May 15, 2007 (Vol. 27, No. 10)
Getting There Quickly and Cost Effectively
Creating analytical tools that support the advancement of knowledge can often come about through the integration of existing information and techniques, theoretical and practical, directed toward a desired outcome. While the resulting construct itself may not be considered revolutionary, it can facilitate the pursuit of science and technology applications that may well be. This is a fair description of the developments leading to new instrumentation supporting affinity tag protein purification.
Protein purification methods typically seek to recover the protein analyte free of contaminants, in good yield, without denaturing the analyte’s biological activity or, alternatively, rendering the purified protein in a renaturable form. Accordingly, separation methods must be sufficiently mild so as not to irreversibly alter the protein’s structure. Techniques that meet this requirement include affinity-based methods that depend on noncovalent bonding interactions between a protein analyte and an immobilizing substrate.
Other methods that may support this objective include size exclusion chromatography; ion exchange chromatography; hydrophobic interaction chromatography; semipermeable membrane filtration; electrophoretic methods, such as SDS-PAGE; and various hybrid techniques employing variations and combinations of the foregoing.
While each of these techniques has its advantages and drawbacks, all are burdened with the requirement that each instance of application be tailored to a particular protein or class of proteins with common binding properties or structure. This requirement for specificity in method, materials, and purification conditions introduces a substantial bottleneck in the time required to carry out purifications across a large universe of differing protein types. As a result, significant time and labor that should be directed toward downstream applications are sacrificed in carrying out purifications.
Affinity tagging overcomes the limitations imposed by purification methods that must be adapted to protein sequence and conformation. The tag, a generic amino acid sequence, is designed for affinity interactions with a binding substrate that is independent of the parent protein structure. The tag sequence is incorporated into the recombinant protein by means of an expression vector positioned alongside the DNA sequence encoding the protein itself. Induction of the vector results in expression of the protein fused to the affinity tag, which can then be purified from the cell lysate.
After purification, the tag can be cleaved from the fusion protein at appropriately engineered molecular sites, typically in a buffer exchange step. Affinity tagging has made possible the expression and purification of large numbers of proteins by a single purification scheme. Starting from the crude biological recombinant product, nearly homogeneous proteins with levels of purity ranging from 90–99% can be routinely produced.
Of the number of affinity tag purification systems available, the most frequently employed utilize polyhistidine (His) or glutathione S-transferase (GST) tags. His binds with good selectivity to matrices incorporating Ni+2 ions, typically immobilized with either iminodiacetic acid or nitrilotriacetic acid chelating groups. The technique is known as immobilized metal affinity chromatography (IMAC; Figure 1). Absorption of the His-tagged protein is performed at neutral to slightly alkaline pH to prevent protonation and loss of binding capacity of the weakly basic histidine imidazole groups. Elution of the bound protein is caused by displacement with imidazole or low pH conditions.
IMAC is applicable to proteins with a broad range of physiochemical properties and structural variations. An important advantage of the technique is the possibility of purifying proteins under both native as well as denaturing conditions. IMAC is widely used in core labs that produce proteins for structural and functional studies.
In contrast to His-IMAC chelate binding, GST-tagged proteins bind noncovalently to an affinity matrix composed of glutathione immobilized on a chromatographic support. The tag is displaced and the protein eluted with solutions of glutathione. GST can be used alone or in combination with His tags to provide an alternate affinity purification regime when IMAC conditions are not suitable.
Limitations of First-generation Affinity Tag Technology
When first introduced, systems for carrying out affinity tag purifications were adapted from existing affinity purification instrumentation and techniques. As these modified systems and methods were deployed, their limitations became evident. First-generation instrumentation tended to be either complex automated fast protein liquid chromatographic (FPLC) systems or simpler manual methods. FPLC instrumentation is expensive and can be challenging to set up and operate. Such systems are generally not suited to a user community oriented toward the biological sciences and many of its associated laboratory techniques.
Manual protein purification techniques, such as those based on the use of spin and gravity-flow columns, are burdened, in part, by the lack of standardized affinity tag purification protocols. While these methods are relatively easy to run and far less expensive than FPLC, they are labor and time intensive, prone to handling losses, and often lack reproducibility. Moreover, these systems do not collect data and they require an additional desalting step with centrifugation that further extends the time required to generate purified protein.
Even FPLC systems, despite their automation, require some manual operations that lengthen purification processing times. These include lengthy setup procedures as well as a necessary interval for column regeneration and maintenance. While FPLC systems do automate the desalting process, manual intervention is needed to set up and program a separate desalting method and column and to physically transfer the purified protein onto the desalting column.
The foregoing assessment points to the need for relatively inexpensive turnkey instrumentation specifically designed for affinity tag protein purification. Such a system would avoid the need for complex method development and set-up protocols, thereby meshing with the skill set of the predominant user community. System automation would eliminate the time- and labor-intensive characteristics of current manual methods and enable the systematization and acceleration of affinity tag purifications without compromising protein quality or yield.
Bio-Rad Laboratories (www.bio-rad.com) recently introduced the Profinia Protein Purification System to address these requirements. Selectable methods are preprogrammed in conjunction with prepackaged buffer and cartridge kits (Figure 2). By collecting and integrating protocols that specifically support affinity tag purification, the system enables comparably rapid and efficient protein purification without sacrificing performance or product quality.
Purification methods for the most common affinity tag applications are factory programmed into the instrument and accessed through a touch-screen interface that guides selection and setup. The system’s plug-in purification kits with standardized buffers are integral to execution of the programmed methods. For example, matching coded labeling between kit components and instrument compartments ensures proper component installation. The instrument is also automated throughout and includes self-cleaning protocols that regenerate the system and prepare the purification cartridges for future use.
A dual-column design permits purification of two samples in series or automated purification and desalting plus buffer exchange of a single sample without user intervention. The main affinity peak is automatically detected and diverted to the desalting column. Column washes and elution fractions are collected in predesignated tubes for easy location of purified protein without needing to pool fraction tubes. Purifications can be conducted at ambient conditions or in a cold room. An accessory is also available to keep samples and collected fractions cold on the benchtop.
Yields and purity of processed proteins are equal to or better than results from either FPLC systems or labor-intensive manual methods. Considerable time savings are realized by elimination of complex system set up or the need for elaborate FPLC method development. System automation obviates much of the hands-on sample transfers and monitoring assessments required with manual procedures. Dependable and reproducible sample purifications (Figure 3) suitable for downstream applications are produced with significantly higher sample throughput compared to the other methods.
Each system was run according to its respective purification and desalting protocols. Purified and desalted protein concentration was determined by SDS-PAGE with UV detection. Post-desalting yield and purity of protein were nearly identical for both systems while the dedicated system purification was capable of a nearly threefold increase in throughput without the complex setups required by the FPLC system.
Results were evaluated with respect to yield, buffer consumption, processing time, and method scalability. Conductivity measurements of the dedicated system were taken in situ while those for the dialysis required sample removal and measurement with a calibrated conductivity meter.
Both procedures successfully desalted the protein sample with equivalent recovery of protein >90%. Desalting by the dedicated system was 8–11 times faster than dialysis and consumed 1/20 the amount of buffer. Concentration of dialyzed protein was twice that of the dedicated system’s gel filtration product, an advantage which must be weighed against the dedicated system’s superior throughput, greatly reduced buffer consumption, and automated, scalable capability.
The manual system was selected for its ability to replicate dedicated system affinity and desalting column volumes, 1 mL and 10 mL, respectively. Results indicate that the dedicated automated system has higher protein yields and superior reproducibility compared to the manual method. Protein product purity of both systems is equivalent.
The dedicated system’s ability to automate the collection of purification data, an option not available with the manual system, simplifies evaluation of the effectiveness of the purification. Unlike the automated system, the manual method requires the user to remain continually on site to apply buffer and collect fractions. It also requires manual transfer of the purified protein to the desalting column and subsequent centrifugation. The dedicated system executes both affinity purification and desalting without user intervention.
Affinity tagging of recombinant proteins enables the purification of a broad range of proteins purifiable by a single technique. As this science and enabling technology become more refined, considerable momentum has been generated toward the development of instrumentation to carry out purifications in a timely manner.
First-generation methodology and instrumentation, borrowed from existing applications, are compromised by limitations in the ability to exploit the affinity tag paradigm of a single purification technique across diverse protein structures. Emerging second-generation instrumentation is more closely tailored to the specific requirements of affinity tag purification.
These systems are helping to realize the promise of affinity tagging: the rapid purification of recombinant proteins at moderate cost and the recapture and deployment of valuable research time directed toward productive downstream applications.
Lee Olech is senior staff scientist at Bio-Rad Laboratories. Web: www.bio-rad.com
E-mail: [email protected].