-free expression systems such as rabbit reticulocyte lysate-based expression systems (Charron et al. 1999; Wang et al. 2001; Pfleger et al. 2001). Cell-free expression is a powerful method for expressing cDNA libraries. This technique is also amenable to high-throughput protein expression and identification. Cell-free expression systems, especially rabbit reticulocyte lysate-based methods, have been extensively used for in vitro pull-down assays because of the ease of performing these experiments (Charron et al. 1999; Wang et al. 2001; Pfleger et al. 2001). There are also reports describing high-throughput identification of protein:protein interactions using TNT^® Rabbit Reticulocyte Lysate (Pfleger et al. 2001).

Mammalian Two-Hybrid Systems

Two-hybrid systems are powerful methods to detect protein:protein interactions in vivo. The basis of two-hybrid systems is the modular nature of some transcription factor domains: a DNA-binding domain, which binds to a specific DNA sequence, and a transcriptional activation domain, which interacts with the basal transcriptional machinery (Sadowski et al. 1988). A transcriptional activation domain in association with a DNA-binding domain promotes the assembly of RNA polymerase II complexes at the TATA box and increases transcription. In the CheckMate™ Mammalian Two-Hybrid System (Cat.# E2440), the DNA-binding domain and transcriptional activation domain, produced by separate plasmids, are closely associated when one protein (“X”) fused to a DNA-binding domain interacts with a second protein (“Y”) fused to a transcriptional activation domain. In this system, interaction between proteins X and Y results in transcription of a reporter gene or selectable marker gene (Figure 11.10).

Originally developed in yeast (Fields and Song, 1989; Chien et al. 1991), the two-hybrid system has been adapted for use in mammalian cells (Dang et al. 1991; Fearon et al. 1992). One major advantage of the CheckMate™ Mammalian Two-Hybrid System over yeast systems is that the protein:protein interaction can be studied in the cell line of choice. The CheckMate™ System also uses the Dual-Luciferase^® Reporter Assay System for rapid and easy quantitation of luciferase reporter gene expression.

Application of the CheckMate™ Mammalian Two-Hybrid System confirms suspected interactions between two proteins and identifies residues or domains involved in protein:protein interactions. When identifying residues or domains involved in an interaction, the GeneEditor™ in vitro Site-Directed Mutagenesis System (Cat.# Q9280) for making site-directed mutants and Erase-a-Base^® Systems for deletion analysis are useful tools. These products are fully compatible with the CheckMate™ Mammalian Two-Hybrid System. Detailed protocol information is available in Technical Manual #TM049.

Assessing Protein:Protein Interactions

cDNA sequences encoding the polypeptides of interest are subcloned into pBIND and pACT Vectors. The insert in each vector must be in the correct orientation and reading frame. See the CheckMate™ System Technical Manual #TM049 for the multiple cloning region following the 3′ end of the GAL4 fragment for pBIND Vector and for the multiple cloning region following the 3′ end of the VP16 fragment for pACT Vector. All vectors in the CheckMate™ Mammalian Two-Hybrid System confer ampicillin resistance and are compatible with E. coli strains such as JM109. We strongly recommend sequencing the 5′ junction between the insert and vector to ensure that the insert is subcloned properly. The T7 EEV Promoter Primer (Cat.# Q6700) can be used for sequence verification.

Certain inserts appear to show vector “directionality” (or preference) in which the interaction between a pair of proteins is fusion vector-dependent (Finkel et al. 1993). Protein:protein interactions may appear stronger given a particular vector context for the inserts. Because of this phenomenon, we advise subcloning each cDNA of interest into both the pBIND and pACT Vectors and testing the two possible fusion protein interactions.

Following the successful subcloning of the test cDNAs into the pBIND and pACT Vectors, the resultant plasmids should be purified such that the DNA is free of protein, RNA and chemical contamination. Before completing any experiments with the CheckMate™ System, optimize the transfection method for the cell type being transfected. The optimization process is easier using a reporter gene and assay system. Many DNA delivery agents exist for transfecting mammalian cells. Transfection of DNA into mammalian cells may be mediated by cationic lipids, calcium phosphate, DEAE-dextran or electroporation. Transfection systems based on cationic lipids (TransFast™ Transfection Reagent, Transfectam^® Reagent, Tfx™-20, Tfx™-50 Reagents) and calcium phosphate (ProFection^® Mammalian Transfection System) are available from Promega. The efficiency of each transfection method is highly dependent upon the cell type. When optimizing a transfection method for a particular cell type, use a reporter gene such as the firefly luciferase gene whose activity is easily and rapidly assayed. The pGL3-Control Vector (Cat.# E1741) expresses the firefly luciferase gene from the SV40 early promoter.

Table 11.3 presents the recommended combinations of vectors to properly control an experiment when using the CheckMate™ System to determine the extent to which two proteins interact in a two-hybrid assay.

Table 11.3. Recommended Experimental Design to Determine the Magnitude of Interaction Between Two Proteins.
Transfection	pBIND Vector	pACT Vector	pG5luc Vector
1	pBIND Vector	pACT Vector	pG5luc Vector
2	pBIND-Id Control Vector	pACT-MyoD Control Vector	pG5luc Vector
3	–	–	–
4	pBIND-X Vector	pACT Vector	pG5luc Vector
5	pBIND Vector;	pACT-Y Vector	pG5luc Vector
6	pBIND-X Vector;	pACT-Y Vector	pG5luc Vector

The amount of vector DNA to use will depend upon the method of transfection. However, we recommend that the molar ratio of pBIND:pACT Vector constructs be 1:1. We have varied the amount of pG5luc Vector in the positive control experiment and have found that the signal-to-noise ratio of firefly luciferase expression does not differ significantly. We routinely use a molar ratio of 1:1:1 for pBIND:pACT:pG5luc Vector in the CheckMate™ Mammalian Two-Hybrid System. Maintain a constant amount of DNA for each transfection reaction within an experiment by adding plasmid DNA such as pGEM^®-3Zf(+) Vector (Cat.# P2271).

We recommend testing a specific cell line with positive and negative control transfection reactions before initiating test experiments. The pBIND Vector encodes the Renilla luciferase gene to normalize for transfection efficiency. Replication of pBIND and pACT Vectors and their recombinants is expected in COS cells or other types of cells that express the SV40 large T antigen. Use the Dual-Luciferase^® Reporter Assay System (Cat.# E1910) to quantitate Renilla luciferase and firefly luciferase activities.

HaloTag^® Pull-Down Assays

Traditional protein pull-down approaches rely upon binding to an affinity resin, and often this is not a very efficient process. The HaloTag^® system is similar in that it is based on a protein fusion tag, but its rapid, covalent and irreversible binding sets it apart from other affinity tags. These properties increase the chances of capturing protein complexes and retaining them after capture. In addition, the lack of an endogenous equivalent of the HaloTag^® protein in mammalian cells minimizes the chances of detecting false positives or nonspecific interactions. An overview of the Halotag^® Mammalian Protein Pull-Down System (Cat.# G6500, G6504) is depicted in Figure 11.11. More information and detailed protocol information is available in Technical Manual #TM342.

HaloTag^® Mammalian Pull-Down Protocol

Materials Required:
(see Composition of Solutions section)

• HaloTag^® Mammalian Pull-Down System (Cat.# G6500, G6504) and protocol
• vector encoding HaloTag^® fusion protein (Cat.# G9651, G9661, G1611, G1601, G1591, G1571, G1551, G1321, G2821, G2831, G2841, G2851, G2861, G2871, G2881 or G2981) in the form of transfection-grade DNA
• HaloTag^® Control Vector (Cat.# G6591) in the form of transfection-grade DNA
• cells for transfection or a stable cell line expressing the desired HaloTag^® fusion protein
• cellular growth media
• transfection reagents
• PBS - tissue culture certified
• ethanol
• IGEPAL^® CA-630 (Sigma Cat.# 18896)
• rotating or shaking platform
• microcentrifuge
• cell culture incubator
• glass homogenizer (e.g., 2ml Kontes Dounce Tissue Grinder; Thermo Fisher Scientific Cat.# K885300-0002) or 25- to 27-gauge needle
• disposable cell lifter (e.g., Thermo Fisher Scientific Cat.# 08-773-1)

Phase 1. Equilibrate Resin

1. Mix the HaloLink™ Resin by inverting the bottle to obtain a uniform suspension.
   
   
2. For each pull-down experiment, dispense 200µl of HaloLink™ Resin into two 1.5ml microcentrifuge tubes (one tube for the experimental sample and one tube for the negative control sample). Centrifuge for 1 minute at 800 × g. Carefully remove and discard the supernatant, leaving the resin at the bottom of the tube.
   
   
3. Add 800µl of Resin Equilibration/Wash Buffer. Mix thoroughly by inverting the tube. Centrifuge for 2 minutes at 800 × g. Remove and discard the supernatant, leaving the resin at the bottom of the tube. Repeat this wash step twice more for a total of 3 washes. Do not remove the final wash supernatant until you are ready to bind lysates (Phase 3). This will prevent the resin from drying out.
   
   

Phase 2. Prepare HaloTag^® Fusion Protein and Control Lysates

1. For each sample, grow approximately 1–1.2 × 10⁷ cells.
   
   

   Note: If your protein shows extremely low or high expression, you may need to adjust the amount of starting cells up or down by a factor of two- to fivefold.

2. Add 25–30ml of ice-cold PBS to the cells, and gently scrape to collect cells into conical tubes. Centrifuge the cells at 4°C for 5–10 minutes at 2,000 × g, and discard the PBS. Store the cell pellets at –80°C for at least 30 minutes prior to lysing.
   
   
3. Thaw the frozen cell pellet, and resuspend the cells in 300µl of Mammalian Lysis Buffer; pipette or briefly vortex to mix.
   
   

   Note: Mammalian Lysis Buffer is optimized for total cellular lysis, including the nucleus. If cytoplasmic and nuclear fractions need to remain as separate pools, perform an initial cytoplasmic lysis. If you are processing the cytosolic fraction only, discard the nuclear pellets. If you will be processing both the cytosolic and nuclear fractions, the nuclear pellets can be lysed subsequently with the Mammalian Lysis Buffer as described here. If you know your complex requires certain cofactors or small molecules to maintain complex integrity, please add these to the Mammalian Lysis Buffer and the wash buffer.

4. Add 6µl of the 50X Protease Inhibitor Cocktail (Cat.# G6521) and incubate on ice for 5 minutes.
   
   
5. To reduce lysate viscosity following the incubation, homogenize with a Dounce glass homogenizer (2ml size) on ice using 25–30 strokes using the large pestle (B). Alternatively, pass the cells through a 25- or 27-gauge needle 5–10 times.
   
   

   Note: We do not recommend sonication because protein complexes may fall apart, and overheating may reduce the HaloTag^® protein activity.

6. Centrifuge the sample at 14,000 × g for 5 minutes at 4°C to clear the lysate.
   
   

   Note: If processing nuclear fractions these may be optionally treated with RQ1 RNase-Free DNase (Cat.# M6101) to reduce DNA content in the nuclear lysate. After the standard lysis protocol, add 30µl of 10X RQ1 DNase 10X Reaction Buffer and 3µl of RQ1 RNase-Free DNase. Incubate at room temperature for 10 minutes with gentle shaking. Continue with the standard lysis protocol.

7. Transfer the clear lysate to new tube, and place the tube on ice until Phase 3.
   
   

Phase 3. Bind Protein Complexes

1. Immediately before binding, dilute the 300µl of clear lysate prepared in Phase 1 with 700µl of 1X TBS.
   
   

   Note: We recommend you determine the binding efficiency (see Technical Manual #TM342), and to do so you will need to set aside 10µl of the diluted lysate as the prebinding fraction. Store this fraction on ice.

2. Remove the Resin Equilibration/Wash Buffer supernatant from the equilibrated resin, and add the remainder of the diluted lysate.
   
   
3. Incubate with mixing on a tube rotator (or equivalent device) for 15 minutes at room temperature. Make certain that the resin does not settle to the bottom of the tube; settling will reduce binding efficiency.
   
   

   Note: In most cases 15 minutes binding time is sufficient to capture abundant protein complexes. For low abundance or larger protein complexes, this incubation time can be extended to 30-60 minutes at room temperature. Longer incubation times may increase non-specific binding. To capture membrane associated protein complexes extend the binding incubation time to 60 minutes at room temperature. For unstable or temperature sensitive protein complexes the binding can be performed at 4°C for 2 hours to overnight.

4. Centrifuge the tubes for 2 minutes at 800 × g, and discard the supernatant.
   
   

   Note: To determine the binding efficiency, set aside 10µl of the supernatant as the unbound fraction. Store this fraction on ice.

Phase 4. Washing

1. Add 1ml of Resin Equilibration/Wash Buffer to each tube, and mix thoroughly by gently inverting the tube. Centrifuge for 2 minutes at 800 × g. Discard the wash. Repeat three additional times, for a total of four washes.
   
   
2. Add 1ml of Resin Equilibration/Wash Buffer, and mix thoroughly by inverting the tube. Incubate at room temperature for 5 minutes with mixing. Centrifuge for 2 minutes at 800 × g. Discard the wash.
   
   

   Note: The stability of different protein complexes will depend on the binding affinities between the proteins in the complex, and the washing conditions may need to be optimized.

Phase 5. Protein Elution

1. For each sample, resuspend the resin with 50µl of SDS Elution Buffer. Incubate the tubes for 30 minutes with shaking at room temperature.
   
   

   Note: In some instances, it is possible to substitute the SDS Elution Buffer for an optional urea elution buffer as described in the Technical Manual #TM342. Samples eluted in urea may be directly digested with Lys-C prior to mass spectroscopy analysis.

2. Centrifuge for 2 minutes at 800 × g, and carefully transfer the eluate to a fresh tube leaving the resin at the bottom.
   
   

   Note: Resin particles in the eluted fraction could be problematic if the sample is to be analyzed directly in solution by mass spectroscopy. This elution method releases the interacting protein partners and leaves behind the HaloTag^® fusion protein, which is covalently bound to the resin. Alternatively, TEV Protease cleavage can be used to isolate the entire complex including the bait protein originally fused to the HaloTag^® protein.

In Vitro Pull-Down Assays

Glutathione-S-Transferase (GST) Pull-Down Assays

The glutathione-S-transferase (GST) pull-down assay (Kaelin et al. 1991) is an important tool to validate suspected protein:protein interactions and identify new interacting partners (Benard and Bokoch, 2002; Wang et al. 2000; Wada et al. 1998; Malloy et al. 2001). GST pull-down assays use a GST-fusion protein (bait) bound to glutathione (GST)-coupled particles to affinity purify any proteins that interact with the bait from a pool of proteins (prey) in solution. Bait and prey proteins can be obtained from multiple sources, including cell lysates, purified proteins and in vitro transcription/translation systems.

The MagneGST™ Pull-Down System (Cat.# V8870) is optimized for detection of protein:protein interactions where the bait protein is prepared from an E. coli lysate and mixed with prey protein synthesized in the TNT^® T7 Quick Coupled Transcription/Translation System (Cat.# L1170). The magnetic nature of the MagneGST™ GSH-linked particles in this system offers significant advantages over traditional resins, which require lengthy preparation and equilibration and are hard to dispense accurately in small amounts. The MagneGST™ Particles are easy to dispense in volumes less than 5μl, and equilibration is quick and easy and does not require any centrifugation steps. Another advantage of this system is that the pull-down reaction is performed in one tube. The particles are easily and efficiently separated from supernatants using a magnetic stand without centrifugation, increasing reproducibility and reducing sample loss. The flexible format of the MagneGST™ Pull-Down System allows optimization of experimental conditions, including modification of particle volume, to fit specific requirements of each unique protein:protein interaction. Additionally, the system allows easy processing of multiple samples at once.

The MagneGST™ Pull-Down System provides GST-linked magnetic particles that enable simple immobilization of bait proteins from bacterial lysates and an in vitro transcription/translation system for expressing prey proteins. The MagneGST™ Pull-Down protocol can be divided into three phases: 1) the prey protein is expressed in the TNT^® T7 Quick Coupled System; 2) bait protein present in crude E. coli lysate is immobilized on the MagneGST™ Particles; and 3) the prey protein is mixed with MagneGST™ Particles carrying the bait protein and captured through bait:prey interaction. Nonspecifically bound proteins are washed away, and the prey and bait proteins are eluted with SDS loading buffer. Prey proteins can be analyzed by SDS-PAGE and autoradiography if the prey protein was radioactively labeled during synthesis.

The transcription/translation component of the MagneGST™ Pull-Down System is the TNT^® T7 Quick Master Mix, which allows convenient, single-tube, coupled transcription/translation of genes cloned downstream from a T7 RNA polymerase promoter. The TNT^® System is compatible with circular (plasmid) or linear (plasmid or PCR product) templates. For more information on the TNT^® T7 Quick Coupled Transcription/Translation System, refer to Technical Manual #TM045. An overview of the MagneGST™ Pull-Down System is depicted in Figure 11.12. An animated presentation of the MagneGST™ pull-down process using the TNT^® T7 Quick Coupled System is available. More information and detailed protocol information is available in Technical Manual #TM249.

Example Protein Pull-Down Protocol Using the MagneGST™ Pull-Down System

Materials Required:
(see Composition of Solutions section)

• MagneGST™ Pull-Down System (Cat.# V8870) and protocol
• Magnetic Separation Stand (Cat.# Z5342, Z5343, Z5332, Z5333 or A2231)
• radiolabeled methionine (e.g., ]³⁵S]Met, 10–40μCi per TNT^® reaction) for radioactive detection of prey protein or specific antibodies for detection using Western blot analysis
• RQ1 RNase-Free DNase (Cat.# M6101)
• NANOpure^® or double-distilled water
• SDS loading buffer
• BSA (Cat.# W3841) or IGEPAL^® CA-630 (Sigma Cat.# I3021)

Express Prey Protein using a TNT^® T7 Quick Coupled Transcription/Translation Reaction

1. Remove the reagents from storage at –70°C. (Store the RQ1 DNase at –20°C after first use.) Thaw the TNT^® T7 Quick Master Mix by hand-warming or on ice. The other components can be thawed at room temperature and stored on ice.
   
   
2. Assemble the reaction components as shown in the table below using template DNA encoding your prey protein of interest. Incubate the reaction at 30°C for 60–90 minutes. During this incubation period, prepare the MagneGST™ Particles.
   
   

   Example of a TNT® T7 Quick Reaction Using Plasmid DNA.
   Components	Reaction Using [35S]Methionine	Reaction Using Unlabeled Methionine
   TNT® T7 Quick Master Mix	40μl	40μl
   Methionine, 1mM	–	1μl
   [35S]methionine (1,000Ci/mmol at 10mCi/ml)	2μl	–
   plasmid DNA template(s) (0.5μg/μl)	2μl	2μl
   Nuclease-Free Water to a final volume of	50μl	50μl

Immobilize GST-Fusion Proteins onto MagneGST™ Particles

Lyse Cells

1. Harvest cells from 1ml of bacterial culture.
   
   
2. Add 200μl of MagneGST™ Cell Lysis Reagent to each fresh or frozen cell pellet. Resuspend the cell pellet at room temperature (20–25°C) by pipetting or gentle mixing.
   
   
3. Add 2μl of RQ1 RNase-Free DNase.
   
   

   Note: Addition of DNase reduces viscosity and can increase the purity of GST-fusion proteins. Up to 5μl of RQ1 RNase-Free DNase can be added to reduce viscosity. The DNase can be omitted, if desired.

4. Incubate the cell suspension at room temperature for 20–30 minutes on a rotating platform or shaker. During this incubation, begin the particle equilibration procedure.
   
   

Equilibrate Particles

1. Thoroughly resuspend the MagneGST™ Particles by inverting the bottle several times to obtain a uniform suspension.
   
   
2. Pipet 20μl of MagneGST™ Particles into a 1.5ml tube.
   
   
3. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Magnetic capture will typically occur within a few seconds.
   
   
4. Carefully remove and discard the supernatant.
   
   
5. Remove the tube from the magnetic stand. Add 250μl of MagneGST™ Binding/Wash Buffer to the particles, and resuspend by pipetting or inverting.
   
   
6. Repeat Steps 3–5 two more times for a total of three washes.
   
   

Bind Protein

1. After the final wash, resuspend the particles in 100µl of MagneGST™ Binding/Wash Buffer.
   
   

   Note: Adding up to 1% BSA may reduce nonspecific binding and potential problems with background. IGEPAL^® CA-630 (NP40 analog) at final concentration 0.5% may have the same effect. The amount of BSA used may need to be optimized for your particular protein.

2. Add 200μl of cell lysate containing the GST-fusion protein or GST control to the MagneGST™ Particles.
   
   
3. Incubate (with constant gentle mixing) for 30 minutes at room temperature on a rotating platform.
   
   

   Note: Do not allow the MagneGST™ Particles to settle for more than a few minutes during capture of the bait protein as this will reduce binding efficiency.

Wash

1. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Carefully remove the supernatant, and save for gel analysis (optional).
   
   
2. Add 250μl of MagneGST™ Binding/Wash Buffer to the particles, and gently mix. Incubate at room temperature for 5 minutes while mixing occasionally by tapping or inverting the tube.
   
   
3. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Carefully remove the supernatant, and discard (or save if analysis of wash is desired).
   
   
4. Add 250μl of MagneGST™ Binding/Wash Buffer to the particles, and mix gently by inverting the tube. (The 5-minute incubation is not required at this wash step.)
   
   
5. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Carefully remove the supernatant, and discard (or save if analysis of wash is desired).
   
   
6. Repeat Steps 4–5 for a total of three washes.
   
   
7. After the last wash, resuspend the particles in 20μl of MagneGST™ Binding/ Wash Buffer.
   
   
8. We recommend using 5μl of the immobilized GST-fusion or GST control for the pull-down assay. Thus, 20μl of particles will provide sufficient material for more than one set of pull-down reactions. However, in some cases more than 5μl may be required for one pull-down reaction.
   
   

Capture, Wash and Analysis of Prey Protein

Capture

1. Add 20µl of the TNT^® T7 Quick coupled transcription/translation reaction from Phase 1 to each 5μl aliquot of particles carrying GST-fusion protein (or GST control).
   
   
2. Add 155μl MagneGST™ Binding/Wash Buffer and 20μl 10% BSA (or 175μl MagneGST™ Binding/Wash Buffer if BSA is omitted) to a final volume of 200μl for each pull-down reaction.
   
   

   Note: MagneGST™ Binding/Wash Buffer is a neutral PBS buffer, allowing the user to optimize buffer conditions for each specific protein:protein interaction. Some protein interactions will require the presence of various cofactors, salts and detergents.

3. Incubate for 1 hour (with gentle mixing) at room temperature on a rotating platform.
   
   

   Note: Do not allow the MagneGST™ Particles to settle for more than a few minutes during capture of the prey protein, as this will reduce binding efficiency.

4. Place the tube in a magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet.
   
   

Washing

1. Add 400μl of MagneGST™ Binding/Wash Buffer, and mix gently by inverting the tube.
   
   
2. Incubate at room temperature for 5 minutes while mixing occasionally by tapping or inverting the tube.
   
   
3. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Remove the supernatant, and save for analysis (it is especially important to keep this fraction during initial optimization).
   
   
4. Add 400μl of MagneGST™ Binding/Wash Buffer, and mix gently by inverting the tube. (The 5-minute incubation is not required at this wash step.)
   
   
5. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet.
   
   
6. Repeat Steps 4 and 5 three more times for a total of five washes.
   
   

Elution

1. Add 20μl of 1X SDS loading buffer.
   
   
2. Incubate for 5 minutes at room temperature with mixing.
   
   
3. Place the tube in the magnetic stand, and allow the MagneGST™ Particles to be captured by the magnet. Remove the eluate for analysis.
   
   

Analysis

Prepare samples for SDS-PAGE analysis. For radioactively labeled prey proteins, we recommend loading 1–2% of each sample volume.

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Analysis of DNA:Protein Interactions

Regulation of chromatin structure and gene expression is essential for normal development and cellular growth. Transcriptional events are tightly controlled both spatially and temporally by specific protein:DNA interactions. Currently there is a rapidly growing trend toward genome-wide identification of protein-binding sites on chromatin to characterize regulatory protein:DNA interactions that govern the transcriptome. Common methods to examine protein:DNA interactions include the electrophoretic mobility shift assay, also known as the gel shift assay, and chromatin immunoprecipitation (Solomon et al. 1985; Solomon et al. 1988) coupled with DNA microarray or ultrahigh-throughput sequencing analysis.

Gel Shift Assays

Electrophoretic mobility shift assays (EMSA) or gel shift assays can be used to analyze protein:DNA complexes expressed in vitro. The proteins are incubated with an oligonucleotide containing a target consensus sequence site, and DNA binding is detected by gel shift. An animated presentation of protein:DNA interaction detection using the TNT^® Systems and Gel Shift Assay is available. The gel shift assay provides a simple and rapid method to detect DNA-binding proteins (Ausubel et al. 1989). This method is used widely in the study of sequence-specific DNA-binding proteins such as transcription factors. The assay is based on the observation that complexes of protein and DNA migrate through a nondenaturing polyacrylamide gel more slowly than free DNA fragments or double-stranded oligonucleotides. The gel shift assay is performed by incubating a purified protein, or a complex mixture of proteins (such as nuclear or cell extract preparations), with a ³²P end-labeled DNA fragment containing the putative protein-binding site. The reaction products are then analyzed on a nondenaturing polyacrylamide gel. The specificity of the DNA-binding protein for the putative binding site is established by competition experiments using DNA fragments or oligonucleotides containing a binding site for the protein of interest or other unrelated DNA sequences.

Promega gel shift assay systems contain target oligonucleotides, a control extract containing DNA-binding proteins, binding buffer and reagents for phosphorylating oligonucleotides. The Gel Shift Assay Core System (Cat.# E3050) includes sufficient HeLa nuclear extract to perform 20 control reactions, Gel Shift Binding 5X Buffer, an SP1 Consensus Oligo and an AP2 Consensus Oligo. The complete Gel Shift Assay System (Cat.# E3300) contains five additional double-stranded oligonucleotides that represent consensus binding sites for AP1, NF-κB, OCT1, CREB and TFIID. These oligonucleotides can be end-labeled and used as protein-specific probes or as specific or nonspecific competitor DNA in competition assays. A detailed protocol is available in Technical Bulletin #TB110.

Chromatin Immunoprecipitation

Chromatin immunoprecipitation (ChIP) is an experimental method used to determine whether DNA-binding proteins, such as transcription factors, associate with a specific genomic region in living cells or tissues. Cells are treated with formaldehyde to form covalent crosslinks between interacting proteins and DNA. Following crosslinking, cells are lysed, and the crude cell extracts are sonicated to shear the DNA. The DNA:protein complex is immunoprecipitated using an antibody that recognizes the protein of interest. The isolated complexes are washed, then eluted. The DNA:protein crosslinks are reversed by heating and the proteins removed by proteinase K treatment. The remaining DNA is purified and analyzed by various ways, including PCR, microarray analysis or direct sequencing.

Antibody-Based ChIP

The standard ChIP assay requires 3–4 days for completion (Figure 11.13). The procedure requires antibodies highly specific to the protein of interest to immunoprecipitate the DNA:protein complex. The success of the procedure relies on the ability of the antibody to bind to the target protein after crosslinking, cell lysis and sonication, all of which can negatively affect epitope recognition by the antibody.

HaloCHIP™ System—an Antibody-Free Approach

To address the difficulties that arise when performing ChIP, a novel method that does not require the use of antibodies, the HaloCHIP™ System, has been devised for the covalent capture of protein:DNA complexes. DNA-binding proteins of interest are expressed in cells as HaloTag^® fusion proteins, crosslinked to DNA, then captured on the HaloLink™ Resin, which forms a highly specific, covalent interaction with HaloTag^® proteins. Due to the covalent linkage between the resin and crosslinked protein:DNA complexes, the resin can be stringently washed to remove nonspecifically bound DNA and protein more efficiently than co-immunoprecipitation. The crosslinks are reversed to release purified DNA fragments from the resin. By improving specificity and reducing background during the isolation of protein:DNA complexes, the HaloCHIP™ approach effectively increases the signal-to-noise ratio to permit detection of small changes in protein binding within a genome. The HaloCHIP™ System (Cat.# G9410) is currently available. An animation of this procedure is available.

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Immunoassays: ELISA and Western Blot Analysis

ELISA

Enzyme-linked immunosorbent assay (ELISA) combine the specificity of antibodies with the sensitivity of reporter enzyme assays, using antibodies or antigens conjugated to an easily-assayed enzyme. The purpose of the ELISA is to determine if a particular protein is present in a sample and to quantitate how much. There are two main variations of this method: the ELISA can determine how much antibody is in a sample, or it can quantitate the amount of a specific antigen in the sample.

In general, the first step in an ELISA is to adsorb the analyte (material to be assayed or measured) to a solid support (e.g. 96-well plate). This is followed by sequentialy incubating with primary and secondary antibodies.The secondary antibody is linked to an enzyme, which will react with its substrate to produce a detectable signal, such as a color change in the substrate, or a florescent or chemiluminescent signal. Alternatively, antibodies could be linked to fluorescent or radioactive labels and detected using the appropriate instrumentation. ELISAs are one of the most widely used molecular biology techniques. For example, ELISAs are used in antibody production to identify which clones or animals are are producing a high level of antigen-specific antibody. In addition, they can be used to quantitate the amount of a specific antigen in a sample since the intensity of signal will be directly proportional to the amount of antigen captured and bound by the secondary antibodies.

Indirect ELISA

In an indirect ELISA, the antigen is immobilized on a solid surface such as a 96-well plate, the surface is then blocked to prevent non-specific binding of downstream reagents. The blocked plate is then incubated with primary antibody that will specifically bind to the antigen. The appropriately labeled secondary antibody is then added to bind to and detect the primary antibody; the enzyme label on the secondary antibody reacts with the detection substrate and generates a visible signal. in this method it is critical that the secondary antibody be specific for the primary antibody only. This can be achieved by using secondary antibodies that have been raised against the host species of the primary antibody (e.g., goat anti-mouse IgG secondary antibodies would bind all mouse antibodies). The detection substrate reacts with the enzyme label on the antibody (e.g., horseradish peroxidase) and generates either a chromogenic (color change), fluorescent or luminescent signal. The presence of the signal indicates secondary antibody has bound to primary antibody; the higher the concentration of the primary antibody, the greater the signal. One core disadvantage of the indirect ELISA approach is that the method of antigen immobilization is nonspecific. If crude serum is used as the source of antigen analyte, then all the proteins in the serum will adsorb to the plate, and the antigen must compete with other serum proteins when binding to the surface of the plate.

Capture ELISA (Sandwich ELISA)

In a sandwich ELISA a capture antibody specific to the antigen analyte is first coated on to the microplate surface. The sample containing the antigen is added and allowed to bind to the capture antibody on the surface. This is followed by adding a primary antibody that will bind to the antigen. The monicker, "sandwich ELISA", is based on that the analyte antigen is sandwiched between two primary antibodies. This is followed by addition of appropriately labeled secondary antibodies and the corresponding detection substrate as discussed above. It is important to use the secondary antibody against the same species as the primary antibody and not the capture antibody. The capture antibody and the primary antibodies need to come from two different species (e.g. mouse and rabbit). The main advantage of the sandwich ELISA approach is that it greatly enhances the specificity and sensitivity of the assay. The main disadvantage is that it requires a matched pair of antibodies that will bind to two different sites (epitopes) on the antigen to form a "sandwich".

Direct ELISA

The direct ELISA approach uses a directly labeled primary antibody that reacts with the antigen. Direct detection is performed in lieu of using a labeled secondary antibody. The main advantages of this approach are that it is simpler and more quantitative compared to using a secondary antibody. The main disadvantages are that direct labeling of the primary antibodies (or antigens) is time-consuming and may negatively affect their reactivity.

Competitive ELISA

The steps for a competitive ELISA are slightly different than for the previously mentioned methods. For a competitive ELISA, unlabeled (primary) antibody is incubated with the sample containing the antigen and then the mixture is added to an antigen-coated well. Only the free antibody that did not previously bind to the antigen in the sample will bind to the antigen-coated well. Washing removes any unbound material, leaving bound only the antibody that was not competed away by the antigen in the sample. The term competitive is based on the fact that the more antigen is present in the sample, the less antibody will be able to bind to the antigen in the well. As described before, the secondary antibody, conjugated to an enzyme, is added and followed by the substrate to generate a signal. In this type of ELISA, the higher the antigen concentration in the sample, the weaker the eventual signal. The competitive ELISA method offers the advantage of using crude or impure samples and still selectively binding any antigen that may be present. The main dissadvantage of this approach is that it is more difficult to setup and optimize.

Common Enzyme labels

Enzyme-conjugated secondary antibodies offer the most flexibility in detection and documentation methods for ELISA because of the variety of substrates available for chromogenic, fluorescent and chemiluminescent imaging. The most commonly used enzyme labels used for ELISA are horseradish peroxidase (HRP) and alkaline phosphatase (AP). There are a large variety of substrates available for performing ELISA with an HRP or AP conjugates. The substrate choice depends upon the desired assay sensitivity and the instrumentation available for signal-detection. In addition, fluorescent tags, radioactive tags and other alternatives to enzyme-based detection methods can be used for plate-based ELISAs.

Although chromogenic ELISA substrates are not as sensitive as fluorescent or chemiluminescent substrates, they are economical and still enable kinetic studies. Furthermore, chromogenic ELISA substrates are detected with standard absorbance plate readers that are common in many laboratories. In contrast, fluorescent ELISA substrates require a fluorescent plate reader or scanner with appropriate filters for the excitation and emission wavelengths of the detected fluorophores. Chemiluminescent substrates can be detected by various means including digital camera systems although they are best used with a luminometer. However, the signal intensity can vary more with chemiluminescent substrates compared to other substrates. This may be problematic for high-throughput assays requiring accurate measurement across a large number of plates.

Western Blot Analysis

The western blot (sometimes called the protein immunoblot) is a widely used analytical technique performed to detect specific proteins in a sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins either dependent on the length of the polypeptide chain (denaturing conditions) or on the structure and charge of proteins (native/ non-denaturing conditions). This technique provides information about protein molecular weight and presence of different protein isoforms (e.g., glycosylation) of the proteins under study. Following electrophoresis, the proteins are transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target proteins.

SDS-PAGE

By far the most common type of gel electrophoresis is Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). This method uses polyacrylamide gels and buffers containing sodium dodecyl sulfate (SDS) to analyze and isolate small amounts of protein. During SDS-PAGE, proteins are denatured and coated with detergent by heating in the presence of SDS and a reducing agent. The SDS coating gives the protein a high net negative charge that is proportional to the length of the polypeptide chain and denatures the protein by interfering with the non-covalent interactions (e.g., ionic, hydrophobic) that stabilize protein structure. The sample is loaded on a polyacrylamide gel, and high voltage is applied, causing the proteins to migrate as unfolded peptide chains toward the positive electrode (anode). SDS-PAGE is commonly performed in reducing conditions (e.g., presence of DTT or beta-mercaptoethanol) in order to reduce the disulfide bonds found within some proteins and to facilitate protein denaturation.

Since the proteins have a net negative charge that is proportional to their size, proteins are separated solely on the basis of their molecular mass—a result of the sieving effect of the gel matrix. The molecular mass of a protein can be estimated by comparing the gel mobility of a band with those of protein standards. Sharp protein bands are achieved by using a discontinuous gel system, having stacking and separating gel layers that differ in either salt concentration or pH or both (Hanes, 1981).

Materials Required:
(see Composition of Solutions section)

• SDS-polyacrylamide gel running 1X buffer
• loading 2X buffer
• trichloroacetic acid (TCA) (optional)
• acetone, ice-cold (optional)
• pre-cast acrylamide gels (gradient of defined percentage) –or–
• acrylamide solution, 40%
• upper gel 4X buffer
• lower gel 4X buffer
• ammonium persulfate, 10%
• TEMED

This gel system uses the method described by Laemmli (Laemmli, 1970). Formulations for preparing resolving and stacking minigels are provided in Tables 11.4 and 11.5. The amounts of reagents indicated in Tables 11.4 and 11.5 are sufficient to prepare two 7 × 10cm gels, 0.75–1.00mm thick. Add ammonium persulfate and TEMED just prior to pouring the gel, as these reagents promote and catalyze polymerization of acrylamide. Pour the resolving gel mix into assembled gel plates, leaving sufficient space at the top for the stacking gel to be added later. Gently overlay the gel mix with 0.1% SDS, and allow the gel to polymerize for at least 15–30 minutes. After polymerization, remove the SDS overlay, and rinse the surface of the resolving gel with water to remove any unpolymerized acrylamide. Rinse one more time with a small volume of stacking gel buffer. Fill the remaining space with the stacking gel solution, and insert the comb immediately. After the stacking gel has polymerized, remove the comb, and rinse the wells with water to remove unpolymerized acrylamide. At least 1cm of stacking gel should be present between the bottom of the loading wells and the resolving gel.

Table 11.4. Formulation for Stacking Gel.
Component	Volume
upper gel 4X buffer	2.5ml
water	6.6ml
acrylamide solution, 40%	0.8ml
APS, 10% 1	100μl
TEMED 2	10μl

¹ammonium persulfate (always prepare fresh)

²N,N,N′,N′-tetramethylethylenediamine

Table 11.5. Formulation for Resolving Gel.
	Volume for Different Percentages of Acrylamide
Component	8%	10%	12%	15%	20%
lower gel 4X buffer	2.5ml	2.5ml	2.5ml	2.5ml	2.5ml
water	5.4ml	4.9ml	4.4ml	3.65ml	2.4ml
acrylamide solution, 40%	2.0ml	2.5ml	3.0ml	3.75ml	5.0ml
APS, 10%1	50.0μl	50.0μl	50.0μl	50.0μl	50.0μl
TEMED 2	5.0μl	5.0μl	5.0μl	5.0μl	5.0μl

¹ammonium persulfate (always prepare fresh)

²N,N,N′,N′-tetramethylethylenediamine

Prepare Samples

1. Add an equal volume of loading 2X buffer to the sample.
   
   
2. Incubate the sample at 95°C for 2–5 minutes, mix by vortexing and load onto the gel.
   
   

Optional Protein Precipitation

1. If the sample is very dilute or contains salts that may interfere with gel analysis, the protein can be precipitated and resuspended prior to SDS-PAGE analysis.
   
   

   Note: The precipitated protein is denatured and needs to be resuspended in a detergent buffer, a chaotropic salt or an organic solvent.

2. Add 150µl of dilute protein to a microcentrifuge tube, add 600µl of methanol, and vortex. .
   
   
3. Add 150µl of chloroform, and vortex.
   
   
4. Add 450ul of Nuclease-Free Water, and vortex.
   
   
5. Centrifuge for 2 minutes at 14,000 × g. An interface will form between the aqueous (top) and organic phases. The protein is in the interface layer.
   
   
6. Carefully remove the top aqueous layer. The salts, detergents, sugars are in the aqueous layer. Note: There is no need to remove the entire aqueous layer. Remove as much as possible with out disturbing the interface layer.
   
   
7. Add 600µl of methanol and vortex.
   
   
8. Centrifuge the sample at 14,000 × g for 5–10 minutes. The protein will form a tight pellet.
   
   
9. Remove the supernatant and air dry the pellet.
   
   
10. Resuspend the protein in a suitable volume (15–20µl) of loading 1X buffer (prepared by adding an equal volume of water to loading 2X buffer).
    
    
11. Incubate the sample at 95°C for 2–5 minutes, mix by vortexing and load onto the gel.
    
    

Blotting

Following electrophoresis and before detection, the proteins must be transferred from the gel onto a nitrocellulose or polyvinylidene difluoride (PVDF) membrane. The membrane is placed on top of the gel, and a stack of filter papers placed on top of that. When the gel/filter/paper stack is placed in a buffer solution, capillary action pulls the proteins out of the gel and into the membrane. Alternatively, the proteins can be transferred by electroblotting, which uses an electric current to pull proteins from the gel into the membrane. With either blotting method, the proteins maintain the relative position they had within the gel but now they are embedded in a thin surface layer that is more suitable for detection. Both membrane material have nonspecific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic and ionic interactions between the membrane and protein.

Blocking

The nonspecific protein binding properties of membranes require that the membrane is blocked prior to addition of any antibodies. Blocking this nonspecific binding is achieved by placing the membrane in a dilute solution of protein such as 3-5% Bovine serum albumin (BSA; Cat.# W3841) or non-fat dry milk in Tris-Buffered Saline (TBS), with a small percentage of detergent such as Tween 20 or Triton X-100. The protein in this dilute solution attaches to the membrane where ever the target proteins have not attached. As a result, when the antibody is added, there is no where for it to attach other than the binding sites of the target protein. This reduces background in the final western blot, leading to clearer results, and fewer false positive results.

Detection

Following blotting, the membrane is probed for the protein of interest. Typically, this is done using a primary antibody specific to the protein of interest and a secondary antibody that is directed towards a species-specific portion of the primary antibody. This secondary antibody is modified so that it is linked to a reporter enzyme that will produce color when exposed to an appropriate substrate. Although this two-step process is the approach most commonly used, there are now one-step detection methods available for certain applications.

While there are many different tags that can be conjugated to a secondary or primary antibody, the detection method used will limit the choice of what can be used in a Western blotting assay. Radioisotopes were used extensively in the past, but they are expensive, have a short shelf-life, offer no improvement in signal:noise ratio and require special handling and disposal. Alternative labels are enzymes and fluorophores.

Enzymatic labels are most commonly used for Western blotting ans can be extremely sensitive when optimized with an appropriate substrate. Alkaline phosphatase (AP) and horseradish peroxidase (HRP) are the two most commonly used enzymes. An array of chromogenic, fluorogenic and chemiluminescent substrates are available for use with either enzyme. AP catalyzes colorimetric reactions using substrates such as the Western Blue^® Substrate (standard BCIP/NBT, Cat.# S3841). It can also drive the chemiluminescent detection reactions involving substrates such as 3-(2´-spiroadamantane)-4-methyl-4-(3´´ phosphoryloxyphenyl-1, 2-dioxetane (AMPPD^®). AP conjugated antibodies offer the advantage that their reaction rates remain linear so that sensitivity can be increased by letting the reaction to proceed for a longer time. Unfortunately, this increased reaction time often leads to high background signal resulting in low signal:noise ratios.

In contrast, Horseradish peroxidase (HRP) conjugated antibodies offer higher specificity compared to AP conjugates due the smaller size of HRP enzyme and compatibility with conjugation reactions. In addition, the high activity rate, good stability, low cost and wide availability of substrates make HRP the enzyme of choice for most applications.

Although, enzyme conjugated antibodies offer the most flexibility in detection and documentation methods for Western blotting because of the variety of substrates available, the simplest detection system is chromogenic substrates such as Western Blue^®Substrate (Cat.# S3841). Chromogenic substrates allow direct visualization of blot development but lack the sensitivity of enzyme conjugated methods. Unfortunately, chromogenic substrates tend to fade as the blot dries, so it is important to make a permanent image of the blot.

Chemiluminescent blotting substrates such as the ECL Western blot substrate (Cat. W1001) are different from other substrates because the signal is a product of the enzyme-substrate reaction that persists only as long as the reaction is occurring. As a result, the signal is lost once the substrate is used up or the enzyme looses activity. However, in well-optimized assays with proper antibody dilutions and sufficient substrate, the reaction can produce stable light output for several hours.

Using fluorophore-conjugated antibodies in a immunoassays requires fewer steps because there is no substrate development step. While the protocol is shorter, special equipment is needed to detect and document the fluorescent signal. Recent advances in digital imaging and the development of new fluorophores has improved the sensitivity and increase the popularity of using fluorescent probes for Western blotting. Finally, this method has the added advantage of multiplex compatibility (using more than one fluorophore in the same experiment).

Western Blot Analysis of Proteins from TNT^® Cell-Free Expression Systems

More information and detailed protocol information is available in Technical Manual #TM045.

Protocol

Materials Required:
(see Composition of Solutions section)

• 1M NaOH/2% H₂O₂
• 25% TCA/2% casamino acids (Difco brand, Vitamin Assay Grade)
• 5% TCA
• Whatman GF/A glass fiber filter (Whatman)
• acetone
• Whatman 3MM filter paper
• 30% acrylamide solution
• separating gel 4X buffer
• stacking gel 4X buffer
• SDS sample buffer
• SDS polyacrylamide gels
• 50mM DTT
• Blot-Qualified BSA (Cat.# W3841)
• PVDF membrane
• iBlot
• SP-antibod
• TBST buffer

1. Add 1µl of the standard, unlabeled translation reaction to 19µl of 1X SDS loading dye with 50mM DTT.
   
   

   Note: Include a no template control on the gel to identify background bands.

2. Incubate at 95°C for 5 minutes. Centrifuge briefly to collect the contents in the bottom of the tube.
   
   
3. Load 20µl onto a 4–20% gradient Tris-glycine SDS polyacrylamide gel.
   
   
4. Following electrophoresis, remove the gel and place it in water.
   
   
5. Transfer the proteins to a PVDF membrane using a Western blotting system (e.g., iBlot^® System; Invitrogen).
   
   
6. Block the membrane using 15ml of 5% Blot-Qualified BSA in TBST (1X TBS + 0.1% Tween^® 20). Incubate for 1 hour with gentle shaking.
   
   
7. Dilute your primary antibody in 1X TBST.
   
   

   Note: We recommend that you titrate your primary antibody dilutions to determine what dilution produces the best results for your protein.

8. Following incubation, remove the blocking solution from the membrane, and add 15ml of diluted primary antibody.
   
   
9. Incubate the membrane with the primary antibody at room temperature for 1 hour with gentle shaking.
   
   
10. Remove the primary antibody solution, and wash the membrane with 15ml of 1X TBST for 5 minutes with gentle shaking.
    
    
11. Repeat the wash 5 more times for a total of six washes.
    
    
12. Dilute your secondary antibody 1:2,500 in 1X TBST.
    
    
13. Following that last wash, remove buffer from the membrane and add 15ml of diluted secondary antibody.
    
    
14. Incubate the membrane with the secondary antibody for 1 hour with gentle shaking.
    
    
15. Following the incubation, remove the secondary antibody solution, and wash the membrane with 15ml of 1X TBST for five minutes. Repeat for a total of six washes.
    
    
16. Proceed to the detection method appropriate for your secondary antibody.
    
    

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Mass Spectrometry Analysis

Mass spectrometry is a powerful tool for protein analysis and the major technique used to study proteins in the field of proteomics (Mann et al. 2001). Mass spectrometry can be used to characterize recombinant proteins, identify unknown proteins and detect and characterize posttranslational modifications.

One method of protein identification uses enzymatic digestion followed by mass spectrometry analysis. In this procedure, complex protein mixtures such as cell extracts are resolved by gel electrophoresis, and the band or spot of interest is excised from the gel and digested with trypsin. Trypsin is a serine protease that specifically cleaves at the carboxylic side of lysine and arginine residues. The distribution of Lys and Arg residues in proteins is such that trypsin digestion yields peptides of molecular weights that can be analyzed by mass spectrometry. The pattern of peptides obtained is used to identify the protein. Database searches can then be performed, using the masses of the identified peptides to identify the protein(s) resolved on the gel (Mann et al. 2001).

The stringent specificity of trypsin is essential for protein identification. Native trypsin is subject to autolysis, generating pseudotrypsin, which exhibits a broadened specificity, including a chymotrypsin-like activity (Keil-Dlouha et al. 1971). Such autolysis products would result in additional peptide fragments that could interfere with database analysis of the mass of fragments detected by mass spectrometry.

Trypsin Gold, Mass Spectrometry Grade (Cat.# V5280), provides maximum specificity. Lysine residues in the porcine trypsin are modified by reductive methylation, yielding a highly active and stable molecule that is extremely resistant to autolytic digestion (Rice et al. 1977). The specificity of the purified trypsin is further improved by TPCK treatment, which inactivates chymotrypsin. The treated trypsin is then purified by affinity chromatography and lyophilized to yield Trypsin Gold, Mass Spectrometry Grade. More information and a detailed protocol are available in Technical Bulletin TB309.

In-Gel Trypsin Digestion of Proteins

Numerous protocols for in-gel protein digestion have been described (Flannery et al. 1989; Shevchenko et al. 1996; Rosenfeld et al. 1992). The following procedure has been used successfully by Promega scientists.

Materials Required:
(see Composition of Solutions section)

• Trypsin Gold, Mass Spectrometry Grade (Cat.# V5280) and protocol
• SimplyBlue™ SafeStain (Invitrogen Cat.# LC6060)
• trifluoroacetic acid (TFA)
• acetonitrile (ACN)
• 200mM NH₄HCO₃ buffer (pH 7.8)
• NANOpure^® water
• ZipTip^® scx pipette tips (Millipore Cat.# ZTSCXS096)
• α-cyano-4-hydroxycinnamic acid (CHCA)
• MALDI target
• ZipTip^® C18 pipette tips (Millipore Cat.# ZTC18S096)

1. Separate protein samples by electrophoresis on an SDS-Tris-Glycine gel.
   
   

   Note: Other gel systems and staining reagents can be used for in-gel digestions but should be tested to ensure compatibility with the protein of interest and detection system being used.

2. Rinse the gel three times, for 5 minutes each rinse, in NANOpure^® water. Stain for 1 hour in SimplyBlue™ SafeStain (Invitrogen Corporation) at room temperature with gentle agitation. When staining is complete, discard the staining solution.
   
   
3. Destain the gel for 1 hour in NANOpure^® water at room temperature with gentle agitation. When destaining is complete, discard the solution.
   
   
4. Using a clean razor blade, cut the protein bands of interest from the gel, eliminating as much polyacrylamide as possible. Place the gel slices into a 0.5ml microcentrifuge tube that has been prewashed twice with 50% acetonitrile (ACN)/0.1% trifluoroacetic acid (TFA).
   
   
5. Destain the gel slices twice with 0.2ml of 100mM NH₄HCO₃/50% ACN for 45 minutes each treatment, at 37°C to remove the SimplyBlue™ SafeStain.
   
   
6. Dehydrate the gel slices for 5 minutes at room temperature in 100μl of 100% ACN. At this point the gel slices will be much smaller than their original size and will be whitish or opaque in appearance.
   
   
7. Dry the gel slices in a Speed Vac^® concentrator for 10–15 minutes at room temperature to remove the ACN.
   
   
8. Resuspend the Trypsin Gold at 1μg/μl in 50mM acetic acid, then dilute in 40mM NH₄HCO₃/10% ACN to 20μg/ml. Preincubate the gel slices in a minimal volume (10–20μl) of the trypsin solution at room temperature (do not exceed 30°C) for 1 hour. The slices will rehydrate during this time. If the gel slices appear white or opaque after one hour, add an additional 10–20μl of trypsin and incubate for another hour at room temperature.
   
   
9. Add enough digestion buffer (40mM NH₄HCO₃/10% ACN) to completely cover the gel slices. Cap the tubes tightly to avoid evaporation. Incubate overnight at 37°C.
   
   
10. Incubate the gel slice digests with 150μl of NANOpure^® water for 10 minutes, with frequent vortex mixing. Remove and save the liquid in a new microcentrifuge tube.
    
    
11. Extract the gel slice digests twice, with 50μl of 50% ACN/5% TFA (with mixing) for 60 minutes each time, at room temperature.
    
    
12. Pool all extracts (from Steps 10 and 11), and dry in a Speed Vac^® concentrator at room temperature for 2–4 hours (do not exceed 30°C).
    
    
13. Purify and concentrate the extracted peptides using ZipTip^® pipette tips (Millipore Corporation) following the manufacturer’s directions.
    
    
14. The peptides eluted from the ZipTip^® tips are now ready for mass spectrometric analysis.
    
    

In-Solution Trypsin Digestion of Proteins

In general, proteins require denaturation and disulfide bond cleavage for enzymatic digestion to reach completion (Wilkinson, 1986). If partial digestion of a native protein is desired, begin this protocol at Step 3.

1. Dissolve the target protein in 6M guanidine HCl (or 6–8M urea or 0.1% SDS), 50mM Tris-HCl (pH 8), 2–5mM DTT (or β-mercaptoethanol).
   
   
2. Heat at 95°C for 15–20 minutes or at 60°C for 45–60 minutes. Allow the reaction to cool.
   
   
3. For denatured proteins, add 50mM NH₄HCO₃ (pH 7.8) or 50mM Tris-HCl, 1mM CaCl₂ (pH 7.6), until the guanidine HCl or urea concentration is less than 1M. If SDS is used, dilution is not necessary. For digestion of native proteins, dissolve in buffer with a pH between 7 and 9.
   
   
4. Add Trypsin Gold to a final protease:protein ratio of 1:100 to 1:20 (w/w). Incubate at 37°C for at least 1 hour. Remove an aliquot, and chill the remainder of the reaction on ice or freeze at –20°C.
   
   
5. Terminate the protease activity in the aliquot from Step 4 by adding an inhibitor. Alternatively, precipitate the aliquot by adding TCA to 10% final concentration. The reaction can also be terminated by freezing at –20°C. Trypsin can also be inactivated by lowering the pH of the reaction below pH 4. Trypsin will regain activity as the pH is raised above pH 4 (Wilkinson, 1986).
   
   

   Note: The following are general trypsin inhibitors: Antipain (50μg/ml), antithrombin (1unit/ml), APMSF (0.01–0.04mg/ml), aprotinin (0.06–2μg/ml), leupeptin (0.5μg/ml), PMSF (17–170µg/ml), TLCK (37–50μg/ml), trypsin inhibitors (10–100μg/ml).

6. Determine the extent of digestion by subjecting the aliquot in Steps 4 and 5 to reverse phase HPLC or SDS-PAGE.
   
   
7. If no inhibitors were added to the remainder of the reaction and further proteolysis is required, incubate at 37°C until the desired digestion is obtained (Sheer, 1994). Reducing the temperature will decrease the digestion rate. Incubations of up to 24 hours may be required, depending on the nature of the protein. With long incubations, take precautions to avoid bacterial contamination.
   
   

Affinity Tag In Vitro Pull-Down Assay with Trypsin Digestion and Protein Analysis

Markillie and associates describe a simple exogenous protein complex purification and identification method that can be easily automated (Markillie et al. 2005). The method uses MagneHis™ Ni Particles (Cat.# V8560, V8565) to pull down target proteins, followed by denaturing elution, trypsin digestion and mass spectrometry analysis (Figure 11.15).

Alternative Proteases

Use of alternative enzymes for protein digestion increases the confidence in mass spectrometry data by confirming the protein sequence and aids in the mapping of post-translational modifications (PTMs). Endoproteinases Asp-N and Glu-C have been used for protein characterization for over 30 years and have gained importance recently due to advancements in mass spectrometry techniques. Asp-N preferentially cleaves proteins at the N-terminus of aspartic and cysteic acid (Drapeau 1980; Ingrosso et al., 1989; Geu et al., 1990). Glu-C cleaves at the C-terminus of glutamic and aspartic residues (Drapeau, 1972; Drapeau, 1978; Drapeau, 1977). Due to their specific cleavage sites, these proteinases create unique peptide fragments that are well-suited for mass spectrometry analysis. Comparing protein sequences or mapping data from Asp-N or Glu-C digests to that of other proteinases promotes higher confidence in data (Fischer and Poetsch, 2006; Wu and MacCoss, 2002; Swaney et al., 2010; Choudhary et al., 2003; Elenitoba-Johnson et al., 2006; Biringer et al., 2006).

Protein digestion is required for either the bottom-up or middle-down approach to protein analysis. In the bottom-up approach, the optimal peptide size range for analysis is 600–5,000Da. Recent advancements in mass spectrometry have expanded the peptide size range to 600–20,000Da. These advancements have promoted the middle-down approach to protein analysis (Young et al., 2010; Mann and Kelleher, 2008; Meyer et al. , 2010; Cannon et al., 2010; Swaney et al., 2010). However, some peptides still may fall above or below the desired peptide size range. This results in decreased protein coverage and incomplete data collection.

To increase protein coverage, additional proteinases have been used. Using alternative proteinases individually or in combination with other proteinases creates a unique peptide map that may include sequences not seen in standard trypsin digestions. Overlaying peptide mass data obtained from digestion with alternative proteinases increases protein coverage and overall confidence in protein identification. When alternative enzymes are used, larger peptides are cleaved into smaller fragments, which are more manageable for the instrumentation. In addition, protein sections cleaved into peptides too small for analysis by one enzyme might be cleaved into larger peptides by a different enzyme. Alternative proteinases also help to overcome incomplete digestion caused by PTMs, which prevent the proteinases from accessing that particular site. By using alternative enzymes, the protein might be cleaved at sites further away from the PTM. The examples above show that alternative proteinases are beneficial to protein analysis (Young et al., 2010; Mann and Kelleher, 2008; Meyer et al. , 2010; Cannon et al., 2010; Swaney et al., 2010).

Asp-N and Glu-C Proteases

Asp-N (Cat.# V1621) is a metalloprotease purified from Pseudomonas fragi. Asp-N cleaves at the N-terminus of aspartic and cysteic acid residues with high specificity (Drapeau 1980; Ingrosso et al., 1989; Geu et al., 1990). Cysteic acid residues are an oxidized form of the cysteine residues. Additional cleavage has been reported at glutamic residues; however, the rate of cleavage at aspartic acid is 2,000-fold higher than at the glutamyl residues (Ingrosso et al., 1989; Geu et al., 1990). Glu-C (Cat. # V1651) is a serine proteinase purified from Staphylococcus aureus V8. It cleaves specifically at the C-terminus of glutamic and aspartic acids (Drapeau, 1972; Drapeau, 1978; Drapeau, 1977). Buffer composition affects the specificity of Glu-C. In phosphate buffers, both glutamic and aspartic residues are cleaved; however, in ammonium bicarbonate and ammonium acetate buffers (pH 4.0), only the glutamic residues are cleaved (Drapeau, 1977)

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Composition of Solutions

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References

1. Armstrong, R.N. (1997) Structure, catalytic mechanism, and evolution of the glutathione transferases. Chem. Res. Tox. 10, 2–18.
2. Ausubel, F.M. et al. (1989) In: Current Protocols in Molecular Biology Volume 2,John Wiley and Sons, New York .
3. Benard, V. and Bokoch, G.M. (2002) Assay of Cdc42, Rac, and Rho GTPase activation by affinity methods. Methods Enzymol. 345, 349–59.
4. Betz, N. (2004) Efficient purification of his-tagged proteins from insect and mammalian cells. Promega Notes 87, 29–32.
5. Biringer, R.G. et al. (2006) Enhanced sequence coverage of proteins in human cerebrospinal fluid using multiple enzymatic digestion and linear ion trap LC-MS/MS. Brief Funct. Genomic. Proteomic. 5, 144–53.
6. Cannon, J. et al. (2010) High-throughput middle-down analysis using an orbitrap. J. Prot. Res. 9, 3886–90.
7. Charron, F. et al. (1999) Cooperative interaction between GATA-4 and GATA-6 regulates myocardial gene expression. Mol. Cell Biol. 19, 4355–65.
8. Chien, C. et al. (1991) The two-hybrid system: A method to identify and clone genes for proteins that interact with a protein of interest. Proc. Natl. Acad. Sci. USA 88, 9578–82.
9. Choudhary, G. et al. (2003) Multiple enzymatic digestion for enhanced sequence coverage of proteins in complex proteomic mixtures using capillary LC with ion trap MS/MS. J. Proteome Res. 2, 59–67.
10. Cronan, J.E. (1990) Biotination of proteins in vivo. A post-translational modification to label, purify, and study proteins. J. Biol. Chem. 265, 10327–33.
11. Dang, C.V. et al. (1991) Intracellular leucine zipper interactions suggest c-Myc hetero-oligomerization. Mol. Cell. Biol. 11, 954–62.
12. Drapeau, G.R. (1980) Substrate specificity of a proteolytic enzyme isolated from a mutant of Pseudomonase fragi. J. Biol. Chem. 255, 839–40.
13. Drapeau, G.R. (1972) Purification and properties of an extracellular protease of Staphylococcus aureus. J. Biol. Chem. 247, 6720–6.
14. Drapeau, G.R. (1978) The primary structure of staphylococcal protease. Can. J. Biochem. 56, 534–44.
15. Drapeau, G.R. (1977) Cleavage at glutamic acid with staphylococcal protease. Methods Enzymol. 47, 189–91.
16. Elenitoba-Johnson, K.S. et al. (2006) Proteomic identification of oncogenic chromosomal translocation partners encoding chimeric anaplastic lymphoma kinase fusion proteins. Proc. Natl. Acad. Sci. USA 103, 7402–7.
17. Fearon, E.R. et al. (1992) Karyoplasmic interaction selection strategy: A general strategy to detect protein-protein interactions in mammalian cells. Proc. Natl. Acad. Sci. USA 89, 7958–62.
18. Fields, S. and Song, O. (1989) A novel genetic system to detect protein-protein interactions. Nature 340, 245–6.
19. Finkel, T. et al. (1993) Detection and modulation in vivo of helix-loop-helix protein-protein interactions. J. Biol. Chem. 268, 5–8.
20. Fischer, F. and Poetsch, A. (2006) Protein cleavage strategies for an improved analysis of the membrane proteome. Proteome Sci. 4, 2–14.
21. Flannery, A.V. et al. (1989) "Proteolysis of proteins for sequencing analysis and peptide mapping." In: Proteolytic Enzymes: A Practical Approach. R.J. Beynon and J.S. Bond, eds., IRL Press, Oxford, 45.
22. Forler, D. et al. (2003) An efficient protein complex purification method for functional proteomics in higher eukaryotes. Nat. Biotechnol. 21, 89–92.
23. Geisse, S. et al. (1996) Eukaryotic expression systems: a comparison. Protein Expr. Purif. 8, 271–82.
24. Geu, B.U. et al. (1990) Characterization of glutamyl cleavage activity of endoproteinase Asp N. J. Prot. Chem. 9, 299–300.
25. Giniger, E. et al. (1985) Specific DNA binding of GAL4, a positive regulatory protein of yeast. Cell 40, 767–74.
26. Gosh, S. et al. (1998) NFκB and Rel proteins: Evolutionarily conserved mediators of immune responses. Annu. Rev. Immunol. 16, 225–60.
27. Hall, D.A., Ptacek, J. and Snyder, M. (2007) Protein microarray technology. Mech. Ageing Dev. 128, 161–7.
28. Hall, D.A. et al. (2004) Regulation of gene expression by a metabolic enzyme. Science 306, 482–4.
29. Hanes, B.D. (1981) In: Gel Electrophoresis of Proteins, Hanes, B.D. and Rickwood, D., ed.,IRL Press, Oxford, UK.
30. Hudson, M.E. and Snyder, M. (2006) High-throughput methods of regulatory element discovery. Biotechniques 41, 673, 675, 677 passim.
31. Hutchens, T.W. and Yip, T.T. (1990) Differential interaction of peptides and protein surface structures with free metal ions and surface-immobilized metal ions. J. Chromatogr. 500, 531–42.
32. Huzbun, T. et al. (2003) Assigning function to yeast proteins by integration of technologies. Mol. Cell. 12, 1353–65.
33. Ingrosso, D. et al. (1989) Specificity of endoproteinase Asp-N (Pseudomonas fragi): cleavage at glutamyl residues in two proteins. Biochem. Biophys. Res. Commun. 162, 1528–34.
34. Kaelin, W.G. et al. (1991) Identification of cellular proteins that can interact specifically with the T/E1A-binding region of the retinoblastoma gene product. Cell 64, 521–32.
35. Keil-Dlouha, V. et al. (1971) Proteolytic activity of pseudotrypsin. FEBS Lett. 16, 291–5.
36. Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–5.
37. Lane, D., Prentki, P. and Chandler, M. (1992) Use of gel retardation to analyze protein-nucleic acid interactions. Microbiol. Rev. 56, 509–28.
38. Li, S. et al. (2004) A map of the interactome network of the metazoan C. elegans. Science 303, 540–3.
39. LiCata, V.J. and Wowor, A.J. (2008) Applications of fluorescence anisotropy to the study of protein-DNA interactions. Methods Cell Biol. 84, 243–62.
40. Lin, Y.-S. et al. (1988) GAL4 derivatives function alone and synergistically with mammalian activators in vitro. Cell 54, 659–64.
41. Lonnerdal, B. and Keen, C. (1982) Metal chelate affinity chromatography of proteins J. Appl. Biochem. 4, 203–8.
42. Malloy, P.J. et al. (2001) A novel inborn error in the ligand-binding domain of the vitamin D receptor causes hereditary vitamin D-resistant rickets. Mol. Genet. Metabol. 73, 138–48.
43. Mankan, M.A. et al. (2009) NF-kappaB regulation: the nuclear response. J. Cell. Mol. Med. 13, 631–43.
44. Mann, M. and Kelleher, N.L. (2008) Precision proteomics: The case for high resolution and high mass accuracy. Proc. Natl. Acad. Sci. USA 105, 18132–8.
45. Mann, M. et al. (2001) Analysis of proteins and proteomes by mass spectrometry. Ann. Rev. Biochem. 70, 437–73.
46. Mannervik, B. and Danielson, U.H. (1988) Glutathione transferases—structure and catalytic activity. Crit. Rev. Biochem. 23, 283–337.
47. Markillie, L.M. et al. (2005) Simple protein complex purification and identification method for high-throughput mapping of protein interaction networks. J. Proteome Res. 4, 268–74.
48. Meyer, B. Papasotiriou, D.G. and Karas, M. (2010) 100% protein sequence coverage: A modern form of surrealism in proteomics. Amino Acids (Epub ahead of print).
49. Nilsson, J. et al. (1997) Affinity fusion strategies for detection, purification, and immobilization of recombinant proteins. Prot. Exp. Purif. 11, 1–16.
50. Pfleger, C. et al. (2001) Substrate recognition by the Cdc20 and Cdh1 components of the anaphase-promoting complex. Genes Dev. 15, 2396–407.
51. Porath, J. et al. (1975) Metal chelate affinity chromatography, a new approach to protein fractionation. Nature 258, 598–9.
52. Rice, R.H. et al. (1977) Stabilization of bovine trypsin by reductive methylation. Biochem. Biophys. Acta 492, 316–21.
53. Rosenfeld, J. et al. (1992) In-gel digestion of proteins for internal sequence analysis after one- or two-dimensional gel electrophoresis. Anal. Biochem. 203, 173–9.
54. Sadowski, I. et al. (1988) GAL4-VP16 is an unusually potent transcriptional activator. Nature 335, 563–4.
55. Sheer, D.G. (1994) Techniques in Protein Chemistry, Volume V, J.W. Crabb ed.,Academic, San Diego , 243.
56. Shevchenko, A. et al. (1996) Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68, 850–8.
57. Smith, D.B. and Johnson, K.S. (1988) Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione S-transferase. Gene 67, 31–40.
58. Solomon, M. et al. (1985) Formaldehyde-mediated DNA-protein crosslinking: A probe for in vivo chromatin structures. Proc. Natl. Acad. Sci. USA 82, 6470–4.
59. Solomon M.J., Larsen, P.L. and Varshavsky, A. (1988) Mapping protein-DNA interactions in vivo with formaldehyde: Evidence that histone H4 is retained on a highly transcribed gene. Cell 53, 937–47.
60. Stevens, J. and Kobs, G. (2004) FastBreak™ Cell lysis reagent for protein purification. Promega Notes 86, 23–24
61. Stevens, R.C. et al. (2001) Global efforts in structural genomics. Prot. Sci. 294, 89–92.
62. Swaney, D.L., Wenger, C.D. and Coon, J.J. (2010) Value of using multiple proteases for larger-scale mass spectrometry-based proteomics. J. Proteome Res. 9, 1323–9.
63. Terpe, K. (2002) Overview of tag protein fusions: From molecular and biochemical fundamentals to commercial systems. Appl. Microbiol. Biotechnol. 60, 523–33.
64. Tyers, M. and Mann, M. (2003) From genomics to proteomics. Nature 422, 193–7.
65. Wada, H. et al. (1998) Cleavage of the C-terminus of NEDD8 by UCH-L3. Biochem. Biophs. Res. Commun. 251, 688–92.
66. Wang, S. et al. (2001) Identification of a cellular protein that interacts and synergizes with the RTA (ORF50) protein of Kaposi's sarcoma-associated herpesvirus in transcriptional activation. J. Virol. 75, 11961–73.
67. Wang, Y. et al. (2000) The RIM/NIM family of neuronal C2 domain proteins. Interactions with Rab3 and a new class of Src homology 3 domain proteins. J. Biol. Chem. 275, 20033–44.
68. Wilkinson, J.M. (1986) “Fragmentation of polypeptides by enzymic methods." In: Practical Protein Chemistry: A Handbook, A. Darbre, ed., John Wiley and Sons, New York, 122.
69. Wilson, K.P. et al. (1992) Escherichia coli biotin holoenzyme synthetase/biorepressor crystal structure delineates the biotin- and DNA-binding domains. Proc. Natl. Acad. Sci. USA 89, 9257–61.
70. Wu, C.C. and MacCoss, M.J. (2002) Shotgun proteomics: Tools for the analysis of complex biological systems. Curr. Opin. Mol. Ther. 4, 242–50.
71. Wurm, F.M.et al. (1996) Production of recombinant protein therapeutics in cultivated mammalian cells. Nat. Biotechnol. 22, 1393–8.
72. Xu, Y. and Beckett, D. (1994) Kinetics of biotinyl-5′-adenylate synthesis catalyzed by the Escherichia coli repressor of biotin biosynthesis and the stability of the enzyme-product complex. Biochem. 33, 7354–60.
73. Yip, T.T. et al. (1989) Evaluation of the interaction of peptides with Cu(II), Ni(II), and Zn(II) by high-performance immobilized metal ion affinity chromatography. Anal. Biochem. 183, 159–71.
74. Young, N.L., Plazas-Mayorca, M.D. and Garcia, B.A. (2010) Systems-wide proteomic characterization of combinatorial post-translational modification patterns. Expert Rev. Proteomics 7, 79–92.
75. Zhu, H. et al. (2003) Proteomics. Annu. Rev. Biochem. 72, 783–812.

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