Protein Science Encyclopedia - PDF Free Download

1 Microcontact Printing of Proteins Emmanuel Delamarche IBM Research, Z¨urich Research Laboratory, R¨uschlikon, Switzer...

Author: Sir Alan Fersht

54 downloads 1554 Views 131MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

1

Microcontact Printing of Proteins Emmanuel Delamarche IBM Research, Z¨urich Research Laboratory, R¨uschlikon, Switzerland

Originally published in: Nanobiotechnology. Edited by Christof M. Niemeyer and Chad A. Mirkin. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30658-9

1 Introduction

Biomolecules on surfaces have applications that range from medical diagnostics, analytical chemistry, and culturing and studying cells on surfaces, to synthesizing or engineering DNA, carbohydrates, polypeptides, or proteins. Defining patterns of biomolecules – and of proteins in particular – on surfaces is no simple task considering how complex and fragile these molecules can be. Photolithography – the art of structuring surfaces at lateral scales of less than 1 micrometer – was used recently to create DNA microarrays. Photolithography affords the capability of synthesizing strands of DNA using lithographic masks and photochemistry, but it is unlikely that a similar approach would permit the fabrication of arrays of proteins, which cannot be synthesized block-by-block at present. In photolithography, UV light, organic solvents, photoresists, and resist developers can compromise the structure and function of even a simple protein. For these reasons, novel approaches to patterning proteins include defining regions on surfaces that attract, bind, or repel proteins from solution, or in a more direct manner, by delivering small volumes of a solution of protein to a surface using drop-on-demand systems or microfluidic devices [1–3]. This chapter describes the patterning of proteins on surfaces by means of microcontact printing (µCP), where proteins are applied like ink to the surface of a stamp and transferred to a substrate by printing. Microcontact printing was originally developed by Whitesides and coworkers at Harvard for printing alkanethiols on gold with spatial control [4]. Many variants of µCP were later developed and collectively termed “soft lithography” [5]. The central element in µCP is the stamp. This is a silicone-based elastomer that is microstructured by curing liquid prepolymers of poly(dimethylsiloxane) (PDMS) Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Microcontact Printing of Proteins

on a lithographically fabricated master (or mold). Once cured, the stamp is peeled off the mold by hand; the stamp then bears an inverted pattern of that of the mold. One mold can be used to replicate many stamps. The relative softness of the stamp, compared to that of a lithographic mask, allows it to follow the contours of surfaces onto which it is applied. It is the work of adhesion between the stamp and the substrate that drives the spreading of the initial zones of contact at the expense of an elastic adaptation of the stamp [6]. In µCP – and soft lithography in general – the contact between the elastomer and a substrate occurs at the molecular scale and is termed “conformal”; it ensures the homogeneous transfer of ink from the stamp to the printed areas of the substrate [4]. PDMS materials have the following features. They are transparent to optical light and even UV down to ∼240 nm, resistant to many chemicals and pH environments, good electrical insulators, thermally stable, nontoxic, and can have tailored mechanical properties using various degrees of crosslinking and amounts of resin fillers [7]. A PDMS stamp can be a simple piece cut from a PDMS slab or accurately molded and mounted on a stiff backplane [8, 9]. It can be composed of PDMS layers having different mechanical characteristics, or shaped like a paint roller [10]. Handling stamps with tweezers and printing by hand is sufficient for most needs of experimentalists. Mounting a stamp on a printing tool, however, provides the ability to vary and control the pressure applied during printing, and to align the stamp with the substrate. PDMS stamps have advancing and receding contact angles with water of ∼115◦ and ∼95◦ and thus are hydrophobic and promote the spontaneous deposition of proteins from solution [11]. This deposition is nonspecific and self-limiting to a monolayer of proteins if the stamp is rinsed after the inking step. An important difference between microcontact printing proteins on surfaces and alkanethiols on noble metals is the limited amount of proteins on the stamp. Alkanethiols can diffuse inside a PDMS stamp in sufficient amounts for multiple prints, whereas reinking stamps with proteins is necessary after each print unless hydrogel “stampers” are used [12]. These stamps can carry a large reserve of protein solution used for the contact processing (CP) of substrates. PDMS stamps derivatized with biomolecules provide the basis for selective inking strategies and lead to the affinity-contact printing (αCP) technique [13]. Figure 1 delineates the operations that CP, µCP, and αCP involve. These techniques are described in detail in the following sections.

2 Strategies for Printing Proteins on Surfaces 2.1 Contact Processing with Hydrogel Stamps

Contact processing (left-hand panel in Figure 1) mimics the deposition of proteins from an aqueous environment to a surface by utilizing a hydrogel swollen with a solution of protein [12–14]. The proteins can diffuse through this hydrophilic matrix and adsorb onto the substrate without uncontrolled spreading. The stamp

2 Strategies for Printing Proteins on Surfaces

Fig. 1 Three related methods can pattern proteins from a stamp to a surface. In contact processing (left), a hydrogel stamp mediates the diffusion of proteins from its bulk to a surface. Microcontact printing (center) utilizes an elastomeric stamp inked with

proteins to print the proteins on a substrate without having a liquid. The stamp in affinity contact printing (right) is derivatized with capture proteins, which allows it to be selectively inked with target proteins released to a substrate during printing.

consists of two parts. The first is a reservoir above the hydrogel containing proteins dissolved in a biological buffer. The second is the hydrogel that makes contact with the substrate and mediates the transport of proteins to the substrate. A stamp that has a hydrogel made of poly(6-acryloyl-β-O-methyl-galactopyranoside), for example, embedded in a fine capillary can pattern proteins with a resolution of ∼20 µm [14]. Hydrogels having a refined composition and a greater degree of crosslinking exhibited better mechanical resistance, and were patterned by replication of a mold [15]. The latter approach should allow a protein to be patterned on a surface with a resolution better than 20 µm. CP based on hydrogel stamps has interesting features. First, biomolecules remain in a biological buffer until the stamp is removed and the substrate dried. Denaturation of proteins in this case should be minimal, and may be similar to that of proteins adsorbed from solution onto polystyrene microtiter plates. Second, it is straightforward to reuse such stamps for multiple CP experiments [14]. 2.2 Microcontact Printing

Microcontact printing of proteins uses PDMS stamps replicated from a mold (middle panel in Figure 1). Inking the stamp with proteins is simple, and analogous to depositing a layer of capture antibody (Ab) on polystyrene for conducting a solidphase immunoassay. The duration of inking and the concentration of protein in the

3

4 Microcontact Printing of Proteins

ink solution determine the coverage of protein obtained on the stamp [16]. Inking a stamp can be local and/or involve multiple types of proteins when the stamp is locally exposed using a microfluidic network (µFN) or microcontainers to one or more solutions of protein [17]. The transfer of proteins can be remarkably homogeneous and effective, depending on the wetting properties of the substrate [18]. The large area of interaction of proteins with substrates and their high molecular weight account for the high-resolution potential of µCP of proteins. At the limit, single protein molecules can be printed as arrays on a surface [16], whereas the diffusion of alkanethiols on noble metals or the reactivity of silanes with themselves limit the practical resolution achieved for microcontact printing self-assembled monolayers on surfaces. Microcontact printing proteins on surfaces appears to be limited by the resolution and mechanical stability of the patterns on the stamp. Stamps made of Sylgard 184 and using masters prepared using rapid prototyping or photolithography can have micrometer-sized patterns on fields even larger than 10 cm2 [19]. Microcontact printing proteins with arbitrary patterns and submicrometer resolution benefits from the use of a PDMS elastomer stiffer than Sylgard 184 and masters patterned using electron-beam lithography [8]. 2.3 Affinity-Contact Printing

Tailoring the surface chemistry of stamps to ink a particular type of biomolecule is crucial for αCP (right-hand panel in Figure 1). The chemical stability of silicone elastomers is both an advantage for preparing chemically resistant stamps and an obstacle to modifying the surface of PDMS stamps. Exposing PDMS to an oxygen-based plasma forms a glassy silica-like surface layer [20]. The oxidized layer is a few nanometers thick and contains silanol groups ( Si OH), which are useful for anchoring organosilanes [21]. Oxidized PDMS can thus be derivatized similarly to glass or SiO2 in a few chemical steps using silane monolayers and with crosslinkers for proteins [22]. Affinity-contact printing is the technique of covalently immobilizing ligand biomolecules onto a PDMS stamp, and using them to ink a stamp selectively with receptor molecules. A stamp for αCP is roughly analogous to a chromatography column due to its ability to extract proteins selectively from a mixture, although releasing them involves printing them onto a surface [13]. Biomolecules that are naturally present in crude solutions and have a function on a surface are ideal candidates for applications of αCP. Cell adhesion molecules is one example that has already been demonstrated, but αCP could well be extended to a large variety of biomolecules for which ligands exist. Stamps in αCP are reusable and may include sites of different affinity to capture and print multiple types of protein in parallel [23]. 3 Microcontact Printing Polypeptides and Proteins

Many different types of proteins can be inked from an aqueous solution onto a hydrophobic silicon rubber such as PDMS [24]. Hydrophobic polymers in general

3 Microcontact Printing Polypeptides and Proteins

promote the deposition of proteins from solution through a variety of interactions, and slight or pronounced conformational changes of the protein structure can accompany this adsorption process. The kinetics of formation of a layer of protein on hydrophobic surfaces is often compared to a Langmuir-type isotherm: the rate of deposition of the protein molecules scales with their concentration in the bulk of the solution and reaches a plateau when all sites on the substrate become occupied [25]. Hydrophobic substrates, as a general rule, have stronger interactions with hydrophobic proteins, and their adsorption process is less influenced by the pH and ionic strength of the solution and by the isoelectric point of the protein than when polar or charged substrates are employed [25]. The size of the protein does not seem to play an important role on their inking behavior. A wide range of proteins in terms of structure and functions has been microcontact printed, which includes cytochrome c (12.5 kDa) [11], streptavidin and bovine serum albumin (BSA; ∼60 kDa) [26–28], protein A and immunoglobulins G (150 kDa) [11], glucose oxidase (160 kDa) [29], laminin (∼210 kDa) [30], and fibronectin (440 kDa) [31]. It is sometimes necessary to employ stamps with a hydrophilic surface to ink hydrophilic polypeptides such as polylysine (with MW ranging from 38 to 135 kDa) [32] or lipid bilayers [33]. In other cases, small biomolecules such as amino-derivatized biotins were inked and printed onto surfaces reactive to amino groups [34, 35]. In general, the derivatization of biomolecules with thiol groups allows the printing of biomolecules on gold substrates [36], where patterning by printing can be complemented by the adsorption of other types of molecules from solution. The chemisorption of small biomolecules on surfaces might be necessary for efficient transfer from the stamp and to prevent rinsing the printing molecules during subsequent steps. 3.1 Printing One Type of Biomolecule

Immunoglobulins G (IgGs) are interesting candidate molecules for µCP: these Abs are useful on surfaces for heterogeneous immunoassays. Their numerous disulfide linkages make them robust, they adsorb from biological buffers to PDMS in a nonreversible manner [24], and they can be conjugated to fluorescent centers, metal particles, enzymes, or ligands such as biotin. Fluorescence microscopy is a versatile method to follow the results of microcontact printing IgGs onto a glass surface (Figure 2). There, TRITC-labeled anti-chicken Abs were inked everywhere on the stamp but transferred to glass only in the regions of contact [11]. The patterns on the glass are accurate and correspond to zones of the stamp where the inked proteins are missing. The contrast of the 1 µm-wide features in the pattern is high and accurate and, as no fluorescence above background is measured in the nonprinted regions, it is clear that no transfer of Ab occurred in the recessed areas of the stamp. This might not always be the case, because small features have limited mechanical stability [6]. Demolding the stamp from the mold, capillary effects during inking and drying the stamp, and the printing itself may compromise the mechanical stability of patterns [37]. Implementing support structures in the design of the pattern, controlling the forces exerted during printing and affixing a

5

6 Microcontact Printing of Proteins

Fig. 2 Microcontact printing proteins on glass. Fluorescence microscopy images revealing TRITC-labeled chicken Abs on a stamp after inking and accurately transferred in the regions of contact to a glass substrate. Reproduced with permission from Ref [11]. (Copyright 1998 American Chemical Society.)

stiff backplane to the stamp improve the stability of patterns. Stamps can be very large and have features measuring from micrometers to centimeters, making it possible to print proteins of one kind on large substrates to pattern cells indirectly. Examples include microcontact printing fibronectin [31], polylysine [30, 38, 39], laminin [40], and adhesion peptides [41]. 3.2 Substrates

Substrates for biomolecules cover a wide range of materials, from simple glass slides to complex functional microelectronic devices or sensors. Having conformal contact between the stamp and substrate during printing is the first requirement for microcontact printing biomolecules. For this reason, the substrate should not be too rough [6], or have too prominent structures [39]. Polystyrene, poly(styrene terephthalate), glass, amphiphilic comb polymers, Si wafers, and substrates covered with a thin evaporated metal and/or a self-assembled monolayer can be microcontact printed with proteins and stamps made of Sylgard 184 [11, 29, 34, 42]. The printing time does not seem to play a role, and takes the few seconds necessary to propagate the initial contact to the entire substrate. The details of how and why proteins transfer from a stamp to a surface were intriguing until the recent discovery that the difference in wettability by water between the stamp and the surface determines whether transfer occurs [18]. Proteins tend to transfer when the substrate is more wettable, or has a higher work of adhesion for water, than the stamp. In this respect, the chemical composition of the surface does not seem to play a particular role other than defining the wettability of the surface (Figure 3). The surface of the stamp can be derivatized with fluorinated silanes to raise the wettability threshold

3 Microcontact Printing Polypeptides and Proteins

7

8 Microcontact Printing of Proteins

of the substrate below which transfer remains effective, for example. A remarkable incidence of printing proteins occurs with poly(ethylene glycol) (PEG)-derivatized surfaces [18]. Surfaces covered with a sufficient density of PEGs resist the deposition of proteins from solution because in order to interact with the PEG layer, proteins have to remove water solvat-ing EG repeat units and reduce the number of possible conformations of the PEG chains [43]. Both of these requirements are energetically unfavorable to the deposition of proteins from solution onto PEG-treated surfaces [44]. The mechanism accounting for the transfer of protein in µCP might thus involve the dry state of the PEG layer during printing [43], the local pressure exerted by the stamp at the line front propagation of the conformal contact [45], or some contamination of the PEG layer by low-molecular-weight silicone residues from the stamp. The deposition of proteins from solution or by printing exhibits antagonistic behaviors: proteins are more difficult to print on a hydrophobic surface than on a hydrophilic one whereas the opposite situation generally occurs in solution with, as an extreme case, PEG surfaces, which are protein-repellant [46]. Printing proteins is not limited to the patterning of planar substrates but is possible on curved surfaces, structured surfaces, and over large areas [5]. A stamp can be molded directly curved or planar and then curved and rolled over a surface [10]. Large stamps (≥ 10 cm) can be molded with a pattern having an accuracy of better than 1 mm [47]. The mechanical properties of stamps can be varied from 1 MPa (Young’s modulus) to over 30 MPa by adjusting the formulation of the polymer with respect to its average molecular weight between junctions, the junction functionality, and the density and size of filler particles added to the polymer [8]. The hardness of a stamp, its work of adhesion with the substrate, the pressure applied during printing, and the topography and work adhesion of the substrate all determine whether conformal contact will occur. The stability of features on the stamp might be compromised, however, when the stamp is made too soft and pressed too hard during printing [6, 9, 48]. 3.3 Resolution and Contrast of the Patterns

High resolution in lithography refers to patterning features of arbitrary shape at a length scale where it becomes crucial to optimize all parameters of the technique ← Fig. 3 Influence of the wettability of the substrate by water on the degree of transfer of proteins from a PDMS stamp to a Au surface. (A) The substrate is derivatized with SAMs comprising variable mole fractions of two constituents having different end-groups. (B) Proteins are adsorbed from solution onto the stamp. (C) The resulting printed patterns are analyzed using fluorescence microscopy. The fluorescence micrograph corresponds to fluorescently la-

beled proteins printed onto a 100% COOHterminated SAM. The transfer of proteins followed on SAMs having hydrophilic components functionalized with (D) COOH, (E) OH, or (F) EG correlates with the wettability of the mixed SAM (G). Figure kindly provided by J. L. Tan, J. Tien and C.S. Chen, and reprinted with permission from Ref. [18]. (Copyright 2002 American Chemical Society.)

3 Microcontact Printing Polypeptides and Proteins

(e. g., conditions for exposing and developing the resist, transfer of the resist pattern onto the substrate). Electron-beam lithography has a high-resolution regime for making features < 100 nm, photolithography for features < 250 nm, and µCP for features < 500 nm. In conventional lithography, shrinking the dimensions of patterns is driven by the necessity to improve the performance of integrated circuits at invariant or lower cost. The resolution of lithographic techniques limits the smallest sizes of components made today. It will be the physics of tomorrow’s devices that will ultimately be the limiting obstacle to further integration. Patterning biomolecules has a different paradigm for the resolution limit than conventional lithography because single functional elements, an enzyme for example, are available but do not have to be constructed. Microcontact printing meets several requirements that are necessary to place single proteins at predefined positions on a surface: (i) it is possible to fabricate Si molds with features as small as 40 nm using electron-beam lithography [16]; (ii) PDMS-replicated structures can be 80 nm and even smaller [9, 47]; (iii) proteins remain in the areas of contact, unlike alkanethiols and monolayerforming molecules which generally diffuse away from the initial printed zones on the substrate when an excess ink is present on the stamp; and (iv) the solution of protein used to ink the stamp can be diluted to limit the number of proteins inked per feature on the stamp [16]. Figure 4 shows high-resolution patterns of Abs on Si and glass and how a highresolution stamp can look. Each feature in the atomic-force microscopy (AFM) image in Figure 4A comprises ∼1000 Abs of the same type that were printed on a Si wafer using a PDMS stamp made of Sylgard 184 [17]. The structures have a width of 500 nm and an edge resolution better than 50 nm. The contrast of the patterns seems perfect because no Ab is present outside of the printed areas. An excellent contrast, together with specific binding events between printed ligands and receptors from solution, are desirable for high-sensitivity biological assays. The photography in Figure 4B shows a 8 × 4 cm2 stamp composed of a 30 µm-thick layer of PDMS attached to a flexible glass backplane 100 µm thick [8]. The PDMS layer of this stamp has numerous fields with 250 nm-wide lines, is about five times harder than Sylgard 184, but is also more brittle. The patterns are consequently more stable against collapse, and the glass backplane contributes significantly to the long-range accuracy of the pattern while making the stamp simple to handle, mount and align on a printer tool [47]. The surface tension of the polymer is an important limiting factor for the resolution of µCP. Features as small as 5 nm can be written by electron-beam lithography in PMMA and developed. PMMA is brittle, however, and hence not soft enough to form a reliable contact over surfaces. Unmolding even relatively hard PDMS stamps (having a Young’s modulus > 12 MPa) from a master structured with 40 nm-wide lines results in their broadening by 20 nm owing to surface tension effects [16]. Such a broadening is relatively less important for stamps with features ≥ 100 µm, and can be compensated in the electron-beam lithography layout. The stability of small features on stamps limits the freedom of design for high-resolution patterns. Dots, lines, and meshes do not have all the same mechanical stability against pressure; some recessed areas may collapse during printing. Incorporating support structures with micrometer dimensions around the high-resolution fields can remedy these problems [6]. Another strategy

9

10 Microcontact Printing of Proteins

Fig. 4 Microcontact printing proteins at high resolution. (A) AFM images showing that only ∼1000 chicken Abs were printed onto a Si wafer in each element of this pattern. (B) High-resolution :CP is best done using stamps harder than Sylgard 184 and

having a flexible glass backplane. (C) AFM images showing rabbit Abs printed on glass as a mesh comprising 100 nm-wide lines (left) or on regularly spaced areas that have none, one or a few Ab molecules (right).

4 Activity of Printed Biomolecules

is to transfer the resist pattern into the Si master with a reactive ion etching, where the etch rates depend on the geometry of the features. When large structures are made deeper in the master than smaller structures, a larger part of the load during printing is exerted on the larger structures without inducing collapse of the smallest features [16]. The AFM images in Figure 4C correspond to Ab molecules microcontact printed from a PDMS stamp (material B) [8] onto glass using a mesh of 100-nm-wide lines (left image) and 100-nm hemispherical posts [16]. Both the posts and the lines were 60 nm high. The detail of a mesh reveals that two to four Ab molecules define the width of the lines. In the case of posts, one to three Ab molecules occupy each visible printed site, and the statistical analysis of larger printed zones revealed that sites could have none, one or a few printed Ab molecules [16]. There, the resolution limit for microcontact printing a single molecule is reached while still leaving space for improvement to form homogeneous arrays having only one biomolecule per site. A high concentration of protein in the ink, a long inking time, a further reduction of the dimensions of the posts, and a substrate with a high work of adhesion could help printing large arrays of single protein.

4 Activity of Printed Biomolecules

Many studies emphasize that while the adsorption of a protein on a surface is simple to perform, it is nevertheless a complicated phenomenon in which the biological activity of the immobilized biomolecule might be lost or significantly altered [25]. Microcontact printing biomolecules harbors this risk twice: when proteins are inked onto the stamp, and when they are printed. In principle, the deposition of proteins from solution onto PDMS should be analogous to that of proteins on hydrophobic surfaces [24]. The second concern is more difficult to weigh. Transferring a protein by printing implies that the adhesion of the protein with the substrate overcomes that of the protein with the stamp. At the limit, this might create a mechanical stress on the protein and could lead to irreversible conformational changes. It might be interesting to evaluate the yield of transfer as a function of the peeling rate to better characterize the transfer mechanism [49]. Comparing the activity of different types of biomolecules printed or adsorbed onto polystyrene suggests that enzymes are more susceptible to denaturation during printing than during adsorption from solution [11]. A layer of printed proteinase K displayed half the activity of a layer deposited from solution. Abs are more robust against loss of function; the ability of printed polyclonal Abs to capture antigens from solution was similar to that where the captured Abs were adsorbed. Printed monoclonal Abs seemed to have a ∼10% loss of capture efficiency compared to ones adsorbed from solution. The surface activity of microcontact printed biomolecules belonging to three important biological classes is illustrated in Figure 5. Cell adhesion molecules are ideal candidates for printing biomolecules because these molecules are useful on surfaces as they can direct the adhesion and growth of cells to specific regions of a substrate. Moreover, many are “simple” polypeptides. Polylysine [30, 32, 38, 39],

11

12 Microcontact Printing of Proteins

Fig. 5 Microcontact printed proteins preserve a sufficiently high degree of activity for (A) promoting cell adhesion, (B) performing immunoassays, and (C) performing surface enzymatic catalysis. (A) The adhesion peptide PA22-2, which was printed onto a thiol-reactive surface, was immunostained (left-hand fluorescence micrograph) in two steps using an anti-PA22-2 Ab and a secondary Ab labeled with fluorescein, and is useful for attaching hippocampal neurons with spatial control (phase-contrast micrograph on the right). (B) These AFM images

illustrate three steps of an immunoassay in which chicken Abs were printed on a Si wafer (left), BSA was adsorbed from solution during the blocking step (middle), and the printed Abs were recognized by anti-chicken Abs. (C) This fluorescence image shows the deposition from solution of a fluorescent product made insoluble by printed alkaline phosphatase. The images ¨ in (A) were kindly provided by Offenhausser et al. and reprinted from Ref. [41]. (Copyright 2000 with permission from Elsevier Science.)

laminin [50], polylysine fused with laminin [40], fibronectin [31], specific adhesion peptides [41, 42], and neuron-glial cell adhesion molecules [13] (NgCAM), for example, were microcontact printed to promote or guide the attachment of cells to surfaces. In other examples, Ab–cell interactions were used to pattern cells on patterns of printed Abs [51, 52]. In some instances, the stamp was made hydrophilic and the substrate activated with a crosslinker to increase, respectively, the inking

5 Printing Multiple Types of Proteins

and transfer efficiency. The left-hand fluorescence image in Figure 5A corresponds to the immunostaining of the adhesion peptide PA22-2 that was microcontact printed onto a glass surface activated with amine-reactive crosslinkers [41]. The phase-contrast image (right) shows that the printed pattern of peptide was suitable to grow viable hippocampal neurons in the printed regions specifically These results illustrate well the conservation of the function of printed adhesion molecules. The capability of printing a pattern in registry with structures predefined on a substrate opens the way to placing cells wherever desired on a complex surface to study their function and to form networks of immobilized cells [39, 50, 53]. The next example (Figure 5B) is a printed polyclonal Ab, which serves as antigen to bind polyclonal Abs from solution [11]. AFM reveals the printed regions of a Si surface, each of which comprises ∼1000 molecules of chicken Ab molecules. Blocking the free Si surface with BSA is the next step necessary to prevent nonspecific deposition of proteins during the recognition step. After the blocking step, the Si surface is either covered with BSA or printed Abs; the prints are no longer visible. Recognition of the printed chicken Abs by anti-chicken Abs faithfully reflects the printed pattern. This experiment is an example of a highly miniaturized surface immunoassay in which the printed antigens were recognized by their specific Abs. Enzymes are probably more fragile than Abs and suffer more from a random orientation on a surface with respect to their activity than antigens for polyclonal Abs. The activity of printed enzymes can be evaluated using flat stamps, polystyrene substrates and colorimetric measurements [11]. It can be useful, however, to keep enzymatic products near their sites of production and to assess the activity of the enzymes with high spatial resolution. This is possible by using precipitating fluorescent products that accumulate in the regions of the substrate having enzymatic activity (Figure 5C) [17]. Real-time analysis of the development of fluorescence on the printed sites is even possible with this method. At least a part of the alkaline phosphatases printed on the glass surface in Figure 5C are active. This suggests that enzyme-linked immunosorbent assays can be performed using captured Abs that are printed. Interestingly, the same type of reporter enzyme can unambiguously reveal an ensemble of binding events, which are discernible through their localization. The challenge in this case remains to derivatize a surface with several types of protein.

5 Printing Multiple Types of Proteins 5.1 Additive and Subtractive Printing

An obvious application for microcontact printing proteins is the preparation of protein microarrays that can be used to screen different analytes in parallel while conserving reagents and still obtaining high-quality signals [54–56]. The simplest method to place two types of protein on a surface is to print one and adsorb the other from solution, as has been done with two different types of Abs [11]. This strategy requires that the printed layer of protein be complete enough to limit the

13

14 Microcontact Printing of Proteins

adsorption of Abs from solution into the printed regions. The fabrication of arrays comprising n types of protein is possible using additive patterning steps: once a substrate is microcontact-printed, more proteins can be printed next to or over the previously printed ones (Figure 6A) [17, 57]. The transfer of proteins from a hydrophobic stamp to a more wettable substrate accounts for this finding because surfaces covered with printed proteins are more hydrophilic than PDMS stamps. This ability to stack proteins on top of each other is peculiar to µCP and might be useful for constructing protein-based architectures. It is also possible to place a variety of proteins on a substrate without the need for precise alignment during printing. Stamps with parallel lines can, for example, be inked and printed with a rotation between each print [17, 57]. Subtractive approaches are also possible (Figure 6B). One strategy is to ink a flat stamp homogeneously and to remove a subset of the proteins by printing them onto a structured surface. The sites on the stamp made free for adsorption are then covered by proteins from solution and the operation can be repeated [17]. An original way to pattern a surface with multiple types of protein is to fabricate a three-dimensional stamp and ink the different layers of the stamp with different types of proteins. Applying increasing pressure to the stamp brings each layer of the stamp successively in contact with the substrate [58]. The different patterns of protein are inherently aligned, and the difficulty of fabricating and inking the stamp might be compensated for by the relatively simple printing operation. 5.2 Parallel Inking and Printing of Multiple Proteins

Serial methods are simple, but probably not suitable, to pattern substrates with a large number of different proteins. A parallel inking approach of a stamp using a µFN can solve the problem of inking a stamp with different types of proteins (Figure 6C) [17, 59]. With such a strategy, a µFN having an ensemble of independent flowing zones is placed on a flat PDMS stamp [60, 61]. Sealing the microchannels results from the conformal contact between the µFN and the stamp. When solutions of proteins are flushed through the microchannels, proteins deposit in the areas of the stamp exposed to the conduits. Filling a µFN can be done serially, or with an array of dispensing heads or tips. The deposition of protein on the PDMS might be as fast as a few seconds when it is not limited by the mass transport of proteins from solution or the depletion of proteins from the channels. There, inking the stamp with or without a µFN could involve pin spotting, drop-on-demand, or microinjection techniques. Prefilling individual wells of a structured surface and applying it to a PDMS surface is also suited to locally derivatize a stamp with different types of proteins [23]. 5.3 Affinity-Contact Printing

The inking of a stamp with a large number of different types of protein before each print can quickly become cumbersome when it is desirable to print a large series

5 Printing Multiple Types of Proteins

Fig. 6 Fluorescence microscopy images illustrating three strategies for printing several types of protein on a surface. (A) Two fluorescently labeled proteins were printed subsequently on a glass surface. Proteins on the stamp transferred during the second print both to glass and to the lines of proteins already patterned. (B) This pattern includes two fluorescently labeled protein and BSA printed simultaneously from a flat stamp. First, BSA was inked homogeneously by adsorption from solution onto

the stamp and patterned by subtractive printing. Proteins labeled with fluoroscein isothiocyanate (FITC) were then adsorbed in the regions complementary to the BSA pattern. This was repeated to remove BSA and the second protein along lines that were filled with TRITC-labeled protein adsorbed from solution. (C) A stamp was inked with different lines of proteins using independent channels of a :FN and printed onto a polystyrene surface in one step. Reproduced from Ref. [17].

15

16 Microcontact Printing of Proteins

of substrate with the same pattern of protein. The ability to attach biomolecules to PDMS covalently yields the opportunity to define zones on a stamp that can actively bind a target molecule from an ink. Crosslinking capture molecules (e.g., Abs or antigens) to a stamp allows inking the stamp by exposing it to a solution containing targets for the surface-bound capture molecules. This inking strategy, termed αCP, is analogous to capturing a protein on a column for affinity chromatography, although release of the captured species can occur during printing (Figure 7A) [13]. The idealized view of αCP is to have an affinity stamp (α-stamp) with multiple sites for capturing target molecules from solution in parallel. This could be used for many cycles of capture and printing. The fluorescence image in Figure 7B corresponds to the detection of fluorescently labeled Abs that were inked onto an α-stamp and printed on a glass substrate. This stamp had two types of binding sites (antigens) that were crosslinked to PDMS with spatial control by means of subtractive printing and ordinary printing [23]. The α-stamp used to print the Abs in Figure 7C was prepared by attaching capture antigens to a stamp activated with a crosslinker for proteins using a µFN. The yellow line corresponds to the printing of TRITC and FITC-labeled Abs that were simultaneously captured on a line of protein A immobilized on the stamp. Affinity contact printing, in particular when it employs stamps having multiple affinity sites, is both powerful and challenging. It is powerful because the ink can be a complex solution of biomolecules, the stamp is reusable, and patterns can have high resolution, as in µCP. The difficulty of αCP lies in the preparation of the α-stamp because the PDMS surface must be derivatized with crosslinkers and the quality of the patterns may degrade when preparation involves a large number of steps. The capture and release of radioactive or fluorescent proteins using αCP demonstrated the specificity of the capture event and the reusability of the α-stamp for at least 10 cycles [13]. Neuron-glial cell adhesion molecules (NgCAM), which are 200 kDa transmembrane proteins present at a concentration of ∼1 mg mL−1 in membrane homogenates of chicken brain, were captured by monoclonal Abs of an α-stamp and patterned on a polystyrene surface. The patterned surface appeared to be suitable for the attachment and growth of dorsal root glial neurons (Figure 7D and 7E), which was not the case where polystyrene was exposed directly to a nonpurified source of NgCAM [13]. Among all the printing methods reviewed here, αCP might be the most powerful method owing to the selective inking step and the reusability of α-stamps. It also has, in principle, the potential to print biomolecules with a defined orientation on a surface.

6 Methods 6.1 Molds and Stamps

Molds for preparing stamps are most often Si wafers patterned with a combination of photolithography and reactive ion etching. Reactive ion etching Si, instead of

6 Methods

Fig. 7 Affinity contact printing. (A) Stamps for "CP are prederivatized with capture sites and alternatively inked selectively and used for releasing the captured molecules on a substrate during the printing step. The fluorescence microscopy images correspond to (B) fluorescently labeled Abs co-captured and co-printed on glass using an "-stamp that had two types of capture sites; (C)

fluorescently labeled proteins captured on lines of an "-stamp and printed on glass; (D) the immunofluorescent detection of NgCAM that was patterned on polystyrene using an "-stamp decorated with lines of anti-NgCAM mAbs; and (E) the staining by immunofluorescence of neurons which attached to and developed on the printed pattern of NgCAM.

17

18 Microcontact Printing of Proteins

using directly the pattern of photoresist as the mold, prolongs the lifetime of molds. Si molds can be washed and cleaned; passivation of the Si surface is necessary before pouring PDMS with a release layer. Passivation can be done in situ after reactive ion etching or using a simple dessicator that can be evacuated. Typical release agents are fluorinated silanes. High-resolution molds are Si wafers patterned using electron-beam lithography. The density and resolution of the high-resolution features determine the price of these molds, which easily reaches $1000 per written cm2 for features having a critical dimension of 100 nm or less. Rapid prototyping is suitable for the preparation of stamps with a resolution of ∼25 µm [62] and only requires access to a high-resolution printer (5000 dpi). Replicating molds to make stamps and other operations such as inking stamps and printing are best done in a clean room or using a laminar flow bench to minimize the contamination of surfaces by dust particles. Sylgard 184 (Dow Corning) is used to prepare stamps in many cases, and comprises two prepolymers which, once mixed at a ratio of 1:10 (catalyst and hydridosiloxanes:vinyl-functionalized siloxanes), are poured on the master and cured at 60◦ C overnight. The formulation of harder, mechanically more stable PDMS is sometimes necessary when stamps have tall, isolated features [8]. Stamps can be cured on a backplane made of a thin steel plate or glass sheet [47], or on another PDMS layer [63]. Backplanes are typically flexible, but nevertheless give dimensional and long-range stability to stamps, which can then be mounted on a mask aligner, a printer, or handled by hand more conveniently. Design rules to make stamps are described jointly with the analytical description of the formation of conformal contact between stamps and substrates [6, 48]. This information is valuable to estimate the mechanical stability of stamps against pressure and to predict whether conformal contact will occur on rough surfaces or surfaces having topography. Hydrogel stamps are composed of a polymer that is crosslinked to the desired value (2–4%) and embedded in a microcapillary [12] or patterned by photocuring the hydrogel precursor sandwiched between a slide and a mold [15]. Both the inking and printing with hydrogel stamps rely on the diffusion of proteins through the hydrogel medium. 6.2 Surface Chemistry of Stamps

PDMS is hydrophobic and promotes the adsorption of proteins from solution in a manner analogous to polystyrene. Modifying the surface chemistry of stamps is necessary in two cases. Polypeptides and homogeneously polar biomolecules require stamps to have a hydrophilic surface for inking [38]. Affinity stamps must be derivatized with a ligand specific for biomolecule targets [13]. Exposing a PDMS stamp to an O2 -based plasma creates a silica-like layer on PDMS in a self-limiting manner. Stamps should be freshly inked (within ∼5 min) after the plasma treatment to prevent the recovery of their hydrophobic character [20, 22, 46]. This hydrophobic recovery originates from the migration of low-MW silicone residues from the bulk to

6 Methods 19

the surface. The plasma-induced scission of some polymer chains might also create mobile residues. It is impractical to extract these residues from the prepolymer components or after polymerization. Instead, plasma-treated stamps can be kept under water for long periods of time (more than days). Anchoring crosslinkers for proteins on plasma-treated stamps in one or more steps permits attaching covalently ligands onto the stamps [23]. Unreacted crosslinkers can be quenched with chemicals or reacted with noninterfering proteins such as BSA. 6.3 Inking Methods

The time to ink a stamp with a full monolayer of protein can be relatively long: up to 30 min at room temperature to obtain a monolayer of Ab using a concentration in phosphate-buffered saline (PBS) of 1 mg mL−1 , and 45 min with a solution of 5 µg mL−1 [11, 16]. The stamp is rinsed and dried after the inking step and then placed in contact with a substrate. The inking of hydrogel stamps with 1 mg mL−1 solutions of Abs in PBS takes similar times, and might even be faster if the gel is initially dry [12]. Shortening the inking time of PDMS stamps and localizing the inking is possible using µFNs [59]. In this case, the adsorption of proteins to the stamp might not be mass transport-limited, and is local in the regions of the stamp exposed to the channels. Localized inking is equally possible using microcontainers. These are small reservoirs microfabricated in Si, for example, and filled with the same, or different, solutions of proteins by hand or using pipetting robots [23]. Subtractive inking corresponds to inking entirely a flat PDMS stamp with proteins and transferring a subset by printing on a structured target. This strategy can remove proteins from areas of the stamp that become free for the inking of other proteins from solution [17, 23, 57]. This method can be repeated to form patterns with different types of protein next to each other but it requires an alignment step. Inking a stamp for αCP is analogous to linking a protein to a chromatography column via NH2 residues. Crosslinking protocols are usually well detailed by chemical suppliers or reviewed elsewhere [64]. 6.4 Treatments of Substrates

Surfaces more wettable by water than PDMS stamps are, in principle, suited for printing proteins [18]. Otherwise, they can be derivatized appropriately using plasma deposition techniques, oxidizing methods, or by grafting ultrathin organic layers. The wettability criteria may not apply when hydrophilic stamps are used to print certain polar biomolecules. In this case, derivatization of the substrate might also be necessary. Polylysine has been printed on glass directly [38] and on glass derivatized with glutaraldehyde [32], biotin on amine-reactive substrates [34, 35, 65, 66], and the adhesion peptide PA22-2 on a thiol-reactive surface [41]. In principle, crosslinkers can be attached to many types of substrates to bind proteins from stamps with high efficiency.

20 Microcontact Printing of Proteins

6.5 Printing

Handling a stamp with tweezers is the simplest approach to microcontact print proteins on surfaces both at low and high resolution. Typically, the stamp is brought close to the surface at an angle and set down gradually to ensure that conformal contact propagates from the initial contact areas without trapping air. Occasionally, (dust) particles or topography on the substrate are an obstacle to the propagating contact; applying gentle pressure to the stamp helps spread the contact to the rest of the substrate. Hybrid stamps are convenient to mount on printing tools [50] such as modified mask aligners, or on home-built printers using step motors to print substrates (up to 40 cm in lateral dimensions) with curved stamps while controlling the pressure applied to the stamp during printing [67]. Alignment of the stamp to preexisting structures on the substrate is desirable to print cell adhesion molecules on electrodes or to pattern substrates with multiple types of proteins, for example [39, 50]. Other methods already employed in soft lithography might be applicable for printing proteins. One example is printing surfaces and curved surfaces with cylindrical stamps [5, 10].

6.6 Characterization of the Printed Patterns

Surface-sensitive techniques such as ellipsometry, contact angle microscopy, and X-ray photoelectron spectroscopy can provide chemical information about surfaces printed with proteins. Patterns are best characterized using: (i) AFM, for which no labeling of the proteins is necessary; (ii) fluorescence and scanning confocal fluorescence microscopy; (iii) scanning electron microscopy, provided that the protein layer attenuates the emission of secondary electrons sufficiently to yield fair contrast [68]; or (iv) time-of-flight secondary ion mass spectroscopy [69]. AFM yields rich data concerning the contrast and resolution of the patterns, as well as the appearance and height of the printed layer, but suffers from the difficulty in localizing small printed areas [16]. Fluorescence microscopy is conversely effective in localizing signals from fluorescently labeled proteins, albeit with much less resolution than AFM. Colocalization of fluorophores having different spectral properties allows the detection of successively different types of proteins forming complex patterns. Optimization of the fluorescent signals is greatly facilitated by a` priori knowledge of the geometry of a printed pattern. Detection of unlabeled proteins is possible by immunoassays with detecting Abs either fluorescently labeled or conjugated with a reporter enzyme. In the latter case, the enzymatic conversion of chromogenic precursor into a precipitating fluorescent product keeps the fluorescent signal localized to the printed areas [17]. Staining using electroless deposition also keeps signals local [23]. Microcontact printing proteins onto diffraction gratings [51] or surfaces suited for plasmon resonance [26] offers the exciting capability of following binding events in real time and over sites in parallel.

7 Acknowledgments 21

7 Conclusion

The possibilities of microcontact printing biomolecules on a variety of surfaces with spatial control and resolution down to single protein molecules is unprecedented. Many fields could benefit in principle from these achievements. Surfaces could be decorated with high-quality patterns of microcontact-printed proteins for diagnostic applications. Very small volumes and quantities of reagents would be necessary for this purpose, using inking strategies based on microfluidic networks. In this case, numerous different types of protein could be patterned next to each other and used for surface ligand assays [70]. Printing proteins with tools that control the pressure during printing – hybrid stamps that are accurate and mechanically stable – and in alignment with predefined structures on the substrate has been demonstrated. The next steps are to build on these concepts, and mass fabricate high-quality arrays of proteins on surfaces such as glass slides or polystyrene surfaces for diagnostic applications. Stamps of various types can be devised. Some incorporate proteins in solution in a hydrogel-based reservoir, some have sites capable of binding target molecules from a complex ink, and others display numerous inked sites of micrometer lateral dimensions. Wherever biomolecules can be used on a surface, they might be placed advantageously by means of µCP. This is clearly the case for the growth of cells on surfaces, for which it becomes possible to construct hybrid architectures with cells connected to parts of electronic devices [53]. Positioning biomolecules on a biosensor surface is equally interesting. Microcontact printing can pattern areas of a sensing element with great precision and contrast, enabling the real-time monitoring of binding events on a surface. Possibly, the interaction between a large number of analytes and multiple printed sites could be screened for multianalyte immunoassays or drug screening. Microcontact printing is also well suited for preparing samples to investigate the biophysical properties of single biomolecules. Arrays of single biomolecules provide the advantage of having multiple sites to study each immobilized molecule with easy localization, without suffering from averaging effects, and with minimal signal degradation (photobleaching). It appears that although µCP was originally developed as a tool for applications in lithography [5], it is such a versatile technique that chemists and biologists have diverted it towards many more purposes. Microcontact printing clearly has unique, impressive features to manipulate and pattern biomolecules on surfaces, and it will be interesting to see how these will translate into firmly established applications for diagnostics, and biology in general.

Acknowledgments

I am very grateful to my colleagues Bruno Michel for his indefectible support of the work, to Andr´e Bernard and Jean Philippe Renault for having pioneered and carried out challenging experiments on microcontact printing proteins, and to Sergei Amontov, Helen Berney, Hans Biebuyck, Alexander Bietsch, Hans

22 Microcontact Printing of Proteins

Rudolf Bosshard, Isabelle Caelen, Dora Fitzli, Matthias Geissler, Bert Hecht, David Juncker, Max Kreiter, Heinz Schmid, Peter Sonderegger, Richard Stutz, Heiko Wolf, and Marc Wolf for their close collaboration on our biopatterning activities.

References 1 See for example the proceedings on µTAS: Micro Total Analysis Systems 2002: Y. BABA, S. SHOJI, A. van den BERG (eds), Kluwer Academic Publishers, Dordrecht, The Netherlands. 2 R. S. KANE, S. TAKAYAMA, E. OSTUNI, D. E. INGBER, G. M. WHITESIDES, Biomaterials 1999, 20, 2363–2376. 3 (a) D. R. REYES, D. IOSSIFIDIS, P.-A. AUROUX, A. MANZ, Anal. Chem. 2002, 74, 2623–2636; (b) P.-A. AUROUX, D. IOSSIFIDIS, D. R. REYES, A. MANZ, Anal. Chem. 2002, 74, 2637–2652. 4 A. KUMAR, H. A. BIEBUYCK, G. M. WHITESIDES, Langmuir 1994, 10, 1498–1511. 5 Y. XIA, G. M. WHITESIDES, Angew. Chem., Int. Ed. Engl. 1998, 37, 550–575. 6 A. BIETSCH, B. MICHEL, J. Appl. Phys. 2000, 88, 4310–4318. 7 S. J. CLARSON, J. A. SEMLYEN (eds), Siloxane Polymers, PTR Prentice Hall, Englewood Cliffs, NJ, 1993. 8 H. SCHMID, B. MICHEL, Macromolecules 2000, 33, 3042–3049. 9 T. W. ODOM, J. C. LOVE, D. B. WOLFE, K. E. PAUL, G. M. WHITESIDES, Langmuir 2002, 18, 5314–5320. 10 Y. XIA, D. QIN, G. M. WHITESIDES, Adv. Mater. 1996, 8, 1015–1017. 11 A. BERNARD, E. DELAMARCHE, H. SCHMID, B. MICHEL, H. R. BOSSHARD, H. A. BIEBUYCK, Langmuir 1998, 14, 2225–2229. 12 M. A. MARKOWITZ, D. C. TURNER, B. D. MARTIN, B. P. GABER, Appl. Biochem. Biotechnol. 1997, 68, 57–68. 13 A. BERNARD, D. FITZLI, P. SONDEREGGER, E. DELAMARCHE, B. MICHEL, H. R. BOSSHARD, H. A. BIEBUYCK, Nature Biotechnol. 2001, 19, 866–869. 14 B. D. MARTIN, B. P. GABER, C. H. PATTERSON, D. C. TURNER, Langmuir 1998, 14, 3971–3975.

15 B. D. MARTIN, S. L. BRANDOW, W. J. DRESSICK, T. L. SCHULL, Langmuir 2000, 16, 9944–9946. 16 J. P. RENAULT, A. BERNARD, A. BIETSCH, B. MICHEL, H. R. BOSSHARD, E. DELAMARCHE, M. KREITER, B. HECHT, U. P. WILD, J. Phys. Chem. B 2003, 107, 703–711. 17 A. BERNARD, J.-P. RENAULT, B. MICHEL, H. R. BOSSHARD, E. DELAMARCHE, Adv. Mater. 2000, 12, 1067–1070. 18 J. L. TAN, J. TIEN, C. S. CHEN, Langmuir 2002, 18, 519–523. 19 A. KUMAR, G. M. WHITESIDES, Appl. Phys. Lett. 1993, 63, 2002–2004. 20 H. HILLBORG, U. W. GEDDE, IEEE Trans. Dielectrics and Electrical Insulation 1999, 6, 703–717. 21 G. S. FERGUSON, M. K. CHAUDHURY, H. A. BIEBUYCK, G. M. WHITESIDES, Macromole-cules 1993, 26, 5870–5875. 22 C. DONZEL, M. GEISSLER, A. BERNARD, H. WOLF, B. MICHEL, J. HILBORN, E. DELAMARCHE, Adv. Mater. 2001, 13, 1164–1167. 23 J. P. RENAULT, A. BERNARD, D. JUNCKER, B. MICHEL, H. R. BOSSHARD, E. DELAMARCHE, Angew. Chem., Int. Ed. Engl. 2002, 41, 2320–2323. 24 B. R. YOUNG, W. G. PITT, S. L. COOPER, J. Colloid Interface Sci. 1988, 124, 28–43. 25 J. D. ANDRADE, V. HLADY, Adv. Polym. Sci. 1986, 79, 1–63. 26 H. B. LU, J. HOMOLA, C. T. CAMPBELL, G. G. NENNINGER, S. S. YEE, B. D. RATNER, Sensors and Actuators B 2001, 74, 91–99. 27 L. A. KUNG, L. KAM, J. S. HOVIS, S. G. BOXER, Langmuir 2000, 16, 6773–6776. 28 D. STAMOU, C. DUSCHL, E. DELAMARCHE, H. VOGEL, Angew. Chem., Int. Ed. 2003, 42, 5580–5583. 29 D. LOSIC, J. G. SHAPTER, J. J. GOODING, Langmuir 2001, 17, 3307–3316.

References 30 D. W. BRANCH, B. C. WHEELER, G. J. BREWER, D. E. LECKBAND, IEEE Trans. Biomed. Eng. 2000, 47, 290–300. 31 M. NISHIZAWA, K. TAKOH, T. MATSUE, Langmuir 2002, 18, 3645–3649. 32 D. W. BRANCH, J. M. COREY, J. A. WEYHENMEYER, G. J. BREWER, B. C. WHEELER, Med. Bio. Engineer. Comput. 1998, 36, 135–141 33 J. S. HOVIS, S. G. BOXER, Langmuir 2000, 16, 894–897. 34 Z. YANG, A. CHILKOTI, Adv. Mater. 2000, 12, 413–417. 35 J. LAHANN, I. S. CHOI, J. LEE, K. F. JENSEN, R. LANGER, Angew. Chem., Int. Ed. Engl. 2001, 40, 3166–3169. 36 A. T. A. JENKINS, N. BODEN, R. J. BUSHBY, S. D. EVANS, P. F. KNOWLES, R. E. MILES, S. D. OGIER, H. SCHO¨ NHERR, G. J. VANCSO, J. Am. Chem. Soc. 1999, 121, 5274–5280. 37 E. DELAMARCHE, H. A. BIEBUYCK, H. SCHMID, B. MICHEL, Adv. Mater. 1997, 9, 741–746. 38 C. D. JAMES, R. C. DAVIS, L. KAM, H. G. CRAIGHEAD, M. ISAACSON, J. N. TURNER, W. SHAIN, Langmuir 1998, 14, 741–744. 39 C. D. JAMES, R. D. DAVIS, M. MEYER, A. TURNER, S. TURNER, G. WITHERS, L. KAM, G. BANKER, H. CRAIGHEAD, M. ISAACSON, J. TURNER, W. SHAIN, IEEE Trans. Biomed. Eng. 2000, 47, 17–21. 40 L. KAM, W. SHAIN, J. N. TURNER, R. BIZIOS, Biomaterials 2001, 22, 1049–1054. 41 M. SCHOLL, C. SPRO¨ SSLER, M. DENYER, M. KRAUSE, K. NAKAJIMA, A. MAELICKE, W. KNOLL, A. OFFENHA¨ USSER, J. Neurosci. Methods 2000, 104, 65–75. 42 J. HYUN, H. MA, P. BANERJEE, J. COLE, K. GONSALVES, A. CHILKOTI, Langmuir 2002, 18, 2975–2979. 43 P. HARDER, M. GRUNZE, R. DAHINT, G. M. WHITESIDES, P. E. LAIBINIS, J. Phys. Chem. B 1998, 102, 426–436. 44 K. L. PRIME, G. M. WHITESIDES, J. Am. Chem. Soc. 1993, 115, 10714–10721. 45 S. R. SHETH, D. LECKBAND, Proc. Natl. Acad. Sci. USA 1997, 94, 8399–8404. 46 E. DELAMARCHE, C. DONZEL, F. KAMOUNAH, H. WOLF, M. GEISSLER, R. STUTZ, P. SCHMIDT-WINKEL, B. MICHEL, H. J. MATHIEU, K. SCHAUMBURG, Langmuir 2003, 19, 8749–8758.

47 B. MICHEL, A. BERNARD, A. BIETSCH, E. DELAMARCHE, M. GEISSLER, D. JUNCKER, H. KIND, J.-P. RENAULT, H. ROTHUIZEN, H. SCHMID, P. SCHMIDT-WINKEL, R. STUTZ, H. WOLF, IBM J. Res. Develop. 2001, 45, 697–719. 48 C. Y. HUI, A. JAGOTA, Y. Y. LIN, E. J. KRAMER, Langmuir 2002, 18, 1394–1407. 49 F. SCHWESINGER, R. ROS, T. STRUNZ, D. ANSELMETTI, H.-J. G¨UNTHERODT, A. HONEGGER, L. JERMUTUS, L. TIEFENAUER, A. PLU¨ CKTHUN, Proc. Natl. Acad. Sci. USA 2000, 97, 9972–9977. 50 L. LAUER, S. INGEBRANDT, K. SCHOLL, A. OFFENHA¨ USSER, IEEE Trans. Biomed. Eng. 2001, 48, 838–842. 51 P. M. St. JOHN, R. DAVIS, N. CADY, J. CZAJKA, C. A. BATT, H. G. CRAIGHEAD, Anal. Chem. 1998, 70, 1108–1111. 52 J. LAHANN, M. BALCELLS, T. RODON, J. LEE, I. S. CHOI, K. F. JENSEN, R. LANGER, Langmuir 2002, 18, 3632–3638. 53 A. OFFENHA¨ USSER, K. VOGT in C. M. NIEMEYER, C. A. MIRKIN (eds.): Nanobiotechnology, Chapter 5 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim 2004. 54 S. C. LIN, F. G. TSENG, H. M. HUANG, C. Y. HUANG, C. C. CHIENG, Fresenius J. Anal. Chem. 2001, 371, 202–208. 55 L. G. MENDOZA, P. MCQUARY, A. MONGAN, R. GANGADHARAN, S. BRIGNAC, M. EGGERS, BioTechniques 1999, 27, 778–788. 56 J. B. DELEHANTY, F. S. LIGLER, Anal. Chem. 2002, 74, 5681–5687. 57 H. D. IRENOWICZ, S. HOWELL, F. E. REGNIER, R. REIFENBERGER, Langmuir 2002, 18, 5263–5268. 58 J. TIEN, C. M. NELSON, C. S. CHEN, Proc. Natl. Acad. Sci. 2002, 99, 1758–1762. 59 M. GEISSLER, A. BERNARD, A. BIETSCH, H. SCHMID, B. MICHEL, E. DELAMARCHE, J. Am. Chem. Soc. 2000, 122, 6303–6304. 60 E. DELAMARCHE, A. BERNARD, H. SCHMID, B. MICHEL, H. A. BIEBUYCK, Science 1997, 276, 779–781. 61 D. JUNCKER, H. SCHMID, U. DRECHSLER, H. WOLF, M. WOLF, B. MICHEL, N. de ROOIJ, E. DELAMARCHE, Anal. Chem. 2002, 74, 6139–6144. 62 D. QIN, Y. XIA, G. M. WHITESIDES, Adv. Mater. 1996, 8, 917–919.

23

24 Microcontact Printing of Proteins

63 T. W. ODOM, V. R. THALLADI, J. C. LOVE, G. M. WHITESIDES, J. Am. Chem. Soc. 2002, 124, 12112–12113. 64 G. T. HERMANSON, A. K. MALLIA, P. K. SMITH (eds), Immobilized Affinity Ligand Techniques, Academic Press Inc., San Diego, CA, 1992. 65 Z. YANG, A. M. BELU, A. LIEBMANNVINSON, H. SUGG, A. CHILKOTI, Langmuir 2000, 16, 7482–7492. 66 J. HYUN, Y. ZHU, A. LIEBMANN-VINSON, Jr., T. P. BEEBE, A. CHILKOTI, Langmuir 2001, 17, 6358–6367.

67 E. DELAMARCHE, J. VICHICONTI, S. A. HALL, M. GEISSLER, W. GRAHAM, B. MICHEL, R. NUNES, Langmuir 2003, 19, 6567–6569. 68 G. P. LOPEZ, H. A. BIEBUYCK, R. HAERTER, A. KUMAR, G. M. WHITESIDES, J. Am. Chem. Soc. 1993, 115, 10774–10781. 69 A. M. BELU, Z. YANG, R. ASLAMI, A. CHILKOTI, Anal. Chem. 2001, 73, 143–150. 70 R. P. EKINS, H. BERGER, F. W. CHU, P. FINKCH, F. KRAUSE, Nanobiology 1998, 4, 197–220.

1

Bacteriorhodopsin and Its Potential in Technical Applications Norbert Hampp

Philipps-Universit¨at Marburg, Marburg, Germany

Dieter Oesterhelt Max-Planck Institute for Biochemistry, Planegg-Martinsried, Germany

Originally published in: Nanobiotechnology. Edited by Christof M. Niemeyer and Chad A. Mirkin. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30658-9

1 Introduction

In biotechnology, the production of biological macromolecules for technical processes is state-of-the art. The biosynthetic capabilities of cells go far beyond those of organic chemistry. Materials such as functional biomolecules, enzymes, antibodies, and hormones are indispensable in the food industry, cleaning, medical diagnosis, pharmacy, and therapy, and consequently many companies supplying systems such as DNA chips and readers, and high-throughput screening platforms were established to meet demands. The biotechnology industry is booming – indeed, it is very likely to become the most important high-tech industry within the next few decades. Why is “bio” the coming technology? The advantages and the need for enduring miniaturizations are an increasing challenge as long as conventional lithographic methods are employed. The utilization of self-organization principles and bioengineering of functional biological structures seems to be a promising alternative approach. Re-engineering of biomolecules in order to realize technically desirable functions has become possible. However, there remains another problem – namely, the communication between classical microsystems (in particular electronic systems) and the nanoscaled biomolecules. This interface remains the major challenge in the realization of “cross-technology” products. Last – but not least – there is the problem of stability, as biomaterials are less stable than organic and semiconductor structures. This is of course a problem

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Bacteriorhodopsin and Its Potential in Technical Applications

today, but in future technologies, where repair mechanisms like those in living organisms may be implemented, this obstacle may be overcome. Today, we have the need to seek methods for stabilizing the structures of biomaterials before they can be considered for technical applications. Bacteriorhodopsin has been studied over the past two decades as a material for technical applications. Its stability is adequate, it has several technically interesting functions, tools for both its modiﬁcation and production in technical quantities have been developed, and it offers various interface principles, whether optical, electrical, or chemical. In the following sections, the activities for technical applications of bacteriorhodopsin will be reviewed and future developments will be discussed.

2 Overview: The Molecular Properties of Bacteriorhodopsin

In the ﬁrst section of this chapter, the organisms in which bacteriorhodopsin is found are described, after which the processes used to modify this protein and to produce it in large quantities will be outlined. Finally, the current technical applications of bacteriorhodopsin will be summarized. 2.1 Haloarchaea and their Retinal Proteins

Archaea form, together with Bacteria and Eucarya, the three domains in life. Archaea are unicellular organisms thriving in a variety of habitats. Most of the archaea so far isolated and cultivated prefer extreme environments, although recent investigations have revealed archaea to be a standard component in biomasses of terrestrial and marine environments [1, 2]. Archaea are split into the crenarchaeotal and the euryarchaeotal branches. Members of the crenarchaeotal family comprise the full temperature range of life, from habitats at −1.8◦ C in the Antarctic to hyperthermal springs where Pyrolobus fumarii holds the world record in optimal temperatures for growth at 113◦ C [3]. Typical of the euryarchaeota are the methanogenic archaea. These occur ubiquitously in locations where organic matter is decomposed under strictly anaerobic conditions, such as in aquatic sediments, in marshes, or in the rumen of herbivores, and they contribute heavily to biogenic methane production. A second branch of the euroarchaeota comprises extreme halophilic organisms, the so-called halobacteria; these are ubiquitous on earth wherever salt occurs in solute concentrations close to saturation (Figure 1). Archaea and their proteins won commercial interest for the unusual physicochemical conditions under which they live and reproduce. The best example is bioleaching (biohydrometallurgy) of ore rubble, which was ﬁrst carried out during the 1950s for copper and is now used to produce a variety of metals [4]. Heatstable enzymes are valuable molecular biological tools; examples include heat-stable catalysts for the polymerase chain reaction, or use in washing powders. Further

2 Overview: The Molecular Properties of Bacteriorhodopsin 3

Fig. 1 Halobacterium salinarum is found in nature in concentrated salt solutions as they occur in salines. A purple color is caused by bacteriorhodopsin, and this is the key protein of the photosynthetic capabilities

of H. salinarum. The proton pathway with the amino acids involved and the lysinebound retinylidene residue are shown in the structural model of bacteriorhodopsin (foreground).

applications of archaeal proteins which are halophilic or especially resistant to extreme pH values might become apparent in the near future. A high potential for applications in nanotechnology lies in the fabrication of devices on the basis of archaeal surface-layer proteins [5]. These readily form two-dimensional crystals, and actually occur on the cell surface as natural crystals with pores of precisely deﬁned size. To date, six genera of halophilic archaea have been identiﬁed [6], and retinal proteins occur ubiquitously among these. In two genera, the extreme halophilicity is connected to a second extreme condition of life, alkalophilism, and these archaea are found in large masses in salty natron lakes. Halophilic archaea in general grow on organic substrates and have optimized their bioenergetics during evolution. With a sufﬁcient supply of organic nutrients and oxygen, they respire in standard fashion. However, it must be noted that oxygen solubility in saturated salt solution is about ﬁve times lower than in water. Under the anaerobic conditions that commonly occur in their habitats, these organisms have acquired three alternative means for energy conversion: either they use nitrate, dimethylsulfoxide or trimethylaminoxide as end electron acceptors [7, 8] or they ferment arginine to carbamoyl phosphate for the production of ATP [9–11]. The third choice is a very powerful system, and the second route to photosynthesis in nature. This system does not use chlorophyll-based reaction centers (as do bacteria and plants) but rather

4 Bacteriorhodopsin and Its Potential in Technical Applications

relies on retinal as a photon absorber and retinal-containing proteins as energy transducers [12]. The light-driven proton pump bacteriorhodopsin drives a proton circuit across the cell membrane, and ATP is produced via photophosphorylation as the chemical energy source for cell growth. The process is supported by the chloride pump halorhodopsin, which converts light energy via chloride transport into electrochemical energy used for the maintenance of osmotic balance. Halobacteria in nature grow very slowly, due to the limited supply of organic nutrients in hypersaline lakes. (The reader is referred to Refs. [7, 8] for a detailed description of their ecophysiology and the natural cycle of their “blooms” in salt lakes.) An example of such massive growth density is shown in Figure 1; an indication of heavy growth is the presence of an intense reddish to purple color. These blooms may occur only once each year (or even less frequently), but under optimization of nutrient supply in the laboratory generation times are usually in the order of hours. For biotechnological use, these two facts have to be considered: (i) a salt concentration of 25% is required to prepare peptone media, and this may be detrimental even to stainless steel fermentors; and (ii) peptone media are usually expensive to produce. These two obstacles are respectively overcome by using salt-resistant fermenting units and low-priced yeast extracts to replace the peptone. Another point to consider is the genetic stability of strains. Here, extensive progress has been made in the availability of stable standard strains used in biotechnology. In particular, strains which produce bacteriorhodopsin can be maintained stably by the use of phototrophic selection procedures [13]. Halophilic archaea, among them the genus Halobacteria, are unique in the sense that they are the only group of archaea that contains retinal proteins. While originally thought to occur only in higher animals capable of vision, retinal proteins recently have also been found in unicellular plant organisms [14], in fungi [15], in bacteria [16] and, most diverse in function, in halophilic archaea [12]. While all other groups seem to harbor either one of the two principal functions of retinal proteins – that is, a sensory function (eucaryotes) or a presumed energy-converting function (bacteria) – only halophilic archaea have developed a set of four retinal proteins, two of which serve a sensory function, while two convert light energy to chemical energy. Retinal, or vitamin-A aldehyde, originates from β-carotene by oxidative cleavage in the center of the molecule. The aldehyde in the free state is a chemically labile molecule with ﬁve conjugated double bonds. It is oxygen-sensitive and shows lightinduced isomerization around all double bonds. Light and oxygen together (photooxidation) destroy the free retinal easily. All known proteins containing retinal protect the molecule against photooxidation and select speciﬁc photoisomerization reactions, e. g., from 11-cis retinal to all-trans retinal in visual pigments and all-trans retinal to 13-cis retinal in the haloarchaeal proteins. All retinal proteins known are intrinsic membrane proteins and possess a transmembrane helical topography. Retinal always binds to the ε-amino group of a lysine residue of the seventh transmembrane helix, and a protonated Schiff base results; this becomes embedded in a cage of amino acids, which in turn drastically modiﬁes the spectroscopic, chemical, and photochemical properties. For example visual pigments cover the color range

2 Overview: The Molecular Properties of Bacteriorhodopsin 5

Fig. 2 Bioenergetics of Halobacterium salinarum. Bacteriorhodopsin acts as a light-driven, outward-directed proton pump. The generated proton gradient over the cell membrane drives a membrane-bound ATPase. These two proteins together form the simplest photosynthetic system known.

of the entire visible spectrum, and bacteriorhodopsin is chemically more stable than most proteins. Moreover, its light fastness is greater than that of organic dye molecules. Four molecular structures of retinal proteins are presently known: the visual pigment rhodopsin [17]; bacteriorhodopsin [18], halorhodopsin [19]; and sensory rhodopsin II [20]. Halobacterium salinarum, for example, makes extensive physiological use of retinal proteins (Figure 2). Bacteriorhodopsin drives the above-mentioned photosynthetic process for the production of ATP in light, and is also coupled to a highcapacity energy storage system in the form of a molar potassium gradient [21]. In order to maintain osmotic balance during growth – when a volume increase occurs under conditions of isoosmolar conditions, both inside and outside the cell – halorhodopsin is used as a light-driven anion pump [22]. This transports chloride ions into the cells, against the existing electrochemical potential, and allows a net salt accumulation to occur during the volume increase. This is a second way of avoiding the extensive use of respiratory energy at the expense of organic nutrients, by using light energy instead. Finally, two other retinal proteins occur in the cell which monitor the intensity and wavelength of the environmental illumination. These two photoreceptors – sensory rhodopsin I and II – receive orange light as an attractive stimulant, and blue light and near UV-light as repulsive stimulants (for a review, see Ref. [23]). Both photoreceptors signal to a two-component system of the cell, thereby regulating the frequency of the ﬂagellar motor’s changes in rotational sense for directing halobacterial cells into an environment of optimal conditions. These photoreceptors are only two among a total of 18 receptors, all of

6 Bacteriorhodopsin and Its Potential in Technical Applications

which convert chemical and physical signals of the environment to direct the cell into areas of optimal growth. The structures of all four halobacterial retinal proteins are very similar, and the molecular structures of two – bacteriorhodospin and halorhodopsin – have been elucidated to the atomic level. Moreover, based on details of the sequence alignment of the two ion pumps and the two sensors, together with the details of some two dozen other structures of archaeal retinal proteins, a tree has been created for this unique family of proteins [24]. Of particular interest in biophysical and biochemical terms is that the functions of the proton pump, the chloride pump, and the sensors are largely inter-convertible, either by varying the physical conditions under which the molecules operate, or by introducing minor genetic modiﬁcations. As an example, a point mutation in bacteriorhodopsin will convert this proton pump into a chloride pump [25, 26]. 2.2 Structure and Function of Bacteriorhodopsin

Bacteriorhodopsin is by far the best-studied archaeal retinal protein. It is naturally overproduced in the cells under conditions of illumination and limited aeration of a growing cell culture, but constitutive overproducers have been isolated. These cell lines produce up to 300 000 copies per cell, covering about 80 % of the cell surface as patches of two-dimensional natural crystals of bacteriorhodopsin with speciﬁc lipidic molecular species forming the so-called purple membrane [18]. Being a paradigm of light-driven proton pumps and seven-transmembrane helical proteins, the intense studies on this molecule over the past three decades by dozens of laboratories have produced a large body of knowledge on its biophysics, biochemistry, and molecular biology. Thus, the biotechnology of this “myoglobin” of membrane proteins is well founded on detailed knowledge about this molecule (for a recent review, see Ref. [27]). Like the other archaeal retinal proteins, bacteriorhodopsin is an intrinsic membrane protein with the common seven-transmembrane helix topology (see Figure 1) and an approximate molecular weight of 26 kDa. The seven helices are arranged in two arcs: an inner arc with helices B, C, and D; and an outer arc with helices E, F, G, and A. A transmembrane pore is formed mainly between helices B, C, F, and G. The retinal is bound to Lys216 in helix G as a protonated Schiff base, which interrupts the pore and separates an extracellular (EC) half channel (Figure 1, downward oriented) from a cytoplasmic (CP) half channel (Figure 1, upward oriented). The retinal side chain in the binding pocket is closely packed between four tryptophan residues, and the positively charged Schiff base interacts electrostatically with the protein environment. The chromophore is deﬁned as the retinylidene moiety and the side chains in contact with it. Its color is tuned by electrostatic interactions speciﬁcally with the Schiff base and a complex counterion consisting of several amino acids. Light absorption causes photoisomerization of the retinal (all-trans to 13-cis and vice versa) and energy storage which includes ion-afﬁnity shifts (H+

2 Overview: The Molecular Properties of Bacteriorhodopsin 7

Fig. 3 Photocycle of bacteriorhodopsin. (A) Upon absorption of a photon, the initial $-state of bacteriorhodopsin is converted photochemically to the J-state from where a series of thermal steps leads back to the initial state. The proton transport is intimately coupled to the photocycle, which is observed as a sequence of intermediates which are represented by the common single-letter code with their absorption maxima given as subscripts. In the dark, bacteriorhodopsin relaxes thermally to the D-

state which has 13-cis conﬁguration. The resulting mixture of $- and D-states is called dark-adapted bacteriorhodopsin. From the O-state, a photochemical conversion of alltrans to 9-cis retinal is possible which is not thermally reisomerized to the initial state. (B) The proton transport and related retinal conﬁgurations as well as accessibility of the nitrogen in the Schiff-base linkage between retinal and Lys216 are indicated. This sequence represents several of the molecular changes involved in the proton transport.

in bacteriorhodopsin and Cl− in halorhodopsin) of the Schiff base and internal binding sites as well as conformational changes as central elements of the catalytic cycle. This cycle may formally be represented as a sequence of six steps which are indispensable for transport: the all-trans to 13-cis photoisomerization and its thermal reversal (isomerization, I); a reversible change in accessibility (switch, S) of the Schiff base for ions in the EC and CP channel respectively; and ion transfer (T) reactions to and from the Schiff base active center (Figure 3). Speciﬁcally in

8 Bacteriorhodopsin and Its Potential in Technical Applications

wild-type bacteriorhodopsin the order of these elementary steps is photoisomerization, proton transfer from the Schiff base to the acceptor aspartic acid (D) 85 in the EC channel, change in accessibility of the Schiff base from EC to CP, proton transfer from the donor aspartic acid (D) 96 in the CP channel to the Schiff base and thermal reisomerization, followed by reset of the Schiff base accessibility from CP to EC. The two proton transfer steps are intimately linked to cooperative changes in the EC and CP channels where hydrogen networks exist [28], and proton conduction over the total distance of about 48 Å is very likely based on a Grotthuss-type mechanism [29]. Structural key players in the EC channel are, besides D85, the arginine residue 82 and the two glutamic acid residues 194 and 204. They are connected by a water bridge to form a dyad which, by all likelihood, is the proton-release unit on the EC surface [18]. Other water molecules in the EC channel also play an important role [30, 31] in addition to amino acid side chains. One water molecule is located between D85 and the Schiff base, and disappears in the structure of the key intermediate M [32]. Three more water molecules below D85 are interconnected with side chains to form an extended hydrogen network as suggested by early experiments [33]. The CP channel from which the Schiff base later in the catalytic cycle receives the proton back is functionally dominated by the protonated D96. The distance between D96 and the nitrogen is 15 Å, and the hydrophobic nature of the space between them could provide an insulating layer against the membrane potential of 280 mV in the ground state. One water molecule occurs in contact with D96, and one water molecule is located between D96 and the Schiff base. Again, changes of this part of the molecule are observed in the key intermediate M [32]. At the proton entrance of the CP channel, aspartic acid D38 plays a role in the refeeding mechanism of protons [34]. The most dramatic effect of mutations on the structure and function of bacteriorhodopsin have the two carboxylates at positions 85 and 96. The removal of the proton acceptor D85 prevents the deprotonation of the Schiff base and almost completely annihilates the proton transport function (for a detailed description, see Ref. [35]). The lack of the proton donor D96 slows the catalytic cycle by a factor of up to several hundred, depending on temperature, humidity, azide concentration, etc. [36]. The reason is that the lack of the internal proton donor renders the rate of reprotonation of the Schiff base depending on external pH. Another important feature of the M-intermediate is its capacity to absorb blue light, and by this to reconvert photochemically into the initial state. Thus, in mutants lacking D96 the life-time of the M-intermediate and therefore the speed of color changes can be regulated either by physico-chemical parameters such as pH, temperature, and humidity, or alternatively by simultaneous application of two photons of different quality, for example green and blue photons. The three most important features, which eventually lead to biotechnological use of bacteriorhodopsin on the basis of its catalytic cycle are: 1. The color changes, which can be used for any type of information processing and storage process.

2 Overview: The Molecular Properties of Bacteriorhodopsin 9

2. The photoelectric events which are due to the changing geometry of the Schiff base upon photoisomerization and the movement of the proton. Such electric changes occur from the picosecond to the millisecond time regime. 3. The pH change between the inside and the outside of bacteriorhodopsincontaining membrane systems as the net result of proton translocation.

So far, the color changes are under most intensive investigation because of the velocity of light-triggered reactions in bacteriorhodopsin and the possibility of regulating the speed of the color changes over a very wide range. 2.3 Genetic Modiﬁcation of Bacteriorhodopsin

The molecular biology and genetics of halophilic archaea are developing at a slow, but constant, pace. The development of a transformation system was a major breakthrough, as this opened the door to site-speciﬁc mutagenesis and gene deletion or replacement [37, 38]. The basic process consists of a protoplast (spheroblast) preparation by incubation of cells with EDTA to remove magnesium ions. These are necessary for the integrity of the surface layer formed by the glycoprotein in the cell wall of archaea. Spheroblasts are incubated with a mixture of vector DNA carrying an antibiotic resistance and polyethyleneglycol. After a curing period in complex medium the cells are plated with antibiotic to select transformed clones [39]. Not many antibiotics have been reported as efﬁcient agents against halophiles, and only two resistance genes have been cloned and inserted into suitable transformation vectors. Mevinolin inhibits the β-hydroxymethylglutaryl CoA (HMG) reductase and thus prevents isoprenoid synthesis of archaea. As lipids of halophilic archaea are diphytanoyl ethers of substituted glycerol, mevinolin completely blocks growth [40]. Novobiocin prevents DNA replication by inhibiting Gyrase B. The two selection marker genes used are hmg and gyrB; these were isolated from Haloferax volcanii and should have highly homologous genes in most other halophilic archaea [41]. The creation of site-speciﬁc retinal protein mutants requires appropriate host strains and vectors. The very stable strain H. (halobium) salinarum L33 [42] was introduced as a host for bacteriorhodopsin muteins [39]. It was found as a spontaneous mutation, carries an insertion element in the bop gene, and does not revert to a wild-type phenotype under any condition tested. Further strains were selected after either spontaneous or induced mutation which lack one or more of the retinal proteins, and many more strains with various phenotypes bearing favorable properties for investigations on the bioenergetics and signal transduction of halophiles have been reported. These are not covered in this chapter. More recently, systematic deletions of retinal protein genes have been described; one such example is the strain SNOB (S9 without bop) which is derived from the bacteriorhodopsinoverproducing strain S9 and has the bop gene deleted [43]. In consecutive rounds of a deletion procedure which uses the same antibiotic for selection repeatedly, all four retinal protein genes have been deleted to reach the strain NAOMI (now all

10 Bacteriorhodopsin and Its Potential in Technical Applications

opsins missing; M. Otsuka and D. Oesterhelt, unpublished results). This strain has the advantage that complementation with opsin genes allows the combination of their physiological function at will. Vectors have been introduced ﬁrst on the basis of a replicative element and a mevinolin-resistant determinant from a halophilic cell, together with a replicon and an ampicillin resistance from E. coli to serve as shuttle vectors [44]. In many laboratories, a plethora of vectors was subsequently designed with a size smaller than 10 kb, and without halobacterial origin of replication (suicide vectors) to enforce homologous recombination and therefore stable integration into the halobacterial genome. For interruption of a functional bop gene, the resistance gene used for selection is usually inserted into the coding sequence and remains stably associated with it permanently. Site-speciﬁc mutated genes can be favorably introduced into wildtype (S9) or mutated (L33) background in the following way. The resistance gene is placed next to the mutant bop gene on the suicide vector DNA. After transformation, antibiotic-resistant clones are selected which must result from a single crossover genetic event leading to the integration of vector DNA into the chromosome. These clones are allowed to grow without selection pressure, and are then replica plated with and without antibiotic. Among several genetic results is one which removes the vector DNA with the resistance gene and the original version of bop gene and leaves the mutated bop gene in the correct place stably integrated. These clones are found at various frequencies (usually one among several hundred) as the phenotype which does not grow on the antibiotic-containing plate but on its antibiotic-free counterpart. Besides genetic stability, the procedure provides the additional advantage of repeated use of the same antibiotic in further rounds of genetic alterations. Mutations of bacteriorhodopsin have been produced by the hundred and, once published, are available from the various laboratories. While the genetic background into which the mutated bop genes are introduced is often different, the mutagenesis procedure presently is usually the overlapping PCR method which replaced the preceding approach of gapped duplex DNA and is easily and quickly applied. Recently, a β-galactosidase gene as a reporter gene has been shown to act as an indicator gene for promoter strength and for blue-white selection procedures [45–47]. In conclusion, the production of bacteriorhodopsin muteins has become a routine method, but the maintenance of stable strains overproducing these muteins requires much care. In addition, the maximal level of mutein production can be very different, is unpredictable, and whether a given mutein will produce the crystalline arrangement of the purple membrane is also in doubt. 2.4 Biotechnological Production of Bacteriorhodopsins

The biotechnology of bacteriorhodopsin is based on simplicity of its isolation and chemical and photochemical stability when in the form of purple membranes. As

3 Overview: Technical Applications of Bacteriorhodopsin 11

mentioned, bacteriorhodopsin forms 2D crystals in vivo, and these can be isolated as purple membranes. The isolation is facilitated by two facts: (i) halobacterial cells are unstable in water and cell constituents are released upon lysis; (ii) the cell membrane is fragmented, and for unknown reasons the crystalline patches of the purple membrane are set free as fragments of largest size and highest buoyant density. These two speciﬁc features are used in the isolation procedure, either in a combination of sedimentation and isopycnic gradient centrifugation leading to a product of highest purity, or by a ﬁltration procedure. The puriﬁcation step yields a product which is 95–100% pure depending on the conditions and the mutein under consideration. Although these methods have not yet been exposed to economic competition, it is expected that the biotechnological production of purple membranes might in the future represent a competitive biomaterial in information technology. Certain limits of this material should be mentioned, however. As a membrane protein, bacteriorhodopsin cannot be produced via inclusion bodies as although the refolding and reconstitution of the active chromoprotein into membranes is possible in principle, it is certainly not feasible on an economic basis. Halobacterial cells do not form invaginations of their cell membrane like phototrophic bacteria (e. g., Rhodopseudomonas), which concentrate their reaction centers for photosynthesis in units called chromatophores. Although the purple membrane may occupy about 80 % of the cell membrane area, the amount of purple membranes obtained from cells is comparably low. It is needless to point out that the secretion of the integral membrane protein bacteriorhodopsin from cells into the medium has no biochemical basis. In practical terms, the current production of bacteriorhodopsin is at a level of 25 g m−3 nutrient broth. A 25-g quantity of bacteriorhodopsin would produce 50 L of suspension with an optical density of 1 (at a layer thickness of 1 cm this would allow the passage of only 10 % of light; i.e. it is an intense color). The molecular biology or production of bacteriorhodopsin variants has been established over the years, and at present the genetics is well known, including the sequence of the entire genome [48]. One problem in the molecular genetics of the halophilic archaea, especially H. salinarum, is that of insertion elements, as this sometimes causes instability of strains. However, with increasing knowledge on transformation procedures, vector construction and the creation of new antibiotics for the cells, these problems may be removed, at least for industrial purposes. It should be mentioned again, however, that cells producing bacteriorhodopsin variants do not always overproduce bacteriorhodopsin as do wild-type cells.

3 Overview: Technical Applications of Bacteriorhodopsin

The remarkable physico-chemical properties of bacteriorhodopsin and its numerous technically attractive molecular functions have led to many potential technical uses for this material [49–51]. In most of these applications, bacteriorhodopsin is used in the purple membrane (PM) form because its deliberation from the

12 Bacteriorhodopsin and Its Potential in Technical Applications

crystalline package reduces both the chemical and thermodynamic stability of bacteriorhodopsin to a signiﬁcant degree. Bacteriorhodopsin comprises three basic molecular functions which may be used in technical applications: photoelectric, photochromic, and proton transport properties. The molecular mechanisms responsible for each of these properties have been already discussed, and here we will focus brieﬂy on the related applications.

3.1 Photoelectric Applications

The use of bacteriorhodopsin as a molecular level photoelectric conversion element is one of the ﬁelds where technical applications of the material have been examined. Upon illumination, a photovoltage up to 250 mV per single bacteriorhodopsin layer is generated, and this may be used either as an indicator or control element for various applications. Triggered by the absorption of a photon, the bacteriorhodopsin molecule undergoes a series of very rapid molecular changes, one of which is the generation of a photovoltage caused by changes in the orientation of molecular dipole moments that are triggered by the isomerization of the retinal on the femtosecond scale (Figure 4A). The proton released through the outer proton half channel may be either transferred to the outer medium or conducted along the surface of the PM [52]. The proton conductivity along the PM in H. salinarum cells supports delocalization of protons on the surface and proton conduction to the membrane-bound ATPase molecules. In most of the photoelectric applications of PM, the water content of the ﬁlms is reduced, and this in turn causes proton conduction along the membrane surface. A single PM sheet contains several thousands of unidirectionally oriented BR molecules. Upon illumination, a number of bacteriorhodopsin molecules proportional to the intensity of light will be excited to accomplish a proton transport process. As all of the bacteriorhodopsin molecules in a single PM patch are oriented in the same direction, the voltage generated over a single membrane is independent of the number of active molecules (Figure 4B), but the proton current generated is proportional to the light intensity. The photovoltage generated can be easily measured by embedding the PM layer between two transparent electrodes. The light-triggered photovoltage induces a compensating polarization voltage in the outer electrodes, and this in turn may be detected as a high impedance voltage signal (Figure 4C). In a perfectly capacitive coupled system of this type, an induced photovoltage with the characteristics shown in Figure 4C is observed. A voltage is induced only during the light intensity change. The polarity of the signal is different for the OFF =⇒ ON and the ON =⇒ OFF transition. This is called the “differential responsivity” of PM layers. There are two major issues to be solved for an attractive use of bacteriorhodopsin in photoelectric processing. The ﬁrst is that a high degree of orientation of the PMs is obtained because counter-oriented PMs cancel out each others’ photoelectric effects. The second is that a coupling of the light-dependent proton-motive force

3 Overview: Technical Applications of Bacteriorhodopsin 13

Fig. 4 Photoelectric properties of bacteriorhodopsin. (A) Absorption of a photon by bacteriorhodopsin causes molecular changes which lead on a pico- to nanosecond timescale to the generation of a photovoltage over the molecule. The bacteriorhodopsin molecules in a single purple membrane (PM) patch are unidirectionally oriented. The light-driven vectorial proton transport of all the bacteriorhodopsin molecules is switched in parallel. This means that, over a single PM patch, it is not the voltage but the proton current which is proportional to the light intensity. The protons transported through the bacteriorhodopsin molecules can either be released to the outer medium or move along the surface of the PM patch due to proton conduction. (B) Oriented bacteriorhodopsin molecules, each represented by an arrow, sandwiched in a capacitor structure can be used as a photoelectric indicator cell. Upon illumination, a charge separation over the bacteriorhodopsin layer is generated which

induces a proportional charging of the outer electrode layers. No electric conduction between the bacteriorhodopsin layer and the electrodes is required. The electric ﬁeld of the charge distribution induced in the electrodes compensates the electric ﬁeld caused by bacteriorhodopsin. (C) Depending on the type of outer circuitry, either the induced voltage or the induced charge motion can be measured. In the latter case, a signal is recorded which corresponds to the ﬁrst derivative of the temporal change of the light. (D) In a pixelated structure which is coated with oriented bacteriorhodopsin, photovoltages are measured only in those spots where a change in the light intensity occurs; hence, this is called novelty ﬁltering. (E) In a volume (e. g., in 3-D data storage) the detection of the photovoltage in an outer capacitor structure was considered for readout. If the bacteriorhodopsin in the point of excitation is in the $-state, the absorption of a photon will lead to a photoinduced voltage, but not in the M-state.

14 Bacteriorhodopsin and Its Potential in Technical Applications

of bacteriorhodopsin to the electromotive forces used in conventional electronics needs to be achieved. 3.1.1 Preparation of Oriented PM Layers The photoelectric signal of PM is high enough so that a single layer only of PM is needed for applications where the PM acts as a photoelectric indicator molecule. The Langmuir-Blodgett technique is often used for the preparation of such devices. The main problem is to obtain a high degree of orientation of the PM patches. The PM patches are thin (5 nm), large (up to micrometers), and ﬂexible, and for this reason a mechanical orientation is difﬁcult to obtain. The physical properties of the two sides of the PM are deﬁnitively different, but the differences are not so large that orientation (e. g., in an electric ﬁeld) achieves more than a preorientation. Any preorientation obtained needs to be made permanent, for example, by covalent crosslinking or by polymer embedding of the PM patches. The only reliable method described so far is the coating of a surface with monoclonal antibodies against bacteriorhodopsin [53]. Since the antibodies selectively react with the cytoplasmic or extracellular side of the PM, a highly oriented monolayer of PM can be obtained using this method, although unfortunately it is restricted to a single PM layer. 3.1.2 Interfacing the Proton-Motive Force The other problem in photoelectric applications is that under light exposure bacteriorhodopsin transports protons, but not electrons. The interface between the biological component bacteriorhodopsin and its proton-motive force and the electromotive forces required for conventional electronics needs to be considered in the design of devices. Interfacing the proton-motive force of BR with the electron-conducting outer electrodes requires an electrolyte layer which couples both “worlds”. The balancing between the electronic circuitry and the photoelectric properties of a PM layer is crucial, and is the reason why results from different laboratories are so incomparable. The more than complete review published by Hong [54] is recommended for the reader who seeks much more detail on this subject. 3.1.3 Application Examples Ultrafast photodetection was one of the earliest proposals for the use of bacteriorhodopsin in a technical application [55, 56]. Two-dimensional photoelectric arrays, as artiﬁcial retinas or as control elements for a liquid crystal spatial light modulator, were developed later. Last, but not least, the photoelectric properties of bacteriorhodopsin can be used for indicator purposes in 3-D memories. 3.1.3.1 Artiﬁcial retinas Most devices which make use of the photoelectric properties of bacteriorhodopsin are called “artiﬁcial retinas” [57–59], the reason being that they offer certain preprocessing features known from the retina, including edge detection and novelty ﬁltering. The physical background is called the “differential response” of PM layers. A device (see Figure 4D) comprising an electrode array which is covered with one or more oriented layers of PM may be used for

3 Overview: Technical Applications of Bacteriorhodopsin 15

novelty ﬁltering. Each of the electrodes is connected to an ampliﬁer electronics. First, assume that the dark rectangular structure shown in Figure 4D prevents a set of electrodes from being exposed to light. Then, if this structure is moved over the light-sensitive sensor area (see arrow), it causes a voltage to be induced in each of the pixel electrodes, with a sign proportional to the light change. Because only those electrodes “ﬁre” where a change of the illumination occurs, this type of sensor is called an “artiﬁcial retina”, and this type of preprocessing is called “novelty ﬁltering”. Due to the differential response of bacteriorhodopsin, the polarity of the electrode signal carries the information whether a pixel was switched to “ON” or to “OFF”, and in turn the direction of the movement of the object can be derived. 3.1.3.2 Electro-optically controlled spatial light modulators Another use of the artiﬁcial retina described above may be in optically addressed spatial light modulators (SLMs). The ampliﬁer electronics which detects the photovoltages induced in the electrode pixels may be connected to a SLM device. In particular, liquid crystal (LC)-based SLMs are state-of-the-art. In this case, the electrodes of the bacteriorhodopsin-based artiﬁcial retina are connected one-to-one to the pixels of a LC-SLM. An advanced version omits the wiring and ampliﬁers. The LC-layer of the SLM is controlled directly by the artiﬁcial retina device [60, 61]. 3.1.3.3 Readout in 3-D Memories Another application of the photoelectric properties of PM was investigated for the readout of volume storage units with bacteriorhodopsin. The basis is a cube of oriented PM patches in either the purple initial state or the yellow M state. Upon illumination of a PM patch in the initial state (which may be addressed by actinic light), a photovoltage signal is induced. A PM patch in the M state would not respond to the actinic light, but two electrodes on the outer surfaces of the bacteriorhodopsin cube could detect the photovoltage generated, and by this method the 3-D distribution of the photochemical states of PM patches could be read out. This is a prerequisite for a 3-D memory based on bacteriorhodopsin (Figure 4E). 3.2 Photochromic Applications

Most applications currently investigated utilize the photochromic properties of bacteriorhodopsin [50, 62]. During the photocycle, bacteriorhodopsin cycles through a pair of spectroscopically distinguishable intermediates (see Figure 3A), all of which have an absorption maximum which is different from that of the initial B state. However, in most applications the photochromism of bacteriorhodopsin is used in connection with the purple to yellow absorption change which is related to M state formation. Upon acidiﬁcation, a blue membrane is formed which has a signiﬁcantly different photocycle. The formation of a 9-cis retinal-containing state is observed, and this is thermally stable. In contrast, 13-cis retinal is isomerized by the bacteriorhodopsin molecule to all-trans retinal at room temperature, and the isomerization of 9-cis

16 Bacteriorhodopsin and Its Potential in Technical Applications

Fig. 5 Photochromism of bacteriorhodopsin and its application. (A) The retinylidene residues are strongly anisotropic. (B) Upon illumination with polarized light, the retinylidene residues which are in parallel to the electric ﬁeld vector of the actinic light are preferentially excited and isomerized. (C)

Transient photochromic change of bacteriorhodopsin between the initial purple state and the yellowish M-state (middle). (D) The photochemical formation of 9-cis retinal may be utilized for photochromic long-term storage. (E) Permanent storage of information in bacteriorhodopsin.

retinal is not catalyzed. This pathway opens the route to long-term storage materials based on bacteriorhodopsin. 3.2.1 Photochromic Properties of Bacteriorhodopsin Isomerization from all-trans to 13-cis is the ﬁrst occurrence after the photochemical excitation of bacteriorhodopsin, and this causes signiﬁcant transient shifts in the absorption spectrum. In addition to the isomerization change, deprotonation of the chromophoric group is observed. In the L to M transition, a proton from the Schiff base nitrogen group is transferred to Asp85, and this deprotonation causes a drastic blue shift of the absorption to 410 nm. The photochromism of bacteriorhodopsin is dominated by the intermediate which has the longest life-time; hence, this forms a “bottleneck” in the photocycle. The retinylidene residue inside bacteriorhodopsin which forms part of the photochromic group in the molecule is strongly anisotropic. A PM layer may be considered as a crystalline arrangement of chromophoric groups which are oriented angles of 120◦ between them (Figure 5A). However, in most applications where a statistic number of PM patches is used, the angular chromophore distribution appears anisotropic. Due to the anisotropy of the retinylidene groups, excitation of a random distribution of PM patches with polarized light causes the chromophores to become oriented in parallel to the actinic light polarization such that a

3 Overview: Technical Applications of Bacteriorhodopsin 17

preferentially converted (and in turn photoinduced) anisotropy is obtained (Figure 5B). In solutions containing PM this is masked by diffusion, but in bacteriorhodopsin-ﬁlms where the PM patches are ﬁxed, the photo-induced anistropy can be easily observed and utilized. Today, three types of photochromic changes in bacteriorhodopsin have been described which enable different applications. The ﬁrst is the photochromic shift between the B and M states (Figure 5C), and this is used mainly for optical processing tasks. The second is photoerasable data storage using 9-cis-containing states of blue membrane or suitably modiﬁed BR-variants (Figure 5D). However, the very low quantum efﬁciency for recording of far below 1 % is a major limitation. And last, but not least, permanent photochromic changes obtained through two-photon absorption in bacteriorhodopsin are suitable for long-term data storage (Figure 5E). 3.2.2 Preparation of Bacteriorhodopsin Films Optical ﬁlms are prepared from bacteriorhodopsin by polymer embedding. Optically clear, water-soluble polymers are suitable for this purpose (e. g., polyvinylalcohol, gelatin). The ﬁlm formation is usually carried out by mixing the polymers with PMs and additives in aqueous solution, this being cast on a glass support. The water is generally removed by drying in air, but the ﬁlms may also be sealed with a second glass plate. 3.2.3 Interfacing the Photochromic Changes The reason why photochromic applications of bacteriorhodopsin are more developed than others is because of the ease with which an interface can be implemented between the bacteriorhodopsin ﬁlms and any type of optical system. The bacteriorhodopsin ﬁlm is completely sealed, the only interface being the light which transports energy and information simultaneously. 3.2.4 Application Examples Many applications based on the photochromism of bacteriorhodopsin have been suggested, and some are described brieﬂy here as an indication of the wide range of potential uses. 3.2.4.1 Photochromic color classiﬁer As the different rhodopsins in the eye enable color perception, the use of bacteriorhodopsins with different absorption maxima would allow a biomimetic system for color perception to be set up. The photoelectric response of three sensor elements coated with three different bacteriorhodopsin types (i. e., wild-type and two containing retinal analogues) are combined and coupled to a simple neural network for color recognition properties. This functions quite reliably, though it is unclear whether it has any technical advantages over conventional color sensors. Two limitations can be identiﬁed in this system. First, the absorption maxima of the three BR types used today are relatively similar, and they do not span the visual wavelength range as well as the human rhodopsins, notably in the blue region. Second, the conventional systems which typically comprise three different color ﬁlters and semiconductor light-sensitive elements reliably

18 Bacteriorhodopsin and Its Potential in Technical Applications

supply the required information. At present, it is difﬁcult to identify any advantages of the bacteriorhodopsin-based systems over conventional systems [63]. 3.2.4.2 Photochromic inks Another development is the preparation of photochromic inks. These inks differ from the polymer ﬁlms for optical recording by their rheological properties. Depending on the method of application (e. g., screen printing, offset printing), the viscosity and surface tension must be considered. The basic photochromic properties are quite similar to those of the optical ﬁlms, but auxiliaries in the compositions adjust the required application dependent properties. A major problem is to identify suitable compositions which do not interfere with the photochemical properties of the bacteriorhodopsin embedded [64]. 3.2.4.3 Electrochromic inks The main color shift in bacteriorhodopsin is due not to a primary photochemical reaction but to the protonation change of speciﬁc groups, in particular the Schiff base linkage and the Asp85 residue. As protons are charged particles, their removal from the binding position by electric ﬁelds should be possible. Indeed, this can be demonstrated, though the speed and efﬁciency of the decoloration/coloration process is quite low. Nonetheless, the basic principle was successfully demonstrated [65], indicating a potential development of electrochromic paper using bacteriorhodopsin. 3.2.4.4 Photochromic photographic ﬁlm It is well known that a permanent bleaching of bacteriorhodopsin may be achieved with hydroxylamine. The chemical reaction of hydroxylamine with the retinal binding site occurs in an intermediate state only, and no reaction of BR in the β-state is observed. Due to this ﬁnding, which dates back to the early retinal extraction experiments, it is possible to fabricate nonreversible optical ﬁlms from bacteriorhodopsin [66]. These ﬁlms behave quite similarly to photographic ﬁlms, except that the compounds needed for the chemical development are already contained in the ﬁlm. The reason why this process has not been used technically is that, after image formation, the nonreacted hydroxylamine must be removed from the ﬁlm, or fading of the contrast will occur when stored in light. 3.2.4.5 Long-term photorewriteable storage of information Much interest has centered on photorewriteable optical storage with bacteriorhodopsin. For this purpose, the conventional all-trans to 13-cis conversion is not suitable because of the retinal reisomerization at room temperature in bacteriorhodopsin. In blue membrane, a photochemical conversion from all-trans to 9-cis retinal, which appears pink in bacteriorhodopsin, may be induced by high light intensities. 9-cis retinal is thermally stable and requires photochemical excitation for reconversion of all-trans. The main disadvantage of blue membrane is that its formation from PM requires either acidiﬁcation or the removal of divalent cations, as both cause destabilization of the bacteriorhodopsin molecule. Aggregation of the PM patches is also seen. The requirement is for bacteriorhodopsin variants which have a similar photochemistry but at ambient pH value and without removal of cations. BR variants with alterations

3 Overview: Technical Applications of Bacteriorhodopsin 19

in position 85 show such desired properties, and one of the ﬁrst reported for this purpose was D85N. Another approach is to use wild-type BR at neutral pH values and to switch the material photochemically to the 9-cis state from the so-called O-state [67]. This scheme was named branched photocycle memory [68]. 3.2.4.6 Neural networks Due to the fact that its absorption state may be shifted with blue and yellow light in different directions, bacteriorhodopsin is also a suitable material for neural networks. The output from an absorptive bacteriorhodopsin-cell is used to control the absorption state of another such cell. This has been demonstrated in principle [69], but the bacteriorhodopsin has a “fan-out” of much less than one and for this reason is not really a suitable material for the implementation of optical neural networks. (“Fan-out” is a term from electronics which characterizes the signal power output of a device compared to the required signal power input the same device requires.) A “fan-out” of 10 means that the output terminal of the device supplies enough power that 10 input terminals of identical devices could be supplied with enough energy to signal them the input state reliably. In photochromic devices where no ampliﬁcation occurs, the “fan-out” is generally less than 1. Without external ampliﬁcation, it is almost impossible to set up control loops. 3.2.4.7 3-D information storage The use of bacteriorhodopsin in 3-D information storage has been investigated for some time. Bacteriorhodopsin shows an astonishingly high two-photon absorption cross-section of the initial B state. This is used in a two-photon absorption setup to address the absorption state of bacteriorhodopsin in three dimensions [70]. The recording process is quite well handled, but the readout is an intrinsic problem. The advantage of such a memory device is its tolerance towards electromagnetic radiation. 3.2.4.8 Nonlinear optical ﬁltering The strongly nonlinear optical response of bacteriorhodopsin towards the incident light intensity has given rise to many applications which use the nonlinear response for image processing purposes (e. g., edge enhancement, noise reduction). The response curve of the bacteriorhodopsin can be tuned over several orders of magnitude by changing the lifetime of the M-state. This may be accomplished by changing the pH, as well as using modiﬁed bacteriorhodopsins. 3.2.4.9 Holographic pattern recognition and interferometry The use of bacteriorhodopsin as a short-term memory has been tested in several applications. The most challenging was the construction of a real-time holographic pattern recognition system which operates at video frame rate. In this system, the holographic comparison of images allows similarities between objects used for identiﬁcation purposes for complex images to be quantiﬁed [71, 72]. In the bacteriorhodopsinﬁlm, holograms are recorded, read-out and erased at video frame rate. In the early 1990s, when the system was ﬁrst developed, it was unchallenged by computer systems, but as computing power has advanced the bacteriorhodopsin-ﬁlm method has been abandoned.

20 Bacteriorhodopsin and Its Potential in Technical Applications

Typical photochromic applications include a holographic real-time correlator (Figure 6A) and a holographic camera for nondestructive testing (Figure 6B). Examples of photochromic inks made from bacteriorhodopsin are shown in Figure 6C, with the initial colored (purple) and bleached, yellowish inks in the foreground and background, respectively. An ID card sample with a bacteriorhodopsin-based optical storage in the purple-colored strip is shown in Figure 6D. 3.3 Applications in Energy Conversion

The use of bacteriorhodopsin as a light energy-converting device seems to be ﬁrst choice as with regard to technical applications of the molecule [73, 74]. All applications of this type have as the key element in common a bacteriorhodopsin-driven charge separation over the PM. Although this is the basic function of bacteriorhodopsin embedded in PM, until now it has not been possible to prepare artiﬁcial PMs with technically relevant dimensions. Methods devised to overcome this problem have been unable to provide PMs capable of producing passive proton transport, and research is continuing in this area.

4 Methods

The methods required for this type of research span from biochemistry and bioengineering to various printing and optical techniques. All have been well documented, with the methods of cultivating halobacteria and isolating PMs being common to all applications. A comprehensive laboratory manual on Archaea was produced in 1995 [40]. For speciﬁc conditions of phototrophic growth [13], laboratory protocols of cultivation and media composition, the reader is referred to Ref. [40] (pages 13–21 and 225–230), while for the isolation of PMs from halobacteria, the reader is referred to Ref. [75]. The genetic modiﬁcation of halobacteria is described in Refs. [47, 76]. The very ﬁrst publication on the discovery of the bacteriorhodopsin [77], as well as the latest volume on its applications [78], should also be mentioned in this context. Information on speciﬁc processing steps for the wide range of applications is taken best from the relevant patent applications, and a summary of patents related to applications of bacteriorhodopsin may be found in Ref. [51].

5 Conclusions

Bacteriorhodopsin is today the biological photochromic material for which technical applications in optical information processing are much more developed than for any other biomaterial. The entire sequence, from an analysis of molecular

5 Conclusions 21

Fig. 6 Photochromic applications of bacteriorhodopsin. (A) Holographic correlator; (B) holographic camera for interferometric testing; (C) photochromic inks for security applications; (D) optical storage.

22 Bacteriorhodopsin and Its Potential in Technical Applications

function to its controlled modiﬁcation and the development of suitable applications, has been demonstrated with this molecule. Ideas of utilizing the evolutionary optimized functions of biological molecules in technical processes by using genetic engineering to produce tailor-made modiﬁed versions of natural molecules with improved technical properties have been demonstrated with bacteriorhodopsin for the ﬁrst time. Biomaterials as blueprints for technical materials with nanoscale functions form the basis of the concept of nanobionics – and bacteriorhodopsin was the ﬁrst such example.

References 1 E. DELONG, Science 1998, 280, 542–543. 2 E. F. DELONG, K. Y. WU, B. B. PREZELIN, R. V. M. JOVINE, Nature 1994, 371, 695–697. 3 E. BLO¨ CHL, R. RACHEL, S. BURGGRAF, D. HAFENBRADL, H. W. JANNASCH, K. O. STETTER, Extremophiles 1997, 1, 14–21. 4 R. AMILS, A. BALLESTER (eds.), Biohydrometallurgy and the environment toward the mining of the 21st century. . Elsevier, Amsterdam, Lausanne, New York, Oxford, Shannon, Singapore, Tokyo, 1999. 5 U. B. SLEYTR, B. SCHUSTER, D. PUM, IEEE Eng. Med. Biol. 2003, 22, 140–150. 6 W. D. GRANT, H. LARSEN, in: Bergey’s Manual of Systematic Bacteriology, N. PFENNIG (ed.), Williams Wilkins, Baltimore 1989, pp. 2216–2233. 7 A. OREN, FEMS Microbiol. Rev. 1994, 13, 415–440. 8 A. OREN, H. G. TRU¨ PER, FEMS Microbiol. Lett. 1990, 70, 33. 9 R. HARTMANN, H.-D. SICKINGER, D. OESTERHELT. Proc. Natl. Acad. Sci. USA 1980, 77, 3821–3825. 10 A. RUEPP, H. M¨ULLER, F. LOTTSPEICH, J. SOPPA, J. Bacteriol. 1995, 177, 1129–1136. 11 L. I. HOCHSTEIN, F. LANG, Arch. Biochem. Biophys. 1991, 288, 80. 12 D. OESTERHELT, Curr. Opinion Struct. Biol. 1998, 8, 489–500. 13 D. OESTERHELT, G. KRIPPAHL, Ann. Microbiol. Inst. Pasteur 1983 134B, 137–150. 14 K. W. FOSTER, J. SARANAK, N. PATEL, G. ZARILLI, M. OKABE, T. KLINE, K. NAKANISHI, Nature 1984, 311, 756–759.

15 J. A. BIESZKE, E. N. SPUDICH, K. L. SCOTT, K. A. BORKOVICH, J. L. SPUDICH, Biochemistry 1999, 38, 14138–14145. 16 O. B´EJA` , L. ARAVIND, E. V. KOONIN, M. T. SUZUKI, A. HADD, L. P. NGUYEN, S. B. JOVANOVICH, C. M. GATES, R. A. FELDMAN, J. L. SPUDICH, E. N. SPUDICH, E. F. DELONG, Science 2000, 289, 1902–1906. 17 K. PALCZEWSKI, T. KUMASAKA, T. HORI, C. A. BEHNKE, H. MOTOSHA, B. A. FOX, I. L. TRONG, D. C. TELLER, T. OKADA, R. E. STENKAMP, M. YAMAMOTO, M. MIYANO, Science 2000, 289, 739–745. 18 L.-O. ESSEN, R. SIEGERT, W. D. LEHMANN, D. OESTERHELT, Proc. Natl. Acad. Sci. USA 1998, 95, 11673–11678. 19 M. KOLBE, H. BESIR, L.-O. ESSEN, D. OESTERHELT, Science 2000, 288, 1390–1396. 20 V. I. GORDELLY, J. LABAHN, R. MOUKHAMETHLANOC, R. EFREMOV, J. GRANZIN, R. SCHLESINGER, G. B¨ULDT, T. SAVOPOL, A. J. SCHELDIG, J. P. KLARO, M. ENGELHARDT, Nature 2002, 439, 464–465. 21 G. WAGNER, R. HARTMANN, D. OESTERHELT, Eur. J. Biochem. 1978, 89, 169–179. 22 D. OESTERHELT, Israel J. Chem. 1995, 35, 475–494. 23 W. MARWAN, D. OESTERHELT. ASM-News 2000, 66, 83–90. 24 K. IHARA, T. UMEMURA, I. KATAGIRI, T. KITAJIMA-IHARA, Y. SUGIYAMA, Y. KIMURA, Y. MUKOHATA, J. Mol. Biol. 1999, 285, 163–174. 25 J. SASAKI, L. S. BROWN, Y. S. CHON, H. KANDORI, A. MAEDA, R. NEEDLEMAN, J. K. LANYI, Science 1995, 269, 73–75.

References 26 J. TITTOR, U. HAUPTS, Ch. HAUPTS, D. OESTERHELT, A. BECKER, E. BAMBERG, J. Mol. Biol. 1997, 271, 405–416. 27 U. HAUPTS, J. TITTOR, D. OESTERHELT, Annu. Rev. Biophys. Biomol. Struct. 1999, 28, 367–399. 28 R. RAMMELSBERG, G. HUHN, M. L¨UBBEN, K. GERWERT, Biochemistry 1998, 37, 5001–5009. 29 M. WIKSTRO¨ M, Curr. Opin. Struct. Biol. 1998, 8, 480–488. 30 N. A. DENCHER, H. J. SASS, G. B¨ULDT, Biochim. Biophys. Acta 2000, 1460, 192–203. 31 J. HEBERLE, Biochim. Biophys. Acta 2000, 1458, 135–147. 32 H. LUECKE, B. SCHOBERT, H.-T. RICHTER, J.-P. CARTAILLER, J. K. LANYI, Science 1999, 286, 255–260. 33 J. le COUTRE, J. TITTOR, D. OESTERHELT, K. GERWERT, Proc. Natl. Acad. Sci. USA 1995, 92, 4962–4966. 34 J. RIESLE, D. OESTERHELT, N. A. DENCHER, J. HEBERLE, Biochemistry 1996, 35, 6635–6643. 35 U. HAUPTS, J. TITTOR, E. BAMBERG, D. OESTERHELT, Biochemistry 1997, 36, 2–7. 36 A. MILLER, D. OESTERHELT, Biochim. Biophys. Acta 1990, 1020, 57–64. 37 R. L. CHARLEBOIS, W. L. LAM, S. W. CLINE, W. F. DOOLITTLE, Proc. Natl. Acad. Sci. USA 1987, 84, 8530–8534. 38 S. W. CLINE, W. L. LAM, R. L. CHARLEBOIS, L. S. SCHALKWYK, W. F. DOOLITTLE, Can. J. Microbiol. 1989, 35, 148–152. 39 B. NI, M. CHANG, A. DUSCHL, J. LANYI, R. NEEDLEMAN, Gene 1990, 90, 169–172. 40 F. T. ROBB, A. R. PLACE, K. R. SOWERS, H. J. SCHREIER, S. DASSARMA, E. M. FLEISCHMANN (eds), Archaea. A Laboratory Manual. Cold Spring Harbor Laboratory Press, 1995. 41 M. L. HOLMES, S. D. NUTTALL, M. L. DYALL-SMITH, J. Bacteriol. 1991, 173, 3807–3813. 42 G. WAGNER, D. OESTERHELT, G. KRIPPAHL, J. K. LANYI, FEBS Lett. 1981, 131, 341–345. 43 M. PFEIFFER, T. RINK, K. GERWERT, D. OESTERHELT, H.-J. STEINHOFF, J. Mol. Biol. 1999, 287, 163–171. 44 W. L. LAM, W. F. DOOLITTLE, Proc. Natl. Acad. Sci. USA 1989, 86, 5478–5482.

45 D. GREGOR, F. PFEIFER, MICROBIOLSGM 2001, 147, 1745–1754. 46 N. PATENGE, A. HAASE, H. BOLHUIS, D. OESTERHELT, Mol. Microbiol. 2000, 36, 105–113. 47 M. DYALL-SMITH, The Halohandbook: Protocols for halobacterial genetics, 2001. http://www.microbiol.unimelb.edu.au/ staff/mds/HaloHandbook/index.html 48 [email protected] 49 R. R. BIRGE, Annu. Rev. Phys. Chem. 1990, 41, 683–733. 50 N. HAMPP, Chem. Rev. 2000, 100, 1755–1776. 51 N. HAMPP, Appl. Microbiol. Biotechnol. 2000, 53, 633–639. 52 B. GABRIEL, J. TEISSIE, Proc. Natl. Acad. Sci. USA 1996, 93, 14521–14525. 53 K. KOYAMA, N. YAMAGUCHI, T. MIYASAKA, Science 1994, 265, 762–765. 54 F. T. HONG, Prog. Surf. Sci. 1999, 62, 1–237. 55 H. W. TRISSL, M. MONTAL, Nature 1977, 266, 655–657. 56 F. T. HONG, BioSystems 1995, 35, 117–121. 57 T. MIYASAKA, K. KOYAMA, I. ITOH, Kagaku to Kogyo (Tokyo) 1993, 46, 247–249. 58 Z. CHEN, R. R. BIRGE, Trends Biotechnol. 1993, 11, 292–300. 59 T. MIYASAKA, K. KOYAMA, I. ITOH, Science 1992, 255, 342–344. 60 C. RITZEL, K. MEERHOLZ, C. BRA¨ UCHLE, D. OESTERHELT, Mol. Cryst. Liq. Cryst. Sci. Technol. A 1998, 315, 443–448. 61 C. RITZEL, K. MEERHOLZ, C. BRA¨ UCHLE, D. OESTERHELT, Proc. SPIE-Int. Soc. Opt. Eng. 1998, 3475, 49–55. 62 N. HAMPP, C. BRA¨ UCHLE, D. OESTERHELT, Biophys J. 1990, 58, 83–93. 63 M. FRYDRCH, P. SILFSTEN, S. PARKKINEN, J. PARKINNEN, T. JAASKELAINEN, Biosystems 2000, 54, 131–140. 64 N. HAMPP, T. FISCHER, M. NEEBE, Proc. SPIE 2002, 4677, 121–128. 65 P. KOLODNER, E. LUKASHEV, Y.-C. CHING, I. ROUSSO, Proc. Natl. Acad. Sci. USA 1996, 53, 6012–6015. 66 N. N. VSEVOLODOV, T. V. DYUKOVA, Trends Biotechnol. 1994, 12, 81–88. 67 A. POPP, M. WOLPERDINGER, N. HAMPP, C. BRA¨ UCHLE, D. OESTERHELT, Biophys. J. 1993, 65, 1449–1459.

23

24 Bacteriorhodopsin and Its Potential in Technical Applications

68 N. B. GILLESPIE, K. J. WISE, L. REN, J. A. STUART, D. L. MARCY, Q. LI, L. RAMOS, K. JORDAN, L. REN, R. R. BIRGE, J. Phys. Chem. B 2002, 106, 13352–13361. 69 D. HARONIAN, A. LEWIS, Appl. Opt. 1991, 30, 597–608. 70 R. R. BIRGE, R. B. GROSS, M. B. MASTHAY, J. A. STUART, J. R. TALLENT, C. F. ZHANG, Mol. Cryst. Liq. Cryst. Sci. Technol. 1992, B3, 133–148. 71 R. THOMA, N. HAMPP, Opt. Lett. 1994, 19, 1364–1366. 72 R. THOMA, M. DRATZ, N. HAMPP, Opt. Eng. 1995, 34, 1345–1351.

73 M. EISENBACH, C. WEISSMANN, G. TANNY, S. R. CAPLAN, FEBS Lett. 1977, 81, 77–80. 74 D. OESTERHELT, FEBS Lett. 1976, 64, 20–22. 75 OESTERHELT, D. STOECKENIUS, W. Methods Enzymol. 1974, 31, 667–678. 76 K. J. WISE, N. B. GILLESPIE, J. A. STUART, M. P. KREBS, R. R. BIRGE, Trends Biotechnol. 2002, 20, 387–394. 77 D. OESTERHELT, W. STOECKENIUS, Nature New Biol. 1971, 233, 149–152. 78 N. VSEVOLODOV, Biomolecular Electronics, Birkh¨auser, Boston, Basle, Berlin, 1998.

1

Biomolecular Motors Operating in Engineered Environments Stefan Diez, Jonne H. Helenius, and Jonathon Howard Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany

Originally published in: Nanobiotechnology. Edited by Christof M. Niemeyer and Chad A. Mirkin. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30658-9

1 Overview

Recent advances in understanding how biomolecular motors work has raised the possibility that they might find applications as nanomachines. For example, they could be used as molecule-sized robots that: Ĺ work in molecular factories where small, but intricate structures are made on tiny assembly lines; Ĺ construct networks of molecular conductors and transistors for use as electrical circuits; Ĺ or that continually patrol inside “adaptive” materials and repair them when necessary.

Thus, biomolecular motors could form the basis of bottom-up approaches for constructing, active structuring and maintenance at the nanometer scale. We will review the current status of the operation of biomolecular motors in engineered environments, and discuss possible strategies aimed at implementing them in nanotechnological applications. We cite reviews whenever possible for the biochemical and biophysical literature, and include primary references to the nanotechnological literature. Biomolecular motors are the active workhorses of cells [1]. They are complexes of two or more proteins that convert chemical energy – usually in the form of the high-energy phosphate bond of ATP – into directed motion. The most familiar motor is the protein myosin which moves along filaments, formed from the protein actin, to drive the contraction of muscle. In fact, all cells – not just specialized

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Biomolecular Motors Operating in Engineered Environments

muscle cells – contain motors that move cellular components such as proteins, mitochondria, and chromosomes from one part of the cell to another. These motors include relatives of muscle myosin (that also move along actin filaments), as well as members of the kinesin and dynein families of proteins. The latter motors move along another type of filament called the microtubule. The reason that motors are necessary in cells is that diffusion is too slow to transport molecules efficiently from where they are made (which typically is near the nucleus) to where they are used (which is often at the periphery of the cell). For example, the passive diffusion of a small protein to the end of a 1 meter-long neuron would take approximately 1000 years, yet kinesin moves it in a week. This corresponds to a speed of 1–2 µm s−1 , which is typical for biomolecular motors [2]. Actin filaments and microtubules form a network of highways within cells, and localized cues are used to target specific cargoes to specific sites in the cell [3]. By using filaments and motors, cells build highly complex and active structures on the molecular (nanometer) scale. Little imagination is needed to envisage employing biomolecular motors to build molecular robots [4]. Biomolecular motors are unusual machines that do what no man-made machines do: they convert chemical energy to mechanical energy directly rather than via an intermediate such as heat or electrical energy. This is essential because the confinement of heat, for example, on the nanometer scale is not possible because of its high diffusivity in aqueous solutions [2]. As energy converters, biomolecular machines are highly efficient. The chemical energy available from the hydrolysis of ATP is 100 × 10−21 J = 100 pN nm−1 (under physiological conditions, where the ATP concentration is 1 mM and the concentrations of the products ADP and phosphate are 0.01 mM and 1 mM, respectively). With this energy, a kinesin molecule is able to perform an 8-nm step against a load of 6 pN [2]. The energy efficiency is therefore almost 50%. For the rotary motor F1 F0 -ATPase synthase which uses the electrochemical gradient across mitochondrial and bacterial membranes to generate ATP, the efficiency is reported to be between 80 and 100% [5, 6]. This high efficiency demonstrates that, like other biological systems, the operation of biological motors has been optimized through evolution. High efficiency is but one feature that makes biomolecular motors attractive for nanotechnological applications. Other features are: 1. They are small and can therefore operate in a highly parallel manner. 2. They are easy to produce and can be modified through genetic engineering. 3. They are extremely cheap. For example, 20 × 109 kinesin motors can be acquired for 1 US cent from commercial suppliers (1 mg = 3.3 × 1015 motors cost $1500; Cytoskeleton, Inc., Colorado, USA) and the price could be significantly decreased if production were scaled up. 4. A wide array of biochemical tools have been developed to manipulate these proteins outside the cell.

1 Overview

This review focuses on two broad categories of molecular motors: Ĺ Linear motors generate force as they move along intracellular filaments. In addition to myosin and kinesin mentioned above, linear motors also include enzymes that move along DNA and RNA. Ĺ Rotary motors generate torque via the rotation of a central core within a larger protein complex. They include ATP synthase, mentioned above, as well as the motor that drives bacterial motility.

Representatives of both categories have been used to manipulate molecules and nanoparticles. Mechanical and structural properties of relevant filaments are contained in Table 1, and those of several associated motors in Table 2. The general set-ups for studying motor proteins outside cells – the so-called motility assays – are depicted in Figure 1. In the gliding assay, the motors are immobilized on a surface and the filaments glide over the assembly (Figure 1A). In the stepping assay, the filaments are laid out on the surface where they form tracks for the motors to move along (Figure 1B). In both assays, movement is observed under the light microscope using fluorescence markers or highcontrast techniques. Variations on these assays have been used to reconstitute linear motility on the four types of filaments – actin filaments, microtubules, DNA, and RNA. The gliding motility assay has provided detailed data on the directionality, speed, and force generation of purified molecular motors [2, 7]. However, for use in nanotechnological applications, the movement of gliding filaments must be controllable in space and time. For example, a simple application would be to employ a moving filament to pick up cargo at point A, move it along a user-defined path to point B, and then release it. A number of methods for the spatial and temporal control of filament movement have been developed. Spatial control has been achieved using topographical features [8–11], chemical surface modifications [10, 12–14], and a combination of both [15–18]. Electrical fields [19–21] and hydrodynamic flow [22, 23] have also been used to direct the motion of gliding filaments. An example from our laboratory of gliding microtubules that are guided by channels is shown in Figure 2. Temporal control has been achieved by manipulating the ATP concentration [9, 24]. In addition to these basic techniques for controlling motion, some simple applications of the gliding assay have been demonstrated. These include the transport of streptavidin-coated beads [9], the transport and stretching of individual DNA molecules [25], the measurement of forces in the pN range [26], and the imaging of surfaces [27]. The stepping assay opens up additional possibilities. Initially, micrometer-sized beads were coated with motor proteins and visualized as they moved along filaments. The movement of beads can be tracked with nanometer precision to determine the speed and step size [2], and the use of optical tweezers allows forces to

3

13 2

25 nm 2 nm

2 nm

RNA

2

2

6 nm

Actin filament Microtubule DNA

Strands per filament

Diameter

Filament

0.34 nm

8 nm 0.34 nm

5.5 nm

Repeat length

75 nm

5 mm 50 nm

10 µm

Persistence length

1.5 GPa

2 GPa 1 GPa

2 GPa

Young’s modulus

30 µm

10 cm 100 mm

100 µm

Maximum length

Kinesin, Dynein RNA polymerase, DNA helicase, topoisomerase Ribosome

Myosin

Motors

80

78 79

77

Reference

where k is the Boltzmann constant and T is absolute temperature. Young’s modulus (E) is calculated assuming that the filament is homogenous and isotropic. The repeat length describes the periodicity along a strand of the filament

Table 1 Physical attributes of actin filaments, microtubules, DNA, and RNA. The persistence length (Lp) is related to the flexural rigidity (EI) by: Lp = EI/kT,

4 Biomolecular Motors Operating in Engineered Environments

The sizes refer to the motor domains.

120◦

8 × 14 45

*

0.34 up to 43 nm/turn 0.34

15

DNA DNA DNA pilus NA NA

0.34

5 36 8

16 24 6 24

Actin Actin Microtubule Microtubule DNA

Myosin II Myosin V Conventional kinesin Dynein T7 DNA polymerase (exonuclease activity) RNA polymerase Topoisomerase Bacteriophage portal motor Type IV pilus retraction motor F1 -ATPase Flagellar motor

Step size [nm]

Filament

Motor

Size* [nm]

5 − 100 bps 1000 8 rps 300 rps

30 000 300 800 6400 >100 bps

Maximum speed [nm s−1 ]

57 pN 110 pN 100 pN nm 550 pN nm

25 pN

10 pN 1.5 pN 6 pN 6 pN 34 pN

Maximum force [pN]

80

NA NA

NA

50 50 50

Efficiency [%]

Table 1. The sizes refer to the motor domains. Dynamic parameters were determined by in-vitro experiments at high ATP concentration

87, 88 89, 90 91 92, 93 6 94

2, 81 82 2, 83 84, 85 86

Reference(s)

Table 2 Values characterizing the operation of several important biomolecular motors. The filaments along which the linear motors operate are indicated in

1 Overview 5

6 Biomolecular Motors Operating in Engineered Environments

Fig. 1 Biomolecular motor systems currently applicable for nanotechnological developments. (A) Linear transport of filaments by surface bound motor molecules (gliding assay). (B) Linear movement of motor proteins along filaments (stepping assay). (C) Rotation generated by a rotary motor.

1 Overview

Fig. 2 (A) Directed movement of gliding microtubules along microstructured polyurethane channels on the surface of a coverslip. The initial positions of the microtubules are shown in orange, while the paths they traveled over the subsequent 12 s are shown in green. (B) Scanning electron microscopy image of the polyurethane channels. The channels are a replica mold of a

Si-master (channel width 500 nm, periodicity 1000 nm, depth 300 nm) produced using a poly(dimethylsiloxane) (PDMS) stamp as an intermediate. Note, that the ridges have been “undercut”. This probably aids the guiding of the microtubules in the channels. (Silicon master provided by T. Pompe, Institute of Polymer Research, Dresden, Germany.)

be measured [28]. In addition to beads, 10 (µm-diameter glass particles [29] and Simicrochips [30] have been transported and membrane tubes have been pulled [32]a along filaments. In another variation, high-sensitivity fluorescence microscopy is used to visualize individual motor molecules as they step along filaments [31, 32]. An example from our laboratory of a single kinesin motor fused to the green fluorescent protein moving along a microtubule is shown in Figure 3. Despite the power of single-molecule techniques, they have yet to be exploited for nanotechnological applications. Rotary motors can be studied in vitro by fixing the stator to a surface and following the movement of the rotor (see Figure 1C). Rotation can be visualized under the light microscope by attaching a fluorescent label or a microscopic marker to the rotor. Both techniques have been used to investigate the stepwise rotation generated by F1 -ATPase, which is a component of the F1 F0 -ATP synthesis machinery [5, 33]. Individual motors have been integrated into nanoengineered environments by arraying them on a nanostructured surface and using them to rotate fluorescent microspheres [34] or to drive Ni-nanopropellers [6].

7

8 Biomolecular Motors Operating in Engineered Environments

Fig. 3 Movement of a single kinesin molecule (labeled with the green fluorescent protein) along a microtubule (red). Micrographs were acquired at the indicated times using total-internal-reflection fluorescence microscopy.

2 Methods

There are many challenges in applying biomolecular motors to nanotechnology. Motility must be robust, it must be controlled both spatially and temporally, and the motors must be hitched to and unhitched from their cargoes. This section summarizes key techniques towards these ends. 2.1 General Conditions for Motility Assays

Motility assays are performed in aqueous solutions that must fulfill a number of requirements. We will illustrate these requirements with the kinesin/microtubule system. Kinesin uses ATP as its fuel; the maximum speed is reached at ∼0.5 mM, approximately equal to the cellular concentration. Other nucleotides such as GTP, TTP, and CTP can substitute for ATP, but the speed is lower [35]. Motility also

2 Methods 9

requires divalent cations, with magnesium preferred over calcium, and strontium and barium unable to substitute [36]. Optimal motility, assessed by gliding speed, occurs over a range of pH, between 6 and 9 [35, 37], and over a range of ionic strengths, between 50 mM and 300 mM [37]. The speed increases with temperature, doubling for each 10◦ C between 5◦ C and 50◦ C [24, 38]; motility fails at higher temperatures. The force is independent of temperature between 15◦ C and 35◦ C [39]. When assays are performed in the middle of these ranges, motility is robust and only a small drop in the mean velocities is seen after 3 hours [24, 37]. If fluorescent markers are used, then an oxygen-scavenging enzyme system must be present in order to prevent photodamage. Many experimental details, including a discussion of the densities of the motors, can be found in Ref. [7].

2.2 Temporal Control

Motors can be reversibly switched off and on by regulating the concentration of fuel, or by adding and removing inhibitors. The ATP concentration can be rapidly altered by flowing in a new solution. In such a setup, the kinesin-dependent movement of microtubules can be stopped within 1 s and restarted within 10 s (unpublished data from our laboratory). Similarly, inhibitors such as AMP–PNP (a non-hydrolyzable analogue of ATP [40]), adocia-sulfate-2 (a small molecule isolated from sponge [41]) and monastrol [42] can be perfused to stop motility. An alternative method to control energy supply is to use photoactivatable ATP. In this method, a flash of UV light is used to release ATP from a derivatized, nonfunctional precursor; an ATP-consuming enzyme is also present to return the ATP concentration to low levels following release. Using such a system, microtubule movement has been repeatedly started and stopped [9], though the start-up and slow-down times were slow, on the order of minutes. The advantage of this method is that the solution in the flow cell does not have to be exchanged. Fortuitously, many proteins possess natural regulatory mechanisms and, once understood, these might offer additional means to regulate the motors in vitro. Examples include the regulation of myosins by phosphorylation and calcium/calmodulin [43] and the inhibition of kinesin by its cargo-binding “tail” domain [44]. Because such natural controls might not always be applicable in a synthetic environment, there is strong interest in the development of artificial control mechanisms for motor proteins. Towards this end, metalion binding sites have been genetically engineered into the F1 -ATPase motor. The binding of ions at the engineered site immobilizes the moving parts of the motor, thus inhibiting its rotation [45]. ATP-driven rotation can be restored by the addition of metal ion chelators. Clever genetic engineering of motors could provide temporal control mechanisms that may be switched by temperature, light, electrical fields, or buffer composition.

10 Biomolecular Motors Operating in Engineered Environments

2.3 Spatial Control

In order to control the path along which filaments glide – a process that we call “guiding” – it is necessary to restrict the location of active motors to specific regions of a surface. This can be done by coating a glass or silicon surface with resist polymers such as PMMA, SU–8, or SAL601 and using UV, electron beam or soft lithography to remove resist from defined regions [12–19]. The motor-containing solution is then perfused across the surface. By choosing appropriate properties of this solution [e. g., the concentration of motors, salts, other blocking proteins such as casein and bovine serum albumin (BSA), and detergents such as Triton X-100], motility can be restricted to either the unexposed, resist surface or to the exposed, underlying substrate. For example, it has been found that myosin motility is primarily restricted to the more hydrophobic resist surfaces while kinesin motility is primarily restricted to the more hydrophilic non-resist surfaces. However, the detailed interactions of the motors with these surfaces are not well understood. One limitation of this approach to binding proteins to surfaces is that the motors tend to bind everywhere, so it is difficult to attain good contrast. A proven method to prevent motor binding is to coat a surface with polyethylene oxide (PEO) [10, 46]. Techniques to bind motors and filaments via affinity tags to surfaces are summarized in section 2.4. While chemical patterning can restrict movement of filaments to areas with a high density of active motors, walking off the trails is not prevented. This was demonstrated by Hess et al. [10], who showed that microtubules move straight across a boundary between high motor density (non-PEO) and low motor density (PEO), where they dissociate from the surface. The problem with a purely chemical pattern is that if a rigid filament is propelled by several motors along its length, there is nothing to stop the motors at the rear from pushing the filament across a boundary into an area of low motor density. The behavior of microtubules colliding with the walls of channels imprinted in polyurethane has been studied by Clemmens et al. [11]. They found that the probability of a filament being guided by the walls decreased as the approach angle increased. At high incident angles, guiding was not observed and instead the microtubules climbed the walls. Combining chemical and topographic features – as occurs in the lithographic studies described above – leads to more efficient guiding. For example, in the study of Moorjani et al. [18], filaments remained at the bottom of the channels formed in the SU–8 even when they collided with the walls at angles above 80◦ . When the leading end of the microtubule hits the wall, the motors at the rear force the microtubule to bend into the region of high motor density, and in this way the motion is guided by the boundary (see Figure 4, unpublished results from our laboratory). While it is possible to use chemical and topographical patterning to guide filaments – that is, to restrict their movement to particular paths – it is more difficult to control the direction of movement along the path. The difficulty arises because the orientation in which motors bind to a uniform surface is not controlled. Some motors will be oriented so that they propel filaments in one direction along the path, whereas others will propel filaments in the opposite direction. The reason

2 Methods

Fig. 4 Sequence of fluorescent images showing the kinesin-driven, unidirectional movement of a rhodamine-labeled microtubule (red) along chemically and topographically structured Si-chip. The bottom of the channels (green), the depths of which

are 300 nm, is coated with kinesin. The surrounding regions are blocked by polyethylene glycol. (Research in collaboration with R. M.M. Smeets, M.G.L. van den Heuvel, and C. Dekker, Delft University of Technology, The Netherlands.)

that motors do not counteract each other is that filaments are polar structures: the orientation of the proteins that form up the filaments is maintained all along the length of the filament (see Figure 1). Because the motors bind stereospecifically to the filament, they will exert force in only one direction. Thus, the orientation of the filament determines its direction of motion; one end always leads. The direction of filament gliding can be controlled by the application of external forces. Actin filaments and microtubules both possess negative net charges and, consequently, in the presence of a uniform electric field, will experience a force directed towards the positive electrode. It is possible to apply high enough electric fields to steer motor-driven filaments in a specified direction [19, 21]. Because the refractive index of protein differs from that of water, filaments become electrically polarized in the presence of an electric field, and consequently in a nonuniform field they move in the direction of highest field strength. This so-called dielectrophoretic force has been used to direct the gliding of actin filaments on a myosin-coated substrate [20]. It is even possible to manipulate a microtubule using optical gradients produced by focusing a laser beam (i. e., an optical tweezers) [47]. Directional control of microtubule gliding has also been achieved using hydrodynamic flow fields [23, 29]. An alternative approach to directionality relies on more sophisticated guiding concepts. For example, unidirectional movement of filaments can be achieved if

11

12 Biomolecular Motors Operating in Engineered Environments

guiding geometries based on arrow and ratchet structures are employed [10, 15]. An example of the unidirectional movement of a microtubule on a topographically and chemically structured silicon chip is depicted in Figure 4 To control the direction of motion in stepping assays, the orientation of the filaments on the surface must be controlled. Towards this end, the generation of isopolar filament arrays has been achieved by binding specific filament ends to a surface, and using hydrodynamic flow to align the filaments along the surface to which they are subsequently adhered to [30, 48–50]. Alternatively, moving filaments can be aligned in a particular orientation by a flow field prior to fixation by glutaraldehyde [23, 29], which has been shown not to interfere with kinesin motility [51]. Fluid flow has also been used to align microtubules binding to patterned silane surfaces, though the orientation of the microtubules was not controlled [52]. 2.4 Connecting to Cargoes and Surfaces

Cargoes can be attached to filaments using several different approaches. The prospective cargo can be coated with an antibody to the filament [53] or to a filament-binding protein such as gelsolin [54]. A clever refinement of this technique is genetically to fuse gelsolin with a cargo protein, thereby generating a dual-functional protein [55]. Alternatively, the cargo can be coated with streptavidin which binds to filaments that have been derivatized with biotin [56]. There are many other possibilities which have not yet been realized. Analogous methods can be used to couple motors to surfaces. For example, the motor can be fused with the bacterial biotin-binding protein [57] and in this way bound to streptavidin-coated cargoes or surfaces. There any many peptide tags that can be fused to proteins to aid their purification [58, 59]. These tags can be used to couple these proteins to surfaces coated with the complementary ligand. A popular tag is the hexahistidine tag which binds Ni2+ and other metals that are chelated to nitriloamines (NTA). A nice approach is to couple the NTA to the terminal ethyleneoxides of triblock copolymers containing PEO. In principle, this provides specific binding of a histagged motor (or another protein) to a surface while the PEO groups block nonspecific binding [46, 60]. Controlled unloading of cargo has not been demonstrated, but ought to be feasible. For example, there are biotins that can be irreversibly cleaved by light and reversibly cleaved by reducing agents, and the histidine-Ni2+ -NTA connection can be broken by sequestering the Ni2+ with EDTA.

3 Conclusions

Although the first steps have been made towards the operation of biomolecular motors in engineered environments, many advances are necessary before these

3 Conclusions

motors can be used in nanotechnological applications such as working in molecular factories and building circuits. An immediate task is to improve the spatial and temporal control over the motors. By combining improved surface techniques with the application of external electric, magnetic, and/or optical fields it should be possible, in the near future, to stretch and collide single molecules, to control cargo loading and unloading, and to sort and pool molecules. Another goal is to control the position and orientation of motors with molecular precision. This means placing motors with an accuracy of ∼10 nm on a surface and controlling their orientation within a few degrees. In this way both the location and the direction of motion of filaments can be controlled. One approach to molecular patterning is to “decorate” filaments with stereospecifically bound motors. Once aligned along the filament matrix, the motors can be transferred to another surface. This approach was taken by Spudich et al. [48, 61] and should be followed up. A further development of this idea is to directly produce (perhaps by stamping a mold made with a filament) surfaces that have structures functionally similar to motor-binding sites. An alternative approach is to use dippen lithography or other AFM techniques [62] to directly pattern motors on surfaces. The robustness of motors must be increased. Motors operate only in aqueous solutions and under a restricted range of solute concentrations and temperatures. While it is inconceivable that protein-based motors could operate in a nonaqueous environment, two approaches to increasing their robustness can be envisaged. First, motors could be purified from thermophilic or halophilic bacteria, some of which grow at temperatures up to 112◦ C and salinities above 5 M. There are also extreme eukaryotes that grow at up to 62◦ C or 5 M NaCl. This approach has already been taken for ATP synthase [63], but not with linear motors because no obvious homologues of myosins or kinesins have been found in bacteria. Second, a genetic screening approach might reveal mutations that allow motors to operate in less restrictive or different conditions. A longer-term goal is to use the design principles learnt from the study of biomolecular motors to build purely artificial nanomotors that can operate in air or vacuum. This is a daunting prospect however, and it is not even clear what fuel(s) might be used. A potential way forward is to use chemical energy from a surface: for example, it was demonstrated that tin particles slide across copper surfaces driven by the formation of bronze alloy [64], this being analogous to paraffin-driven toy boats. Besides the motor systems discussed so far, other biomechanical assemblies are good candidates for nanotechnological applications. In addition to providing paths along which motors move, active biological filaments on their own might find use in nanotechnological applications. The pushing and pulling forces generated by the polymerization and depolymerization of actin filaments and microtubules provide an alternative method of moving molecules [2, 65]. This ability is of particular interest because bacteria possess actin- [66] and microtubule-like [67] filaments and, as mentioned above, the proteins of extremophilic bacteria function in extreme environmental conditions. Filaments and motors can also self-organize under certain conditions [68–71]. On a side note, the flagellar filament in conjunction

13

14 Biomolecular Motors Operating in Engineered Environments

with the flagellar motors allow the bacteria to move in three-dimensional liquid space [72]. In addition to the motors that we have described so far, cells contain numerous biomolecular machines that can also be thought of as motors (for example, see Ref. [3]). These machines use chemical energy to replicate DNA (DNA polymerases) and process it (recombinases, topoisomerases and endonucleases), to produce RNA (RNA polymerases) and splice it (spliceosomes), to make proteins (ribosomes) and fold them (chaperones) and move them across membranes (translocases), and finally destroy them (proteasomes). The energy is provided by another group of machines that generate the electrochemical gradients (electron transport system, bacteriorhodopsin) used by the F1 F0 -ATP synthase to make ATP or by flagellar motors to propel bacteria. All these machines are candidates for nanotechnological applications, and a recent report of the use of chaperones to maintain nanoparticles in solution [73] is a step in this general direction. We finish up by pointing out that the high order and nanometer-scale periodicity of DNA, actin filaments and microtubules make them ideal scaffolds on which to erect three-dimensional nanostructures. While these features have been exploited to make DNA-based structures [74], the use of DNA motors to address specific sites (based on nucleotide sequence) has not, to our knowledge, been realized. Some years ago it was proposed that the regular lattice of microtubules might serve as substrates for molecular computing and information storage [75, 76]. While these ideas seem crazy in the context of the living organism, they may be realizable for biomolecular motors operating in engineered environments. At the moment, anything is possible!

Acknowledgements

The authors thank U. Queitsch for help with experiments on guiding microtubules, T. Pompe, R. M.M. Smeets, M.G.L. van den Heuvel, and C. Dekker for fabricating microstructured channels, and F. Friedrich for assistance with the illustrations.

References 1 B. ALBERTS, Cell 1998, 92, 291. 2 J. HOWARD, Mechanics of Motor Proteins and the Cytoskeleton, Sinauer Associates, Sunderland, MA, 2001. 3 B. ALBERTS, Molecular biology of the cell, 4th edition, Garland Science, New York, 2002. 4 M. CRICHTON, Prey, HarperCollins, Toronto, 2002. 5 K. KINOSITA, Jr., R. YASUDA, H. NOJI, S. ISHIWATA, M. YOSHIDA, Cell 1998, 93, 21.

6 R. K. SOONG, G. D. BACHAND, H. P. NEVES, A. G. OLKHOVETS, H. G. CRAIGHEAD, C. D. MONTEMAGNO, Science 2000, 290, 1555. 7 J. M. SCHOLEY, Motility assays for motor proteins, Academic Press, San Diego, 1993. 8 J. R. DENNIS, J. HOWARD, V. VOGEL, Nanotechnology 1999, 10, 232. 9 H. HESS, J. CLEMMENS, D. QIN, J. HOWARD, V. VOGEL, Nano Lett. 2001, 1, 235.

References 10 H. HESS, J. CLEMMENS, C. M. MATZKE, G. D. BACHAND, B. C. BUNKER, V. VOGEL, Appl. Physics A (Materials Science Processing) 2002, A75, 309. 11 J. CLEMMENS, H. HESS, J. HOWARD, V. VOGEL, Langmuir 2003, 19, 1738. 12 H. SUZUKI, A. YAMADA, K. OIWA, H. NAKAYAMA, S. MASHIKO, Biophys. J. 1997, 72, 1997. 13 D. V. NICOLAU, H. SUZUKI, S. MASHIKO, T. TAGUCHI, S. YOSHIKAWA, Biophys. J. 1999, 77, 1126. 14 J. WRIGHT, D. PHAM, C. MAHANIVONG, D. V. NICOLAU, M. KEKIC, C. G. DOS REMEDIOS, Biomed. Microdevices 2002, 4, 205. 15 Y. HIRATSUKA, T. TADA, K. OIWA, T. KANAYAMA, T. Q. P. UYEDA, Biophys. J. 2001, 81, 1555. 16 C. MAHANIVONG, J. P. WRIGHT, M. KEKIC, D. K. PHAM, C. DOS REMEDIOS, D. V. NICOLAU, Biomed. Microdevices 2002, 4, 111. 17 R. BUNK, J. KLINTH, L. MONTELIUS, I. A. NICHOLLS, P. OMLING, S. TAGERUD, A. MANSSON, Biochem. Biophys. Res. Commun. 2003, 301, 783. 18 S. G. MOORJANI, L. JIA, T. N. JACKSON, W. O. HANCOCK, Nano Lett. 2003, 3, 633. 19 D. RIVELINE, A. OTT, F. JULICHER, D. A. WINKELMANN, O. CARDOSO, J. J. LACAPERE, S. MAGNUSDOTTIR, J. L. VIOVY, L. GORRE-TALINI, J. PROST, Eur. Biophys. J. Biophys. Lett. 1998, 27, 403. 20 S. B. ASOKAN, L. JAWERTH, R. L. CARROLL, R. E. CHENEY, S. WASHBURN, R. R. SUPERFINE, Nano Lett. 2003, 3, 431. 21 R. STRACKE, K. J. BOHM, L. WOLLWEBER, J. A. TUSZYNSKI, E. UNGER, Biochem. Biophys. Res. Commun. 2002, 293, 602. 22 P. STRACKE, K. J. BOHM, J. BURGOLD, H. J. SCHACHT, E. UNGER, IOP Publishing. Nanotechnology 2000, 11, 52. 23 I. PROTS, R. STRACKE, E. UNGER, K. J. BOHM, Cell Biol. Int. 2003, 27, 251. 24 K. J. BOHM, R. STRACKE, M. BAUM, M. ZIEREN, E. UNGER, FEBS Lett. 2000, 466, 59. 25 S. DIEZ, C. REUTHER, C. DINN, R. SEIDEL, M. MERTIG, W. POMPE, J. HOWARD, Nano Lett. 2003, 3, 1251–1254. 26 H. HESS, J. HOWARD, V. VOGEL, Nano Lett. 2002, 2, 1113.

27 H. HESS, J. CLEMMENS, J. HOWARD, V. VOGEL, Nano Lett. 2002, 2, 113. 28 A. D. MEHTA, M. RIEF, J. A. SPUDICH, D. A. SMITH, R. M. SIMMONS, Science 1999, 283, 1689. 29 K. J. BOHM, R. STRACKE, P. MUHLIG, E. UNGER, IOP Publishing. Nanotechnology 2001, 12, 238. 30 L. LIMBERIS, R. J. STEWART, Proceedings of Spie: the International Society for Optical Engineering, SPIE Int. Soc. Opt. Eng. 1998, 3515, 66. 31 R. D. VALE, T. FUNATSU, D. W. PIERCE, L. ROMBERG, Y. HARADA, T. YANAGIDA, Nature 1996, 380, 451. 32 A. YILDIZ, J. N. FORKEY, S. A. MCKINNEY, T. HA, Y. E. GOLDMAN, P. R. SELVIN, Science 2003, 300, 2061. [a] A. ROUX, G. CUPPELLO, J. CARTAUD, J. PROST, B. GOUD, P. BASSEREAU, Proc. Natl. Acad. Sci. USA, 2002, 99, 5394–5399. 33 H. NOJI, R. YASUDA, M. YOSHIDA, K. KINOSITA, Jr., Nature 1997, 386, 299. 34 C. MONTEMAGNO, G. BACHAND, Nanotechnology 1999, 10, 225. 35 S. A. COHN, A. L. INGOLD, J. M. SCHOLEY, J. Biol. Chem. 1989, 264, 4290. 36 K. J. BOHM, P. STEINMETZER, A. DANIEL, M. BAUM, W. VATER, E. UNGER, Cell. Motil. Cytoskeleton 1997, 37, 226. 37 K. J. BOHM, R. STRACKE, E. UNGER, Cell Biol. Int. 2000, 24, 335. 38 K. KAWAGUCHI, S. ISHIWATA, Cell. Motil. Cytoskeleton 2001, 49, 41. 39 K. KAWAGUCHI, S. ISHIWATA, Biochem. Biophys. Res. Commun. 2000, 272, 895. 40 B. J. SCHNAPP, B. CRISE, M. P. SHEETZ, T. S. REESE, S. KHAN, Proc. Natl. Acad. Sci. USA 1990, 87, 10053. 41 R. SAKOWICZ, M. S. BERDELIS, K. RAY, C. L. BLACKBURN, C. HOPMANN, D. J. FAULKNER, L. S. GOLDSTEIN, Science 1998, 280, 292. 42 T. U. MAYER, T. M. KAPOOR, S. J. HAGGARTY, R. W. KING, S. L. SCHREIBER, T. J. MITCHISON, Science 1999, 286, 971. 43 J. R. SELLERS, H. V. GOODSON, Protein Profile 1995, 2, 1323. 44 D. L. COY, W. O. HANCOCK, M. WAGENBACH, J. HOWARD, Nature Cell Biol. 1999, 1, 288. 45 H. LIU, J. J. SCHMIDT, G. D. BACHAND, S. S. RIZK, L. L. LOOGER, H. W. HELLINGA, C. D. MONTEMAGNO, Nature Mater 2002, 1, 173.

15

16 Biomolecular Motors Operating in Engineered Environments

46 M. J. DECASTRO, C. H. HO, R. J. STEWART, Biochemistry 1999, 38, 5076. 47 H. FELGNER, R. FRANK, J. BIERNAT, E. M. MANDELKOW, E. MANDELKOW, B. LUDIN, A. MATUS, M. SCHLIWA, J. Cell Biol. 1997, 138, 1067. 48 J. A. SPUDICH, S. J. KRON, M. P. SHEETZ, Nature 1985, 315, 584. 49 L. LIMBERIS, J. J. MAGDA, R. J. STEWART, Nano Lett. 2001, 1, 277. 50 L. LIMBERIS, R. J. STEWART, IOP Publishing. Nanotechnology 2000, 11, 47. 51 T. B. BROWN, W. O. HANCOCK, Nano Lett. 2002, 2, 1131. 52 D. TURNER, C. Y. CHANG, K. FANG, P. CUOMO, D. MURPHY, Anal. Biochem. 1996, 242, 20. 53 D. C. TURNER, C. Y. CHANG, K. FANG, S. L. BRANDOW, D. B. MURPHY, Biophys. J. 1995, 69, 2782. 54 Y. WADA, T. HAMASAKI, P. SATIR, Mol. Biol. Cell 2000, 11, 161. 55 C. VEIGEL, L. M. COLUCCIO, J. D. JONTES, J. C. SPARROW, R. A. MILLIGAN, J. E. MOLLOY, Nature 1999, 398, 530. 56 J. YAJIMA, M. C. ALONSO, R. A. CROSS, Y. Y. TOYOSHIMA, Curr. Biol. 2002, 12, 301. 57 F. GITTES, E. MEYHOFER, S. BAEK, J. HOWARD, Biophys. J. 1996, 70, 418. 58 E. BERLINER, H. K. MAHTANI, S. KARKI, L. F. CHU, J. E. CRONAN, Jr., J. GELLES, J. Biol. Chem. 1994, 269, 8610. 59 J. W. JARVIK, C. A. TELMER, Annu. Rev. Genet. 1998, 32, 601. 60 K. TERPE, Appl. Microbiol. Biotechnol. 2003, 60, 523. 61 J. P. BEARINGER, S. TERRETTAZ, R. MICHEL, N. TIRELLI, H. VOGEL, M. TEXTOR, J. A. HUBBELL, Nature Mater. 2003, 2, 259. 62 F. JIANG, K. KHAIRY, K. POOLE, J. HOWARD, D. MULLER, submitted 2003. 63 A. HAZARD, C. MONTEMAGNO, Arch. Biochem. Biophys. 2002, 407, 117. 64 A. K. SCHMID, N. C. BARTELT, R. Q. HWANG, Science 2000, 290, 1561. 65 M. DOGTEROM, B. YURKE, Science 1997, 278, 856. 66 F. van den ENT, J. MOLLER-JENSEN, L. A. AMOS, K. GERDES, J. LOWE, EMBO J. 2002, 21, 6935. 67 J. LOWE, L. A. AMOS, Nature 1998, 391, 203.

68 F. J. NEDELEC, T. SURREY, A. C. MAGGS, S. LEIBLER, Nature 1997, 389, 305. 69 T. SURREY, M. B. ELOWITZ, P. E. WOLF, F. YANG, F. NEDELEC, K. SHOKAT, S. LEIBLER, Proc. Natl. Acad. Sci. USA 1998, 95, 4293. 70 K. KRUSE, F. JULICHER, Phys. Rev. Lett. 2000, 85, 1778. 71 D. HUMPHREY, C. DUGGAN, D. SAHA, D. SMITH, J. KAS, Nature 2002, 416, 413. 72 W. S. RYU, R. M. BERRY, H. C. BERG, Nature 2000, 403, 444. 73 D. ISHII, K. KINBARA, Y. ISHIDA, N. ISHII, M. OKOCHI, M. YOHDA, T. AIDA, Nature 2003, 423, 628. 74 N. C. SEEMAN, Nature 2003, 421, 427. 75 S. R. HAMEROFF, R. C. WATT, J. Theoret. Biol. 1982, 98, 549. 76 R. PENROSE, Ann. N. Y. Acad. Sci. 2001, 929, 105. 77 P. SHETERLINE, J. CLAYTON, J. SPARROW, Protein Profile 1995, 2, 1. 78 E. NOGALES, Annu. Rev. Biochem. 2000, 69, 277. 79 C. BUSTAMANTE, Z. BRYANT, S. B. SMITH, Nature 2003, 421, 423. 80 P. J. HAGERMAN, Annu. Rev. Biophys. Biomol. Struct. 1997, 26, 139. 81 C. RUEGG, C. VEIGEL, J. E. MOLLOY, S. SCHMITZ, J. C. SPARROW, R. H. FINK, News Physiol. Sci. 2002, 17, 213. 82 A. MEHTA, J. Cell Sci. 2001, 114, 1981. 83 F. J. KULL, Essays Biochem. 2000, 35, 61. 84 C. SHINGYOJI, H. HIGUCHI, M. YOSHIMURA, E. KATAYAMA, T. YANAGIDA, Nature 1998, 393, 711. References 199 85 S. A. BURGESS, M. L. WALKER, H. SAKAKIBARA, P. J. KNIGHT, K. OIWA, Nature 2003, 421, 715. 86 G. J. WUITE, S. B. SMITH, M. YOUNG, D. KELLER, C. BUSTAMANTE, Nature 2000, 404, 103. 87 M. D. WANG, M. J. SCHNITZER, H. YIN, R. LANDICK, J. GELLES, S. M. BLOCK, Science 1998, 282, 902. 88 N. R. FORDE, D. IZHAKY, G. R. WOODCOCK, G. J. WUITE, C. BUSTAMANTE, Proc. Natl. Acad. Sci. USA 2002, 99, 11682. 89 J. J. CHAMPOUX, Annu. Rev. Biochem. 2001, 70, 369. 90 T. R. STRICK, G. CHARVIN, N. H. DEKKER, J. F. ALLEMAND, D. BENSIMON, V. CROQUETTE, Comptes Rendus Physique 2002, 3, 595.

References 91 D. E. SMITH, S. J. TANS, S. B. SMITH, S. GRIMES, D. L. ANDERSON, C. BUSTAMANTE, Nature 2001, 413, 748. 92 A. J. MERZ, M. SO, M. P. SHEETZ, Nature 2000, 407, 98.

93 B. MAIER, L. POTTER, M. SO, H. S. SEIFERT, M. P. SHEETZ, Proc. Natl. Acad. Sci. USA 2002, 99, 16012. 94 D. J. DEROSIER, Cell 1998, 93, 17.

17

1

Biocatalysis in Non-conventional Media Andreas Sebastian Bommarius

Georgia Institute of Technology, Atlanta, USA

Bettina R. Riebel Emory University School of Medicine, Atlanta, USA

Originally published in: Biocatalysis. Andreas Sebastian Bommarius and Bettina R. Riebel. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30344-1

Introduction

Contrary to expectations that enzymes are only active in aqueous solution, activity in almost anhydrous organic solvents was already demonstrated in the 1930s and rediscovered in 1977. It was not water-miscible hydrophilic solvents such as methanol or acetone that proved to be the best reaction media, but hydrophobic water-immiscible solvents such as toluene or cyclohexane. Supposedly, the cause is the partitioning of water between the enzyme surface and the bulk phase of the organic solvent. As comparably hydrophilic solvents such as methanol or acetone can take up basically inﬁnite amounts of water, they strip the remaining water molecules off the enzyme surface. As a consequence, the enzyme is no longer active because it requires a small but measurable amount of water for developing its activity. The advantages of organic media for enzyme reactions are: (i) an enhancement of the solubility of reactants; (ii) a shift of equilibria in organic media; (iii) easier separation of organic solvents than water; (iv) enhanced stability of enzymes in organic solvents; and (v) altered selectivity, including substrate speciﬁcity, enantioselectivity, prochiral selectivity, regioselectivity, and chemoselectivity, of enzymes in organic solvents. In water (with no organic phase present) the yield of the ester is about 0.01%, whereas in a biphasic water–water-immiscible organic solvent system consisting of porous glass impregnated with aqueous buffer solution it is practically 100%. Upon dehydration, many enzymes become extremely thermostable, up to and beyond 100◦ C. Most categories of enzymes have been found to be active in organic solvents, not just lipases but also proteases, dehydrogenases, peroxidases, and several others. Enzymatic reactions in organic solvents follow saturation kinetics; mechanisms Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Biocatalysis in Non-conventional Media

and active sites have been found to be the same as in aqueous solution. Kinetic constants, however, do not correlate with a few simple solvent parameters such as hydrophobicity, dielectric constant, or dipole moment; instead, case-by-case correlations are found. While enzymes in organic solvents often display activities that are orders of magnitude of lower than in water, there are several activation mechanisms which each yield about an order of magnitude of improvement: (i) sufﬁcient hydration in organic solvent (between 1% v/v and saturation); (ii) lyophilization at the pH of maximum activity in water; (iii) addition of lyoprotectants such as polyols; (iv) addition of hydrophobic binding pocket protectors such as phenols or anilines; and (v) lyophilization in strong salts such as KCl (if present at > 90% w/w). Whereas “protein engineering” covers knowledge of improving the biocatalyst, the complementary “medium engineering” optimizes the reaction medium, resulting in improved prochiral, regio-, and enantioselectivity. Many such changes have been observed, demonstrating that not only the catalyst but also the medium exerts a decisive inﬂuence on the outcome of an enzymatic reaction.

1 Enzymes in Organic Solvents

Conventional wisdom says that enzymes are active in water and in water only, just as in nature; according to these ideas, enzymes immediately deactivate in organic solvents. This picture is wrong for several reasons: Ĺ in nature, enzymes are not active in water but in the proximity of cell membranes, so a double layer of lipid molecules or a micelle would be a more suitable environment to emulate than an aqueous bulk phase; Ĺ enzymes do not necessarily deactivate in organic solvents – other phenomena such as inaccessibility owing to insolubility or denaturation during lyophilization can account for lack of activity.

Despite these arguments and classic papers demonstrating activity of enzymes in partially or wholly organic solvents [1], results from the Lomonosov University in Moscow, Russia, at the end of the 1970s came somewhat as a surprise: Klibanov et al. demonstrated esteriﬁcation of N-acetyl-L-tryptophan with ethanol in chloroform [2]. As porous glass was impregnated with aqueous buffer solution, Klibanov et al. termed this system a biphasic water–water-immiscible organic solvent system. They considered the system to consist of two phases, the organic phase and the aqueous phase on the glass bead, noted the shift of equilibria for water-forming reactions in nearly anhydrous media and provided equations modeling the situation. In water (no organic phase) the yield of the ester is about 0.01%, whereas in the biphasic system it is practically 100%. However, it was not water-miscible hydrophilic solvents such as methanol or acetone that proved to be the best reaction media but, on the contrary, hydrophobic

2 Evidence for the Perceived Advantages of Biocatalysts in Organic Media 3

water-immiscible solvents such as toluene or cyclohexane. Supposedly, the cause is the partitioning of water between the enzyme surface and the bulk phase of the organic solvent. As comparably hydrophilic solvents such as methanol or acetone can take up basically inﬁnite amounts of water, they strip the remaining water molecules off the enzyme surface. As a consequence, the enzyme is no longer active because it requires a small but measurable amount of water for developing its activity. What are the advantages of organic solvents as media for biocatalytic reactions? 1. The main advantage is an enhanced solubility of many, mostly hydrophobic, reactants insoluble or only sparingly soluble in water. Even for reactants with a moderate solubility in water, that solubility can often limit the attainable reaction rate. 2. In nearly anhydrous media, equilibria of hydrolytic reactions can be shifted towards condensation. In this fashion, reactions are possible in organic media whose equilibrium constants favor the reactants to such a degree that they alone are present in water. 3. As the enzyme in most cases is not dissolved in hydrophobic media but only suspended, it can be separated relatively easily after the reaction is ﬁnished. As the enzyme is not immobilized, connected disadvantages such as low activity yields or mass transfer limitations are avoided. 4. Enzymes are sometimes more stable in anhydrous organic solvents than in water. Lysozyme in water at 100◦ C was demonstrated to be 50% deactivated after 30 s (pH 8) or after 100 min (pH 4), but after 140 h in cyclohexane or even after 200 h as a dry powder [3]. 5. Owing to the inﬂuence of the medium, enzymes should feature different substrate speciﬁcities and selectivities than water. This would allow for tuning of these parameters under the control of the operator. 6. Organic solvents can be recycled more easily than water with its known high enthalpy of vaporization.

2 Evidence for the Perceived Advantages of Biocatalysts in Organic Media 2.1 Advantage 1: Enhancement of Solubility of Reactants

The rates of asymmetric sulfoxidation of thioanisole in nearly anhydrous (99.7%) isopropyl alcohol and methanol catalyzed by horseradish peroxidase (HRP) were determined to be tens to hundreds of times faster than in water under otherwise identical conditions [4]. Similar effects were observed with other hemoproteins. This dramatic activation is due to a much higher substrate solubility in organic solvents than in water and occurs even though the intrinsic reactivity of HRP in isopropyl alcohol and in methanol is hundreds of times lower than in water.

4 Biocatalysis in Non-conventional Media

In addition, the rates of spontaneous oxidation of the model prochiral substrate thioanisole in several organic solvents was observed to be some 100- to 1000-fold slower than in water. This renders peroxidase-catalyzed asymmetric sulfoxidations synthetically attractive. 2.2 Advantage 2: Shift of Equilibria in Organic Media 2.2.1 Biphasic Reactors Multi-phase reactors are of interest in biocatalytic reactions if one or several components of the reaction are insoluble or insufﬁciently soluble in aqueous phases but if an aqueous phase has to be kept, if only for the biocatalyst. However, a two-phase system can be utilized advantageously for the shifting of an equilibrium; this is demonstrated below. We analyze the simple reaction A ⇐⇒ B in an organic-aqueous two-phase system with the assumption that reactant A and product B partition between the two phases. The partition coefﬁcients Pw and Porg are deﬁned by Eq. (1).

PX = [X]org /[X]w

(1)

In both phases, equilibrium exists between A and B [Eq. (2)]. K w = [B]w /[A]w and K org = [B]org /[A]org

(2)

A biphasic equilibirum constant also can be deﬁned as in Eq. (3). K biphas = [B]tot /[A]tot

(3)

[X]tot can be linked to [X]org and [X]w via a mass balance [Eq. (4)]. [X]tot Vtot = [X]w Vw + [X]org Vorg

(4)

In addition, the sum V tot = V w + V org is valid, so insertion of Eq. (4) into Eq. (3) yields Eq. (5). K biphas = {[B]w Vw + [B]org Vorg {/{[A]w Vw + [A]org Vorg }

(5)

Upon division by [A]w V w and introduction of the phase ratio α (α = V org /V w ) we ﬁnally obtain Eq.: K biphas = K w · {(1 + α PB )/(1 + α PA )}

(6)

It can be seen from this equation that the effective equilibrium constant K bisphas is a function of the phase ratio α. Within limits, the equilibrium can be shifted in the

2 Evidence for the Perceived Advantages of Biocatalysts in Organic Media 5

Fig. 1 Dependence of the effective equilibrium constant K biphas on the phase ratio " (from Chaplin, 1990).

desired direction by controlling and shifting α. If the reaction is of the form A + B ⇐⇒ C + D, Eq. (7) is equivalent to Eq. (6). K biphas = K w · {(1 + α PC )(1 + α PD )/(1 + α PA )(1 + α PB )}

(7)

Figure 1 further elucidates the connection between K biphas and α: 2.3 Advantage 3: Easier Separation

In comparison to the high enthalpy of evaporation of water (57.36 kJ mol−1 ), the respective values are much lower for organic solvents, which are thus much easier to separate than water. Table 1 lists some boiling points and enthalpies of evaporation for common organic solvents. 2.4 Advantage 4: Enhanced Stability of Enzymes in Organic Solvents

Porcine pancreatic lipase catalyzes the transesteriﬁcation reaction between tributyrin and various primary and secondary alcohols in a 99% organic medium [5]. Upon further dehydration, the enzyme becomes extremely thermostable. Not only

6 Biocatalysis in Non-conventional Media Table 1 Boiling points and enthalpies of evaporation for common organic solvents Solvent MTBE Acetone CH3 OH THF Hexane DIPE B.p. [◦ C] 55.2 29.70 Hvap [kJ mol−1 ]

56.2 32.00

65.0 39.26

67.0 28.76

69.0 31.93

EA

C2 H5 OH C6 H12 Toluene DMF

69.0 77.1 32.56 34.75

78.5 40.50

80.7 32.79

110.6 149 39.23 60.45

MTBE: methyl t-butyl ether; THF: tetrahydrofuran; EA: ethyl acetate; C6 H12: cyclohexane; DIPE: diisopropyl ether; DMF: dimethylformamide.

can the dry lipase withstand heating at 100 ◦ C for many hours, but it exhibits a high catalytic activity at that temperature. Reduction in water content also alters the substrate speciﬁcity of the lipase: in contrast to its wet counterpart, the dry enzyme does not react with bulky tertiary alcohols. During transesteriﬁcation of amino acid esters with alcohols in the presence of chymotrypsin and subtilisin, both thermal and storage stabilities of chymotrypsin in non-aqueous solvents are observed to be greatly enhanced compared to water. 2.5 Advantage 5: Altered Selectivity of Enzymes in Organic Solvents

One of the most profound inﬂuences a solvent system can have on a reaction is a change of selectivity. Enzymes in organic solvents have been discovered in many cases to feature altered selectivity, including substrate speciﬁcity, enantioselectivity, prochiral selectivity, regioselectivity, and chemoselectivity [6, 7].

3 State of Knowledge of Functioning of Enzymes in Solvents 3.1 Range of Enzymes, Reactions, and Solvents

Not only do enzymes work in anhydrous organic media, but in this unnatural milieu they acquire remarkable properties such as enhanced stability, altered substrate and enantiomeric speciﬁcities, molecular memory, and the ability to catalyze unusual reactions [8]. Regarding the latter point, hydrolases, such as lipases, catalyze not only transesteriﬁcations in organic media but also other types of reactions, including esteriﬁcation, aminolysis, thiotransesteriﬁcation, and oximolysis. As all of these reactions compete with hydrolyses, which tend to dominate in aqueous media, some of them proceed to an appreciable extent only in non-aqueous solvents. Soon after the initial discovery, it became apparent that neither the source of the enzyme, nor the type of enzyme, nor the type of solvent seem to constrain the

3 State of Knowledge of Functioning of Enzymes in Solvents 7 Table 2 A selection of enzymes active in organic solvents

Class

E.C. number Enzyme

Reaction

Reference

yeast ADH horse-liver ADH alcohol oxidase HR peroxidase PP lipase

ethanol oxidation alcohol oxidation ethanol oxidation sulfoxidation transesteriﬁcation

Deetz [17] Zaks [19] Zaks [19] Dai [4] Zaks [5]

3.2.1.17 3.4.21.1 3.4.21.62 3.4.21.4 3.4.24.27 3.5.1.5 3.5.1.11

lysozyme α-chymotrypsin subtilisin trypsin thermolysin urease Pen G acylase

transesteriﬁcation transesteriﬁcation transesteriﬁcation dipeptide synthesis hydrolysis of urea resolution of β-amino esters

Zaks [3] Zaks [16] many van Unen [36] Bedell [4] Ghatorae [29] Roche [67]

4.1.2.10 4.2.1.84

(R)-oxynitrilase HCN addition to C O nitrile hydratase H2 O addition to nitriles

Oxido-reductases 1.1.1.1 1.1.1.1 1.1.3.13 1.11.1. Hydrolases 3.1.1.3

Lyases

Wajant [80] Thomas [29]

HR peroxidase: horseradish peroxidase; PP lipase: porcine pancreatic lipase.

use of organic solvents [3]. Various types of enzymes, such as lipases, proteases (chymotrypsin, subtilisin), oxidoreductases (alcohol dehydrogenase, oxidases, and peroxidases), and others, react in organic solvents. A selection of enzymes found to be active is listed in Table 2. Lipases from three different sources, porcine pancreatic, yeast, and mold, act vigorously as catalysts in a number of nearly anhydrous organic solvents. The range of solvents encompasses the whole spectrum from hydrophilic ones such as methanol, isopropanol, and acetonitrile via THF to the hydrophobic ones such as dialkyl ethers, toluene, hexane, cyclohexane, and heptane. 3.2 The Importance of Water in Enzyme Reactions in Organic Solvents 3.2.1 Exchange of Water Molecules between Enzyme Surface and Bulk Organic Solvent Early after the (re)discovery of enzyme activity in organic solvents, the higher level of activity in very hydrophobic solvents, such as hydrocarbons, in comparison with rather hydrophilic solvents, such as alcohols, was noted. Soon the hypothesis was raised that water molecules partition between the enzyme surface and the bulk organic solvent, and they are “stripped off” the enzyme surface in dry organic solvents. This idea explained why hydrophilic solvents, which strip water off an enzyme surface more readily than hydrophobic ones, deactivate enzymes much more readily than the latter. The hypothesis was proven correct by exchange of water on the enzyme surface by tritiated water, T2 O: suspension of the tritiated enzyme in the organic solvent, ﬁltration of the enzyme, and measurement of the

8 Biocatalysis in Non-conventional Media

amount of tritiated water in the solvent by scintillation counting demonstrated that several different enzymes (α-chymotrypsin, Subtilisin Carlsberg, and horseradish peroxidase) yielded water molecules immediately after suspension in the solvent. Depending on the solubility of water in the solvent, different amounts of T2 O were desorbed, from 0.4% in hexane to 62% in methanol [9]. The amount of water required by chymotrypsin and subtilisin for catalysis in organic solvents is several hundred molecules per protein molecule, less than required to form a monolayer on the surface. While subtilisin and α-chymotrypsin act as catalysts in a variety of dry organic solvents, the vastly different catalytic activities in these organic solvents are partly due to stripping of the essential water from the enzyme by more hydrophilic solvents and partly due to the solvent directly affecting the enzymatic process. Enzymes as different as yeast alcohol oxidase, mushroom polyphenol oxidase, and horse-liver alcohol dehydrogenase demonstrated vastly increased enzymatic activity in several different solvents upon an increase in the water content, which always remained below the solubility limit [10]. While much less water was required for maximal activity in hydrophobic than in hydrophilic solvents, relative saturation seems to be most relevant to determining the level of catalytic activity. Correspondingly, miscibility of a solvent with water is not a decisive criterion: upon transition from a monophasic to a biphasic solvent system, no signiﬁcant change in activity level was observed [11]. Therefore, the level of water essential for activity depends more on the solvent than on the enzyme.

3.2.2 Relevance of Water Activity While water level itself seems to be of no signiﬁcance for enzyme activity in organic solvents, the relative amount of water with respect to total water solubility seems to be very inﬂuential. Correspondingly, instead of using concentration cw or mole fraction xw of water as a measure of the amount of water, water activity aw should be preferred. [Activity, linked to concentration by the activity coefﬁcient γ w , aw = γ w · cw , written here for the case of water, is the thermodynamically rigorous way to consider amounts per volume in thermodynamic formulae such as mass action laws.] At the solubility limit in any solvent, water activity is always equal to unity, regardless of the mole fraction. The transesteriﬁcation of vinyl butyrate with n-octanol with Pseudomonas cepacia lipase yielded increasing K M values with rising water activity; however, vmax values demonstrated a maximum at water activity levels aw between 0.11 and 0.38 [7]. Activity maxima at certain aw values have also been found with many other enzymes, such as optima at aw = 0.55 for Mucor miehei lipase-catalyzed reaction in several solvents of different polarity from hexane to 3-pentanone [13]. In comparing ﬁve different lipases, considerable variations not just of the optima of aw but also the dependence of activity on aw itself were found [14]. Moreover, in polar solvents, correlations of activity with aw often cannot be found at all, probably owing to solvent effects beyond those controlling hydration of the enzyme [5].

3 State of Knowledge of Functioning of Enzymes in Solvents 9

Summarizing the ﬁndings above, control of water activity aw during enzyme reactions in organic solvents is extremely important, as water activity exerts a crucial inﬂuence, and enzyme reactivity crucially depends on it. 3.3 Physical Organic Chemistry of Enzymes in Organic Solvents 3.3.1 Active Site and Mechanism In organic solvents, enzymes react at the same active site as in water; covalent modiﬁcation of the active center of the enzymes by a site-speciﬁc reagent renders them catalytically inactive in organic solvents [16]. Upon replacement of water with octane as the reaction medium, the speciﬁcity of chymotrypsin towards competitive inhibitors reverses. Early results with enzymes in organic solvents demonstrated that enzymes work best in organic solvents if there are lyophilized from aqueous solution at or near the optimum pH value in water (see Section 3, below). Any speciﬁc pH effect of organic solvents, however, can be dismissed: it was found that the pKa of amino, carboxylic, and phenolic compounds does not differ by more than 0.3 units in the aqueous and lyophilized states [13]. Enzymatic transesteriﬁcations in organic solvents follow Michaelis-Menten kinetics, and in quite a few cases, the values of v/K M roughly correlate with the solvent’s hydrophobicity [16]. The dependence of the catalytic activity of the enzyme in organic media on the pH of the aqueous solution from which it was recovered is bell-shaped, with the maximum coinciding with the pH optimum of the enzymatic activity in water. Hammett analysis and ﬁnding of corresponding LFERs suggests that the mechanism of an enzyme reaction in organic solvents is the same as in aqueous solution [18]. 3.3.2 Flexibility of Enzymes in Organic Solvents From experiments on (i) replacement of water with other hydrogen bond forming additives and (ii) titration of enzyme amino groups in an organic medium, as well as the literature data on dehydrated enzymes, it is concluded that the water required by enzymes in non-aqueous solvents provides them with sufﬁcient conformational ﬂexibility for catalysis [19]. The activity of lyophilized HL-ADH in several solvents increased by orders of magnitude upon addition of small amounts of water to solvents of dielectric constant ε from 1.9 to 36 [20]. Enzyme ﬂexibility, as measured by electron spin resonance (ESR) spectroscopy with a spin label at the active site, did not depend on water content but on ε of the pure solvent: HL-ADH turns less ﬂexible with decreasing ε. Correlating positively with the hydrophobicity of the solvent, different fractions of inactivated active centers were measured in different solvents with solid-state NMR spectroscopy (13 C-cross-polarization/magic angle spinning (MAS) NMR) [9]. Just as with tritiated water (see above), immediate desorption of water molecules from the protein surface was observed after addition to the organic solvent.

10 Biocatalysis in Non-conventional Media

In some instances, a relationship between enzyme enantioselectivity and ﬂexibility has been found: while there are some positive examples in the literature [8], other investigations either found a slightly negative correlation or did not ﬁnd any correlation at all [23]. 3.3.3 Polarity and Hydrophobicity of Transition State and Binding Site As enzyme active sites commonly have to be totally anhydrous to allow chemical catalysis to occur, substrates have to be desolvated when transferring from solvent to active site. The effect of six solvents ranging from water to hexane on the standard test reaction in organic solvents, the transesteriﬁcation of N-acetyl-L-Phe-OEt with n-PrOH, with the help of Subtilisin Carlsberg, has been interpreted by taking into account substrate desolvation, as measured by relative substrate solubility [24]. Analysis of a thermodynamic cycle between ground (E + S) and transition state (ES= ) in the solvent and in acetone yields Eq. (8). =

=

=

G obs = G intr + G s = G ES −G E

(8)

Equation (8) states that the apparent and intrinsic differential activation energies G= obs and G*intr differ by the differential Gibbs free solvation energy Gs , with Gs relating the saturation solubilities of N-Ac-L-Phe-OEt in a solvent to acetone as the standard state [Eq. (9)], so that, with G = H – T · S, Eq. (10) follows. G s = −RT ln([S]solvent /[S]acetone ) G

=

=

obs

= H

intr −T

(9) =

· S

intr

−RT ln([S]solvent /[S]acetone )

(10)

While G= intr failed to yield a straight-line correlation, an LFER, with log ε (manifesting solvent polarity) and G= obs in fact did feature such a correlation. The results indicate that substrate desolvation is important and that the intrinsic activation energy of subtilisin catalysis is lowest in polar organic solvents, which may be due to transition-state stabilization of the enzyme’s polar transition state for transesteriﬁcation. Furthermore, the results strongly point towards the same transition state in water and organic solvents across a wider range of solvent polarity ε. The dramatic activation of serine proteases in non-aqueous media owing to prior lyophilization in the presence of KCl could be due in principle to the relaxation of potential substrate diffusional limitations in clumps of heterogeneous, undissolved enzyme in organic solvent. This hypothesis was checked with a classic experiment for diffusional limitation: Subtilisin Carlsberg was lyophilized with KCl and phosphate buffer in varying proportions ranging from 1 to 99% (w/w) enzyme in the ﬁnal lyophilized solids [4]. The active enzyme content of a given biocatalyst preparation was controlled by mixing native subtilisin with subtilisin preinactivated with the serine protease inhibitor PMSF and lyophilizing the enzyme mixture in the presence of different fractions of KCl and phosphate buffer. Initial catalytic rates, normalized per weight of total enzyme in the catalyst material, were measured as

3 State of Knowledge of Functioning of Enzymes in Solvents 11

a function of active enzyme for biocatalyst preparations containing different ratios of active to inactive enzyme. Plots of initial reaction rates as a function of percentage active subtilisin in the biocatalyst were found to be linear for all biocatalyst preparations. Thus, enzyme activation, as high as 3750-fold in hexane for the transesteriﬁcation of N-Ac-L-PheOEt with n-PrOH, is a manifestation of intrinsic enzyme activation and not of relaxation of diffusional limitations resulting from diluted enzyme preparations. Thus, the above-mentioned hypothesis was refuted. As similar results were found for the metalloprotease thermolysin, activation due to lyophilization in the presence of KCl may be a general phenomenon. 3.4 Correlation of Enzyme Performance with Solvent Parameters

Given the multitude of potential organic solvents for enzyme reactions, design rules or at least heuristics were sought early on for picking organic solvents without trying a large number of them experimentally. Organic solvents can be described and classiﬁed according to several criteria, such as volatility (melting and boiling points, Tm and T boil ), size (molar mass, M [g mol−1 ]), accommodation of charge (dielectric constant ε), hydrophobicity (water/octanol partition coefﬁcient, log P), distribution of charge (dipole moment µ), or polarizability (polarization constant α); for data, see Table 3. The water/octanol partition coefﬁcient log P of a component A is deﬁned by Eq. (11), where A is in dilute solution below the solubility limit. logP = log{[A]water /[A]n−octanol }

(11)

n-Octanol is chosen as a reference because it mimics biological membranes with a hydrophobic tail and a hydrophilic head capable of hydrogen bonding.

Table 3 Some properties of common organic solvents Solvent MTBE Acetone CH3 OH THF Hexane DIPE B.p. [◦ C] M [g/mol] ε[−] log P µ [D]

55.2 88.15 4.5 1.15 1.23

56.2 65.0 67.0 58.08 32.04 72.10 20.7 32.63 7.58 − 0.23 − 0.76 0.49 2.70 1.71 1.74

EA

69.0 69.0 77.1 86.17 102.17 88.10 1.89 3.23 6.02 3.5 1.9 0.68 0.0 1.24 1.83

C2 H5 OH C6 H12 Toluene DMF 78.5 46.07 24.3 − 0.24 1.74

80.7 110.6 153 84.16 92.13 73.09 2.02 2.38 36.7 3.2 2.5 − 1.0 0.0 0.30 3.24

MTBE: methyl t-butyl ether; THF: tetrahydrofuran; EA: ethyl acetate; C6 H12 : cyclohexane; DIPE: diisopropyl ether; DMF: dimethylformamide.

12 Biocatalysis in Non-conventional Media

Fig. 2 Correlation of activity data with solvent parameters [27].

3.4.1 Control through Variation of Hydrophobocity: log P Concept In an inﬂuential early investigation, correlation of biocatalytic activity data of aerobic and anaerobic whole-cell biocatalysis with solvent properties resulted in the strongest correlation for the partition coefﬁcient log P, whereas both Hildebrand’s solubility parameter δ and the dielectric constant ε showed either a weak correlation with activity data or none at all [26, 27] (Figure 2). This simple picture, unfortunately, did not hold for all enzyme reactions in organic solvents. 3.4.2 Correlation of Enantioselectivity with Solvent Polarity and Hydrophobicity The enantioselectivity of the transesteriﬁcation of vinyl butyrate with secphenylethanol was inﬂuenced greatly by the solvent; however, not log P (hydrophobicity) but both the dielectric constant ε and the dipole moment µ of the solvent correlated best with the E values [27]. Table 4 demonstrates the correlation of µ with E. Table 4 Correlation of the enantioselectivity E of subtilisin in the transtesteriﬁcation of

racemic 1-phenylethanol in various organic solvents, and the dielectric constants ε of the solvents [28]

Solvent Dioxane Benzene Triethylamine Tetrahydrofuran pyridine

er

E

Solvent

2.2 2.3 2.4 7.6 12.9

61 54 48 40 31

Dimethylformamide Nitromethane Acetonitrile Methylacetamide

er

E

36.7 35.9 35.9 191.3

9 5 3 3

4 Optimal Handling of Enzymes in Organic Solvents 13 Table 5 Effects of the hydrophobicity (log P) of solvents on the prochiral selectivity (S) of

Pseudomonas sp. lipase in the monohydrolysis of 2-(1-naphthoylamino)trimethylene dibutyrate [29])

Solvent

log P

Acetonitrile Nitrobenzene Acetone Cyclohexanone Butanone 2-Pentanone Chloroform Tetrahydrofuran 2-Hexanone Dioxane t-Butyl acetate t-Butyl alcohol t-Amyl alcohol Triethylamine Toluene Benzene Carbon tetrachloride

−0.33 1.8 −0.23 0.96 0.29 0.80 2.0 0.49 1.3 −1.1 1.7 0.80 1.4 1.6 2.5 2.0 3.0

S >30 >30 18 18 16 16 9.9 9.1 8.8 5.4 5.3 4.9 4.8 4.7 3.5 3.2 2.6

In a third example, the prochiral selectivity results of the Pseudomonas sp. lipasecatalyzed monohydrolysis of 2-(1-naphthoylamino)trimethylene dibutyrate correlate inversely with hydrophobicity, as expressed by log P (Table 5; [29]). The three examples provided in this section demonstrate that each case has to be considered individually. None of the solvent parameters has been found to correlate reliably with enzyme activity or selectivity data, except in individual cases; predictions at this point are not possible.

4 Optimal Handling of Enzymes in Organic Solvents

While different enzyme reactions yield different results in different solvents, thus rendering comparison difﬁcult, a procedure akin to a “standard protocol” for conducting enzyme reactions in organic solvents has been developed. It calls for Ĺ lyophilization at optimum pH in aqueous solution; Ĺ meeting the requirement for water for enzyme function; and Ĺ testing the enzyme with a standard reaction under standard conditions.

In the case of subtilisin, the standard reaction is the transesteriﬁcation of Nacetyl-L-phenylalanine ethyl ester with n-propanol [Eq. (12)]. N−Ac−L−Phe−OEt + n−PrOH → NAc−L−Phe−OPr + EtOH

(12)

14 Biocatalysis in Non-conventional Media

More data have been obtained with this reaction on enzymes in organic solvents than with any other. 4.1 Enzyme Memory in Organic Solvents

Repeatedly, researchers have found the phenomenon that enzyme molecules seem to “remember” the aqueous conditions from which they were prepared [30]. As mentioned above, early results with enzymes in organic solvents demonstrated that enzymes work best in organic solvents if there are lyophilized from aqueous solution at or near the optimum pH value in water. No speciﬁc pH effect of organic solvents on enzyme reactions was found: the pK a of amino, carboxylic, and phenolic compounds does not differ by more than 0.3 units in the aqueous and lyophilized state [13]. Catalytic activities of α-chymotrypsin and Subtilisin Carlsberg in various hydrous organic solvents were measured as a function of how the enzyme suspension had been prepared [31]. Direct suspension of the lyophilized enzyme in the solvent containing 1% water was compared with precipitation of the same enzyme from its aqueous solution by a 100-fold dilution with anhydrous solvent. The reaction rate in a given non-aqueous enzymatic system was found to depend on the nature of both enzyme and solvent, but to depend strongly on the mode of enzyme preparation. The common mode of suspending lyophilized enzymes in organic solvents containing very little water results in just a low speciﬁc activity. Much higher speciﬁc activity can be achieved if the enzymes are dissolved in water ﬁrst and then diluted with anhydrous organic solvent to the same water content [14]; the lower the water content of the medium, the greater the discrepancy. The mechanism of this phenomenon, termed “lyophilization-induced inactivation”, was found to arise from reversible denaturation of the oxidases on lyophilization: because of its conformational rigidity, the denatured enzyme exhibits very limited activity when directly suspended in largely non-aqueous media but renatures and thus yields much higher activity if ﬁrst redissolved in water. Lyophilizationinduced inactivation could be minimized, at least for the case of four oxidases employed, by at least two strategies, both involving the addition of excipients to the aqueous enzyme solution before lyophilization: Ĺ addition of lyoprotectants such as polyols or polyethylene glycol to preserve the overall enzyme structure during dehydration; Ĺ addition of phenolic and aniline substrates that presumably bind to the hydrophobic pocket of the enzyme active site.

Not only was the effect of these excipients found to extend activity by up to orders of magnitude but the effect of the two excipient groups was found to be additive, affording up to complete protection against lyophilization-induced inactivation when representatives of the two are present together.

4 Optimal Handling of Enzymes in Organic Solvents 15

4.2 Low Activity in Organic Solvents Compared to Water

One important issue in the discussion of the biotechnological opportunities afforded by non-aqueous enzymology and its impact on biocatalysis is the often drastically diminished enzymatic activity in organic solvents compared to that in water. The loss of comparative speciﬁc activity can be as high as 103 −105 . Several recent studies have addressed the issue and have made great strides towards both elucidating causes of the phenomenon of activity loss and designing strategies that systematically enhance activity by multiple orders of magnitude [33]. The goal ultimately is to bring the activity level of enzymes in organic solvents to aqueous-like level. Dipeptide synthesis in acetonitrile is found to be enhanced 425-fold in the α-chymotrypsin-catalyzed reaction between the 2-chloroethyl ester of N-acetyl-Lphenylalanine and L-phenylalaninamide upon lyophilization of the enzyme in the presence of 50 equivalents of 18-crown-6 [34]. Acceleration is observed in different solvents and for various peptide precursors. The enhancement of enzyme activity upon addition of 18-crown-6 to the organic solvent can be reconciled with a mechanism in which macrocyclic interactions of 18-crown-6 with the enzyme play an important role [35]. Macrocyclic interactions (e.g., complexation with lysine ammonium groups of the enzyme) can lead to a reduced formation of inter- and intramolecular salt bridges and, consequently, to lowering of the kinetic conformational barriers, enabling the enzyme to refold into thermodynamically stable, catalytically (more) active conformations. This assumption is supported by the observation that the crown-ether-enhanced enzyme activity is retained after removal of the crown by washing with a dry organic solvent. Much stronger crown ether activation is observed when 18-crown-6 is added prior to lyophilization, and this can be explained by a combination of two effects: the beforementioned macrocyclic complexation effect, and a less speciﬁc, non-macrocyclic, lyoprotecting effect. The magnitude of the total crown ether effect depends on the polarity and thermodynamic water activity of the solvent, the activation being highest in dry and apolar media, where kinetic conformational barriers are highest. By determination of the speciﬁc activity of crown-ether-lyophilized enzyme as a function of the enzyme concentration, the macrocyclic crown ether (linearly dependent on the enzyme concentration) and the non-macrocyclic lyoprotection effect (not dependent on the enzyme concentration) could be separated. These measurements reveal that the contribution of the non-macrocyclic effect is signiﬁcantly larger than the macrocyclic refolding effect. Immobilization in a sol–gel matrix accelerated the propanolysis of N-acetyl-Lphenylalanine ethyl ester in cyclohexane for several serine proteases compared to the non-immobilized lyophilized enzymes: 31-fold for Subtilisin Carlsberg, 43-fold for α-chymotrypsin, and 437-fold for trypsin [36]. The activity yield upon immobilization was 90% (α-chymotrypsin). The rate enhancement effect of immobilization on the enzyme activity is highest in hydrophobic solvents.

16 Biocatalysis in Non-conventional Media Table 6 Approximate activation factors for enzymes in organic solvents for various strategies

Action

Approx. activation factor

Sufﬁcient hydration in organic solvent (≥ 1%, < saturation) Lyophilization at pH of maximum activity in water Lyoprotectants such as polyols Hydrophobic binding pocket protectors (phenols, anilines) Strong salts such as KCl (if present at > 90% w/w)

10 10 10 10 10

total:

105

The activation mechanisms, with the corresponding activation ratios by which the maximum attainable rate constants of enzymes in organic solvents can be enhanced, are summarized in Table 6. Activation by a factor of 105 closes most of the gaps between speciﬁc activity levels in water and organic solvents. The state of affairs regarding the preparation of highly active enzyme formulations for use in non-aqueous media is summarized by Lee and Dordick [37]. Improved mechanistic understanding of enzyme function and activation in dehydrated environments will lead to the development of a broad array of techniques for generating more active, stable, and enantioselective and regioselective tailored enzymes for synthetically relevant transformations. This, in turn, should result in an increase in the opportunities for enzymatic processes to be developed on a commercial scale. 4.3 Enhancement of Selectivity of Enzymes in Organic Solvents

Enzymatic enantioselectivity in organic solvents can be markedly enhanced by temporarily enlarging the substrate via salt formation [38]. In addition to its size, the stereochemistry of the counterion can greatly affect the enantioselectivity enhancement [39]. In the Pseudomonas cepacia lipase-catalyzed propanolysis of phenylalanine methyl ester (Phe-OMe) in anhydrous acetonitrile, the E value of 5.8 doubled when the Phe-OMe/(S)-mandelate salt was used as a substrate instead of the free ester, and rose sevenfold with (R)-mandelic acid as a Brønsted-Lewis acid. Similar effects were observed with other bulky, but not with petite, counterions. The greatest enhancement was afforded by 10-camphorsulfonic acid: the E value increased to 18 ± 2 for a salt with its R-enantiomer and jumped to 53 ± 4 for the S. These effects, also observed in other solvents, were explained by means of structure-based molecular modeling of the lipase-bound transition states of the substrate enantiomers and their diastereomeric salts. The relatively meager stereoselectivity of horseradish peroxidase in the sulfoxidation of thioanisole in 99.8% (v/v) methanol is vastly improved when the enzyme forms a complex with benzohydroxamic acid [16]. The generality of the observed “ligand-induced stereoselectivity enhancement” is demonstrated with other hydrophobic hydroxamic acids, as well as with additional thioether

5 Novel Reaction Media for Biocatalytic Transformations 17

substrates, and rationalized by means of molecular dynamics simulations and energy minimization.

5 Novel Reaction Media for Biocatalytic Transformations 5.1 Substrate as Solvent (Neat Substrates): Acrylamide from Acrylonitrile with Nitrile Hydratase

One of the best examples for discussing biotransformations in neat solvents is the enzymatic hydrolysis of acrylonitrile, a solvent, to acrylamide. For several applications of acrylamide, such as polymerization to polyacrylamide, very pure monomer is required, essentially free from anions and metals, which is difﬁcult to obtain through conventional routes. In Hideaki Yamada’s group (Kyoto University, Kyoto, Japan), an enzymatic process based on a nitrile hydratase was developed which is currently run on a commercial scale at around 30 000–40 000 tpy with resting cells of third-generation biocatalyst from Rhodococcus rhodochrous J1. Characteristic of this process are its extremely high concentration level and space-time-yield. Solid nitrile substrates render precipitating product up to a solid medium at high degrees of conversion; liquid nitriles can be run as neat substrate. Figure 3 illustrates the connection between substrate concentration up to 15 M (!) and achievable degree of conversion for the example of the transformation of 3-cyanopyridine to nicotinamide.

Fig. 3 Relationship of degree of conversion with substrate concentration for the reaction of cyanopyridine to form nicotinamide [64].

18 Biocatalysis in Non-conventional Media

5.2 Supercritical Solvents

Supercritical solvents combine extremely low viscosity as well as diffusivity of gases with the high solubility of compounds in solution. In addition, products can be separated by simply depressurizing the reaction vessel and boiling off the solvent. After having been tested for chemical reactions, supercritical solvents were also tested for enzymatic reactions in around 1985. Owing to the expected dependence of the reaction parameters on temperature and pressure, supercritical solvents should allow the investigation of that inﬂuence. Supercritical CO2 (SCCO2 ), especially, with its accessible critical parameters (T crit = 31◦ C, pcrit = 50 bar) offers access to this technique. However, results demonstrated that SCCO2 does not show good results for all reactions. It was found for the synthesis of acrylates by transesteriﬁcation of methylacrylate with lipase from Candida rugosa that owing to its hydrophilic character SCCO2 was inferior to more hydrophobic supercritical solvents such as propane, ethane, or even SCSF6 with respect to reactivity [41]. A comparison between SCCO2 and n-hexane for the esteriﬁcation of oleic acid showed that in SCCO2 inhibition by ethanol was less important but that vmax in nhexane was about an order of magnitude higher. Enzyme stability was comparable and satisfactory in both cases [27]. For a review of the ﬁeld of biocatalysis in supercritical solvents, see Mesiano [85]. 5.3 Ionic Liquids

Ionic liquids are composed of organic cations with a compatible anion; examples are [BMIM][PF6 ] and [MMIM][MeSO4 ] (Figure 4). Unlike conventional organic solvents, ionic liquids possess no vapor pressure but good solvent properties, and a wide range of thermal stability over a wide range of liquid state (≤ 300◦ C); also, they can be used to form two-phase systems with many solvents [42].

Fig. 4 Some ionic liquids [42]: a) 1-butyl-3-methylimidazolium hexaﬂuorophosphate, [BMIM][PF6 ]; b) 1-butyl-3-methylimidazolium bis((triﬂuoromethyl)sulfonyl)amide, [BMIM][CF3 SO2 ]2 N;

c) 1-methyl-3-methylimidazolium methylsulfate, [MMIM][MeSO4 ]; d) 1-butyl-3-methylimidazolium triﬂate, [BMIM][CF3 SO3 ]; e) 1-butyl-4-methylpyridium tetraﬂuoroborate, [MBPy][BF4 ].

5 Novel Reaction Media for Biocatalytic Transformations 19

Given these properties, they can extend the application of solvent engineering to biocatalytic reactions. Instead of just serving as a replacement for organic solvents, they often lead to improved process performance. The ﬁrst report on thermolysincatalyzed Z-aspartame synthesis from Z-L-aspartate and L-phenylalanine methyl ester · HCl in 1-butyl-3-methylimidazolium hexaﬂuorophosphate [BMIM][PF6 ] achieved an initial rate (1.2 nmol (min−1 (mg protein)−1 ) comparable to rates in organic solvents, and remarkable enzyme stability obviating immobilization [25]. Since then, a range of enzymatic reactions and whole-cell processes have achieved remarkable yields [42, 78], (enantio) selectivity [45, 46], or enzyme stability [42, 47]. 5.4 Emulsions [Manufacture of Phosphatidylglycerol (PG)]

Fatty acids are important raw materials which are gained from naturally occurring triglycerides. However, thermal processes to obtain fatty acids result in discoloration, so alternative processes have been sought. Lipases (E.C. 3.1.1.3.) catalyze the hydrolysis of lipids at an oil/water interface. In a membrane reactor, the enzymes were immobilized both on the side of the water phase of a hydrophobic membrane as well as on the side of the organic phase of a hydrophilic membrane. In both cases, no other means for stabilization of the emulsion at the membrane were required. The synthesis reaction to n-butyl oleate was achieved with lipase from Mucor miehei, which had been immobilized at the wall of a hollow ﬁber module. The degree of conversion reached 88%, but the substrate butanol decomposed the membrane before the enzyme was deactivated. Phosphatidylglycerol (PG), which is useful as a lung surfactant for the treatment of respiratory illnesses, can be obtained from phosphatidylcholine (PC) in a transphosphatidylation reaction [Eq. (13)] catalyzed by phospholipase D (PLD, E.C. 3.1.4.4.), with the hydrolysis to phosphatidic acid (PA) as a side reaction [Eq. (14)]. PC + glycerol → PG + choline

(13)

PC + H2 O → PA + choline

(14)

Commonly, the reaction is conducted in an emulsion stabilized by surfactants and containing the substrate (PC). In a microporous membrane reactor, PLD stability could be increased sevenfold by the addition of ethers to the chamber side. Additionally, no surfactants were required as the product could be separated in simple fashion; 20% product was formed, compared with 4% in the simple emulsion system. 5.5 Microemulsions

If the objective is to keep the enzyme active and stable in an aqueous phase but otherwise to use as much organic phase as possible, microemulsions are an option as

20 Biocatalysis in Non-conventional Media

Fig. 5 Microemulsion droplets and possible average location of enzyme molecules [59].

a reaction medium. In contrast to ordinary emulsions they are thermodynamically stable and, at a particle diameter of 1–20 nm, accommodate most often only one enzyme molecule (Figure 5). The microemulsion droplets communicate rapidly and exchange their contents through elastic collisions. The boundary between microemulsions and reversed micelles is not clearly delineated, and the two notions are often used interchangeably. Enzyme of almost all classes and structures have been solubilized in microemulsion systems and used for reactions [48]. Activity and stability are often comparable to values in aqueous media. Many substrates which cannot be made to react in water or in pure organic solvents such as hexane owing to lack of solubility can be brought to reaction in microemulsions. Whereas enzyme structure and mechanism do not seem to change upon transition from water to the microemulsion phase [6], partitioning effects often are very important. Besides an enhanced or diminished concentration of substrates in the vicinity of microemulsion droplets and thus of enzyme molecules, the effective pH values in the water pool of the droplets can be shifted in the presence of charged surfactants. Frequently, observed acceleration or deceleration effects on enzyme reactions can be explained with such partitioning effects [41]. 5.6 Liquid Crystals

As described above, microemulsions feature the disadvantage that the reaction product can be separated only with difﬁculty from other components of the system. This disadvantage can be compensated by employing highly viscous to solid liquidcrystalline phases which consist of the same components as microemulsions (oil, surfactant, water). Many enzymes display activity in several phases of a threecomponent mixture (Figure 6) [51]. At the beginning of the investigation a surfactant/water/oil phase diagram has to be prepared in case it has not been done already. Subsequently, one can generate the reaction medium by mixing corresponding amounts of components according to the phase diagram; in most cases, several phases will form. In cubic inverse phases, which are mechanically very stable, a series of enzymatic reactions have been conducted [51]. Often, stability of enzymes has been found to be signiﬁcantly

5 Novel Reaction Media for Biocatalytic Transformations 21

Fig. 6 Liquid-crystalline phases and catalytic activity of peroxidase in ternary water/surfactant/organic solvent phases [59]. Phases: a = normal micelles; b = reversed micelles; c = lamellar aggregates; d = hexagonal aggregates.

higher in such liquid-crystalline phases than in pure isotropic continuous phases and media. 5.7 Ice-Water Mixtures

Even more than 30 years ago, it had been found that hydroxylaminolysis of amino acid esters with the help of trypsin catalysis is accelerated in frozen aqueous phases at −23◦ C in comparison to liquid aqueous phases at 1◦ C [34]. For kinetically controlled peptide synthesis with different serine or cysteine proteases, strongly enhanced yields of di- and oligopeptides have been observed [53] (Table 7; Figure 7). The strength of nucleophiles seems to be much higher in ice/water phases than in the purely liquid phase, as even unprotected dipeptides render good yields and even unprotected amino acid render at least moderate yields (Table 7; Figure 7). The reverse action of a trypsin-free elastase isolated from porcine pancreas was studied in frozen aqueous systems and was found to catalyze peptide bond formation more effectively than in solution at room temperature [36]. The acceptance of free amino acids as nucleophilic amino components indicates a changed speciﬁcity of the endoprotease in frozen reaction mixtures. In elastase-catalyzed formation of Ser-, Ile-, and Val-X-bonds in frozen aqueous reaction mixtures, peptide yields obtained depended on the P1 amino acid and the acyl donor chain length. The highest yields have been found at temperatures between −20 and −10◦ C and at pH values where more than 90% of the nucleophile was present as free

22 Biocatalysis in Non-conventional Media Table 7 Yields of α-chymotrypsin-catalyzed peptide synthesis in water and ice using maleoyl-

l-tyrosine methyl ester as acyl donor and various peptides, amino acid amides, and amino acids as nucleophiles , [53] Peptide yield [%] Amino component of the reaction H-Gly-Ala-OH H-Gly-Gly-OH H-Gly-Gly-Gly-OH H-D-Leu-NH2 H-L-Leu-NH2 H-Arg-OH H-Lys-OH H-β-Ala-Gly-OH

25◦ C 5.8 2.6 5.1 9.9 79.1 <2 <2 <2

−25◦ C 94.7 95.4 91.2 73.0 91.8 32.7 43.8 78.8

∗ [amino component] = 50 mM (25 mm as free base); [Maleoyl-L-Tyr-OMe] = 2 mM; [α-chymotrypsin] = 0.15 mm at 25◦ C and 0.3 µm at −25◦ C.

base. Beyond a tenfold excess of nucleophile over electrophile, no further increase in yield could be observed. In a typical case, for the α-chymotrypsin (E.C. 3.4.21.1)catalyzed reaction of maleyl-Phe-OMe with H-Leu-NH2 investigated under a range of reaction conditions, a variety of results were demonstrated [28]:

Fig. 7 Left: dependence of the yield of the "-chymotrypsin-catalyzed synthesis of the peptides maleoyl-Tyr-d-Leu-NH2 (◦) and maleoyl-Tyr-$-Ala-Gly-OH () in ice upon the relative amount of nucleophile free base (for conditions, see Table 7). Right: inﬂuence of temperature on the yield of the "-

chymotrypsin-catalyzed peptide synthesis in ice using maleoyl-l-Tyr methyl ester as acyl donor and H-d-Leu-NH2 (◦), H-Gly-Gly-GlyOH (•), and H-Ala-Ala-OH () as amino components (for conditions, see Table 7) [53].

5 Novel Reaction Media for Biocatalytic Transformations 23

Ĺ the peptide yield is independent of the method of shock-freezing; Ĺ the optimal reaction temperature is 248–263 K (−25 to −10 ◦ C), and lower temperatures result in clearly retarded reactions; Ĺ addition of DMSO leads to decreasing peptide yields; and Ĺ peptide bond formation is catalyzed by the active enzyme, since unspeciﬁc protein surface catalysis gave no peptide yields at all.

The best explanation of the good results for peptide syntheses in ice-water mixtures are based on the freeze-concentration-model, which just provides for a volume-reducing function for the ice while the liquid aqueous part is still the only relevant phase for the reaction. All observed enhancements of reaction rate would then have to be attributed to an increase in effective concentration. 1 HNMR relaxation time measurements have been used to determine the amount of unfrozen water in partially frozen systems, thus quantifying the extent of the “freeze-concentration effect” [56]. Comparative studies in ice and at room temperature verify the importance of freeze-concentration which, however, is not sufﬁcient for a complete understanding of the observed effects. 5.8 High-Density Eutectic Suspensions

Enzymatic catalysis in heterogeneous eutectic mixtures, i.e., at comparable mole fractions of each substrate (in the case of peptide synthesis, electrophile and nucleophile) and water, offers advantages over conventional reactions conducted in organic solvents. Such beneﬁts include the avoidance of bulk solvents and the attainment of greatly improved productivities. A wide range of proteases have been shown to retain their catalytic activity in eutectic mixtures of substrates, and have been used to synthesize various model and bioactive oligopeptides [30]. In similar fashion, the results of an initial study of enzymatic catalysis in metastable supersaturated solutions of carbohydrates show that such solutions, formed in the presence of small amounts of water and alcohol as plasticizers, are sufﬁciently stable under ambient conditions to enable enzymatic transformations of substrates [58]. A partial phase diagram for a system consisting of glucose, water, and polyethylene glycol was constructed to identify the regions which are most suitable for biotransformations (Figure 8). It was conﬁrmed that the glass transition in this system occurred below the reaction temperature at any given composition of the constituent components. Several glycosidases were found to be catalytically active in this medium and the activity of β-glucosidase from almond was determined at several compositions of the reaction mixture and related to the corresponding regions of the phase diagram. The synthetic utility of the system was illustrated by glucosylation of several α,ω-alkyldiols, short-chain polyethylene glycols, and hydroxyalkyl and glyceryl monoacrylates. The inﬂuence of eutectic media on the kinetics and productivity of biocatalysts has yet to be fully elucidated. Syntheses in eutectic suspensions have been scaled up to the pilot scale in a rotating drum reactor. The bioactive peptide Nα-Cbz-L-Lys(N-Cbz)-Gly-L-Asp(OAll)-L-Glu(OAll)OEt was synthesized via a

24 Biocatalysis in Non-conventional Media

Fig. 8 Phase diagram for a binary eutectic system [30]. a) E: eutectic point; TA: melting point of component A; TB: melting point of component B; CE: eutectic composition. b) Phase diagram of the system N-Z-l-Tyr-OEt + l-Leu-NH2 .

Curve A: binary mixture of substrates in the absence of adjuvant; curve B: ternary mixture with 3% (w/w) water and 6% (w/w) ethanol; curve C: ternary phase diagram containing 10% (w/w) triethylene glycol dimethyl ether.

sequential N-to-C strategy in a heterogeneous solid-liquid mixture of the substrates in the presence of chymopapain and subtilisin as well as 16–20% (w/w) water and ethanol [31]. At substrate concentrations of around 1 M, yields of 67–74% per step at product concentrations of 0.36, 0.49, and 0.48 kg kg−1 were achieved. The corresponding space–time yields were between 0.30 and 0.64 kg (kg d)−1 and biocatalyst reuse provided productivities of 166–312 kg product (kg enzyme)−1 . 5.9 High-Density Salt Suspensions

A measure as simple as adding certain inorganic salts to aqueous enzyme solutions prior to lyophilization can result in dramatic activation of the dried powder in organic media relative to enzyme lyophilized without added salt [60]. Upon salt-induced lyophilization of Subtilisin Carlsberg, the optimum speciﬁcity constant Kcat /K M for transesteriﬁcation in hexane exceeded the value for saltfree enzyme 20 000-fold, substantially more than the previously reported enhancement of 3750-fold. As for pure enzyme, salt-activated enzyme seems to exhibit the greatest activity when lyophilized from a solution of pH equal to the pH for optimal activity in water. The activation ratio and active-site content in the presence of 98% KCl (1% phosphate buffer and 1% enzyme) are highly sensitive to the lyophilization time and water content of the sample, over a range of up to 13fold for Subtilisin Carlsberg and 11-fold for a lipase from Mucor javanicus. The activity and water content were found to correlate directly with the kosmotropicity of the activating salt (kosmotropic salts bind water molecules strongly relative to the strength of water–water interactions in bulk solution) [60]. Combinations of

5 Novel Reaction Media for Biocatalytic Transformations 25

kosmotropic salts with known lyoprotectants such as polyethylene glycol (PEG) and sugars did not yield an appreciably more active catalyst. However, the combination of the kosmotropic sodium acetate with the strongly buffering sodium carbonate activated the enzyme more than the individual additives alone. Enzyme activity was enhanced further by the addition of small amounts of water to the organic solvent. Substrate selectivity experiments suggest that a mechanism other than selective partitioning of substrate into the enzyme-salt matrix is responsible for salt-induced activation of enzymes in organic solvents. Under optimal conditions, the enzyme activity in hexane was improved over 27 000-fold relative to the salt-free enzyme, reaching a catalytic efﬁciency that was within one order of magnitude of kcat /K M for hydrolysis of the same substrate in aqueous buffer. 5.10 Solid-to-Solid Syntheses

When one is using proteases in a direct reversal of their normal hydrolytic function, the equilibrium position is very important in limiting the attainable yield in equilibrium-controlled enzymatic peptide synthesis. If both reactants and products are largely undissolved in the reaction medium as suspended solids, thermodynamic analysis of such a system shows the reaction will proceed until at least one reactant has dissolved completely, towards either products or reactants (“switchlike” behavior). In case of a favorable equilibrium for synthesis, the yield is maximized in the solvent of least solubility for the starting materials [37]. Thermolysin-catalyzed reactions of X-Phe-OH (X = formyl, Ac, Z) with Leu-NH2 yielded X-Phe-Leu-NH2 with equilibrium yields > 90% over a range of solvents. Some predictions, such as a linear decrease in yield with the reciprocal of the initial reactant concentrations, could be veriﬁed [37]. This approach enables high peptide yields in equilibrium-controlled peptide synthesis in high-density aqueous media with an equimolar supply of substrates. Scale-up to molar amounts veriﬁed the concepts as well as demonstrate the synthetic utility of this approach: Z-His-Phe-NH2 and Z-Asp-Phe-OMe, precursors for cyclo-[His-Phe-] and the low-calorie sweetener Aspartame, respectively, were synthesized in preparative yields of 84–88% [21]. For a review of the ﬁeld of peptide synthesis in unusual media, see Jakubke [40]. In the thermolysin-catalyzed solid-to-solid dipeptide synthesis of equimolar amounts of Z-Gln-OH and H-Leu-NH2 as model substrates, the water content was varied from 0 to 600 mL water (mol substrate)−1 and enzyme concentration in the range 0.5–10 g (mol substrate)−1 to achieve 80% yield and initial rates of 5–20 mmol (s kg)−1 [22]. When the water content is decreased from the 1.6-molal lowest substrate concentration, the initial rate increases tenfold to a pronounced optimum at 40 mL water (mol substrate)−1 and falls to much lower values in a system with no added water, and to zero in a rigorously dried system. The behavior at a higher water content was demonstrated through variation of the enzyme content to be caused by mass transfer limitations; at low water levels, the effects reﬂect the stimulation of the enzymatic activity by water. Preheating of the substrates or ultrasonic treatment had no signiﬁcant effect on the system.

26 Biocatalysis in Non-conventional Media

During melting point experiments, the formation of a third compound, the salt of the two substrates, was discovered and coincided with a very strong dependence of kinetics on the exact substrate ratio: at 60% Leu-NH2 and 40% Z-Gln the rate was twice as high as at an equimolar substrate ratio [23]. The reaction rate of both Z-GlnLeu-NH2 and Z-Asp-Phe-OMe formation is strongly dependent on the addition of basic salts to the reaction system, pointing to the importance of acid-base effects in solid-to-solid systems [25]. If either KHCO3 or K2 CO3 is raised to 2.25-fold the equimolar amount of the Phe-OMe · HCl starting material, the rate increases 20fold and remains at that level with further addition of KHCO3 , but it drops sharply when further K2 CO3 is added. Beyond the addition of 1.5 equivalents of basic salt, the ﬁnal yield of the reaction is affected negatively. These effects can be rationalized using a model estimating the pH of these systems, taking into account the possible formation of up to ten different solid phases. Together with inﬂuences of different mixing and water distribution methods, the pH model seems to explain most of the experimental results. Speeding up of enzymatic conversions of substrate in aqueous suspensions is often attempted by raising the temperature or adding organic solvents to promote dissolution of the substrate. Results of an α-chymotrypsin-catalyzed hydrolysis of dimethylbenzyl methyl malonate demonstrated that, upon addition of organic cosolvents, longer process times were actually required, even though the substrate solubility increased severalfold as expected; however, on raising the temperature from 25 to 37 ◦ C, the substrate solubility, the substrate dissolution rate, and the enzymatic reaction rate all increased, leading to shorter process times [66]. A simple relationship for the prediction of the overall process time was established by evaluating the time constants for three sub-processes involved: substrate dissolution, enzymatic conversion, and enzyme deactivation [Eqs. (15)–(17)]. Substate dissolution:τdis

= 1/(kL a)0

(15)

Enzymatic conversion:τenz = K M /(kcat · [E]0 )

(16)

Enzyme deactivation:τdeact = 1/(kd + [E]0 )

(17)

With the assumptions of second-order deactivation and ﬁrst-order reaction, the regimes are characterized as follows: ([A]solid,0 = initial concentration of solid substrate, [A]lim = substrate solubility): Ĺ In the deactivation-limited regime, if τ deact ≈ [A]lim · τ enz /(0.4 · [A]solid,0 ), the process time is totally dominated by deactivation. If [A]solid,0 > [A]lim · τ enz /(0.4 · τ deact ) the enzyme is deactivated before the reaction is complete. Ĺ In the dissolution-limited regime, if deactivation is insigniﬁcant, τ deact [A]lim · τ enz /(0.4 · [A]solid,0 ), overall product formation rate r P and the overall process time tpro are given by Eqs. (18).

r p = kL a · [A]lim and tpro ≈2τdis ([A]solid,0 /[A]lim )

(18)

6 Solvent as a Parameter for Reaction Optimization (“Medium Engineering”)

Ĺ In the reaction-limited regime, τ deact [A]lim · τ enz /(0.4 · [A]solid,0 ) and τ enz 2 · τ dis , and r P and tpro are then given by Eq. (19).

r p = (kcat /K M ) · [E]0 · [A]lim and tpro ≈τenz ([A]solid,0 /[A]lim )

(19)

In this case, the concentration of dissolved substrate equals [A]lim . The subprocesses, such as dissolution/crystallization of substrates and products, enzymatic synthesis of the product(s), undesired enzymatic hydrolysis of substrates and/or products, and deactivation, often inﬂuence the pH value and are inﬂuenced in turn by it as the reactants are weak electrolytes. Investigating the kinetically controlled synthesis of the β-lactam antibiotic amoxicillin from 6-aminopenicillanic acid and D-p-hydroxyphenylglycine methyl ester in a solid suspension system in which the reaction nevertheless occurred in the liquid phase, Diender et al. found that the pH value and dissolved concentrations took a very different course at different initial substrate amounts [18]. These results were described reasonably well by the model based on mass and charge balances, pH-dependent solubilities of the reactants, and enzyme kinetics. Whereas enzymatic kinetic resolutions in a solid substrate suspension are usually treated as conventional kinetic resolutions of a fully dissolved substrate, a modeling study has demonstrated that there are important differences leading to much better performance in a solid suspension case [68]. In the suspension processes the liquid-phase concentration of substrate enantiomer that should be converted can be kept close to the maximum value, i.e., the solubility, when process conditions are properly chosen, whereas in a conventional process this concentration gradually decreases. Calculations show that this leads to a productivity that is about sixfold higher in the suspension processes. Also, for enzymes with a low enantioselectivity, a severalfold increase in yield of remaining enantiopure substrate is predicted, compared to the conventional kinetic resolution of dissolved enantiomers. Other potential advantages of using suspension reactions are that the initial substrate concentration may be higher [up to 25% (w/w)] and that the desired remaining substrate may be recovered by simply ﬁltering off the solid crystals.

6 Solvent as a Parameter for Reaction Optimization (“Medium Engineering”)

The term “medium engineering” was coined as a complement to protein engineering [6]. Whereas the latter changes the catalyst, the former changes the reaction medium with resulting changes in prochiral, regio-, and enantioselectivity. Many such changes have been observed in the literature [6, 10, 11], however, only a few examples are given here to demonstrate that not only the catalyst but also the medium exerts a decisive inﬂuence on the outcome of an enzymatic reaction.

27

28 Biocatalysis in Non-conventional Media Table 8 Substrate speciﬁcity S (kcat /K M )Ser /(kcat /K M )Phe , in the transesteriﬁcation of N-Ac-

l-amino acid ethyl ester with n-propanol, catalyzed by Subtilisin Carlsberg in various anhydrous organic solvents [69]

Solvent Dichloromethane Chloroform Toluene Benzene N,N-Dimethylformamide tert-Butyl acetate N-Methylacetamide Diethylether Carbon tetrachloride Ethyl acetate

S 8.2 5.5 4.8 4.4 4.3 3.7 3.4 3.2 3.2 2.6

Solvent tert-Butyl methyl ether Octane Isopropyl acetate Acetonitrile Dioxane Acetone Pyridine tert-Amyl alcohol tert-Butyl alcohol tert-Butylamine

S 2.5 2.5 2.2 1.7 1.2 1.1 0.53 0.27 0.19 0.12

6.1 Change of Substrate Speciﬁcity with Change of ReactionM: Speciﬁcity of Serine Proteases

When switching from water to an organic solvent, or switching between organic solvents, the substrate speciﬁcity can change. In the example of the standard reaction, transesteriﬁcation of N-acetyl-L-phenylalanine ethyl ester with n-propanol by Subtilisin Carlsberg, which has been mentioned several times in this chapter already, the relative speciﬁcity between the rather hydrophobic phenylalanine compound and its more hydrophilic analog N-acetyl-L-serine ethyl ester varies with the solvent (Table 8) [69]. Partitioning of the two substrates between the bulk organic solvent and the binding pocket of the enzyme active site can act as an explanation. It is known that any binding pocket of enzymes can be identiﬁed with a value for the hydrophobicity. A hydrophobic substrate in water partitions readily into the binding pocket, but the more hydrophobic the solvent becomes the worse is the partitioning that results. From Table 8 it can be discerned, however, that this monocausal explanation does not sufﬁce, as speciﬁcity S does not follow hydrophobicity very well. 6.2 Change of Regioselectivity by Organic Solvent Medium

Ortho-substituted phenolic diesters, such as bis-p-(1,4-butyroyl)-2octylhydroquinone (Figure 9), can be hydrolyzed in the more hindered 1-position or in the less-hindered 4-position. The regioselectivity of hydrolysis to the monoester, as expressed by the ratio of kcat /K M values, depends approximately linearly on log P over the range from acetonitrile (log P: −0.33) to cyclohexane (log P: 3.2), and is thus controlled by the hydrophobicity of the solvent (Carrea, 2000).

6 Solvent as a Parameter for Reaction Optimization (“Medium Engineering”)

Fig. 9 Regioselectivity of the hydrolysis of hydroquinone bis-esters [? ].

In another case, 4-alkylphenols can be transformed with the help of the ﬂavoenzyme vanillyl alcohol oxidase (VAO) to either (R)-1-(4 -hydroxyphenyl) alcohols or to 1-(4 -hydroxyphenyl)alkenes. Both products pass through a common intermediate, the p-quinone methide, which then either is attacked by water or rearranges. The product spectrum can be controlled by medium engineering: in organic solvents such as acetonitrile and toluene, more cis-alkene but not transalkene, and less alcohol, are produced [71]. A similar shift in cis/trans-alkene was achieved by the addition of monovalent anions that bind speciﬁcally close to the active site.

Fig. 10 Asymmetric hydrolysis of nifedipine diesters; inﬂuence of solvent on stereochemical preferences [11, 38].

29

30 Biocatalysis in Non-conventional Media

6.3 Solvent Control of Enantiospeciﬁcity of Nifedipines

The probably most striking example so far of solvent inﬂuence on enzyme selectivity has been found by Yoshihiko Hirose and co-workers at the Amano Corp. in Nagoya, Japan, studying nifedipines. Nifedipines, structurally 1-substituted dihydropyridine mono- or diesters, are used in cardiovascular therapy, where they are R termed calcium antagonists [32]; the most prominent representative is Adalat from Bayer (Leverkusen, Germany). While the simplest achiral dihydropyridines can be synthesized by simple mixing and heating of benzaldehydes, ammonia, and alkyl ac-etoacetate, the asymmetric – and thus difﬁcult to synthesize – dihydropyridines are therapeutically the most valuable today. The Amano researchers found not just drastically different enantiomeric excesses in the Pseudomonas sp. (Amano AH) lipase-catalyzed hydrolysis of methyleneoxypropionyl or -pivaloyl diesters, but a near-total reversal of speciﬁcity. Whereas in cyclohexane the different esters yielded half-esters with 88.8–91.4% e.e. R-speciﬁcity for a triple mutant (“FVL”) of Amano PS lipase, the same transformation with the same enzyme in diisopropyl ether (DIPE) yielded between 68.1 and > 99% e.e. of the S-product (Figure 10) [7, 38].

Suggested Further Reading G. CARREA, G. OTTOLINA, and S. RIVA, Role of solvents in the control of enzyme selectivity in organic media, Trends Biotechnol. 1995, 13, 63–70. G. CARREA and S. RIVA, Properties and synthetic applications of enzymes in

organic solvents, Angew. Chem. Int. Ed. 2000, 39, 2226–2254. A. M. KLIBANOV, Improving enzymes by using them in organic solvents, Nature 2001, 409, 241–246.

References 1 E. A. SYM, The action of esterase in the presence of organic solvents, Biochem. J. 1936, 30, 609–617. 2 A. M. KLIBANOV, G. P. SAMOKHIN, K. MARTINEK, and I. V. BEREZIN, 1977, A new approach to preparative enzymatic synthesis, Biotechnol. Bioeng. 19, 1351–1361, reprinted in Biotechnol. Bioeng 2000, 67, 737–747. 3 A. ZAKS and A. M. KLIBANOV, Enzyme-catalyzed processes in organic solvents, Proc. Natl. Acad. Sci. USA 1986a, 82, 3192–3196. 4 L. DAI and A. M. KLIBANOV, Peroxidase-catalyzed asymmetric

sulfoxidation in organic solvents versus in water, Biotechnol. Bioeng. 2000, 70, 353–357. 5 A. ZAKS and A. M. KLIBANOV, Enzymatic catalysis in organic media at 100 degrees C, Science 1984, 224, 1249–1251. 6 C. R. WESCOTT and A. M. KLIBANOV, The solvent dependence of enzyme speciﬁcity, Biochim. Biophys. Acta 1994, 1206, 1–9. 7 Y. HIROSE, K. KARIYA, Y. NAKANISHI, Y. KURONO, and K. ACHIWA, Inversion of enantioselectivity in hydrolysis of 1, 4-di-hydropyridines by point mutation of lipase PS, Tetrahedon Lett. 1995, 36, 1063–1066.

References 8 A. M. KLIBANOV, 1989, Enzymatic catalysis in anhydrous organic solvents, Trends Biochem. Sci. 14, 141–144. 9 L. A. S. GORMAN and J. S. DORDICK, Organic solvents strip water off enzymes, Biotechnol. Bioeng. 1992, 39, 392–397. 10 A. ZAKS and A. M. KLIBANOV, Substrate speciﬁcity of enzymes in organic solvents vs. water is reversed, J. Am. Chem. Soc. 1986b, 108, 2767–2768. 11 V. S. NARAYAN and A. M. KLIBANOV, Are water-immiscibility and apolarity of the solvent relevant to enzyme efﬁciency? Biotechnol. Bioeng. 1993, 41, 390–393. 12 R. BOVARA, G. CARREA, G. OTTOLINA, and S. RIVA, Effects of water activity on V max and K M of lipase catalyzed transesteriﬁcation in organic media, Biotechnol. Lett. 1993, 15, 937–942. 13 R. H. VALIVETY, P. J. HALLING, and A. R. MACRAE, Reaction rate with suspended lipase catalyst shows similar dependence on water activity in different organic solvents, Biochim. Biophys. Acta 1992a, 1118, 218–222. 14 R. H. VALIVETY, P. J. HALLING, A. D. PEILOW, and A. R. MACRAE, Lipases from different sources vary widely in dependence of catalytic activity on water activity, Biochim. Biophys. Acta 1992b, 1122, 143–146. 15 G. BELL, A. E. M. JANSSEN, and P. J. HALLING, Water activity fails to predict critical hydration level for enzyme activity in polar organic solvents: interconversion of water concentrations and activities, Enzyme Microb. Technol. 1997, 20, 471–477. 16 A. ZAKS and A. M. KLIBANOV, Enzymatic catalysis in nonaqueous solvents, J. Biol. Chem. 1988a, 263, 3194–3201. 17 H. R. COSTANTINO, K. GRIEBENOW, R. LANGER, and A. M. KLIBANOV, On the pH memory of lyophilized compounds containing protein functional groups, Biotechnol. Bioeng. 1997, 53, 345–348. 18 L. T. KANERVA and A. M. KLIBANOV, Hammett analysis of enzyme action in organic solvents, J. Am. Chem. Soc. 1989, 111, 6864–6865. 19 A. ZAKS and A. M. KLIBANOV, The effect of water on enzyme action in organic media, J. Biol. Chem. 1988b, 263, 8017–8021.

20 R. M. GUINN, P. S. SKERKER, P. KAVANAUGH, and D. S. CLARK, Activity and ﬂexibility of alcohol dehydrogenase in organic solvents, Biotechnol. Bioeng. 1991, 37, 303–308. 21 P. A. BURKE, R. G. GRIFFIN, and A. M. KLIBANOV, Solid-state NMR assessment of enzyme active center structure under nonaqueous conditions, J. Biol. Chem. 1992, 267, 20057–20064. 22 J. BROOS, A. J. W. G. VISSER, J. F. J. ENGBERSEN, W. VERBOOM, A. van HOEK, and D. N. REINHOUDT, Flexibility of enzymes suspended in organic solvents probed by time-resolved ﬂuorescence anisotropy. Evidence that enzyme activity and enantioselecticity are directly related to enzyme ﬂexibility, J. Am. Chem. Soc. 1995, 117, 12657–12663. 23 R. V. RARIY and A. M. KLIBANOV, On the relationship between enzymatic enantio-selectivity in organic solvents and enzyme ﬂexibility, Biocat. Biotransf. 2000, 18, 401–407. 24 J. KIM, D. S. CLARK, and J. S. DORDICK, 2000, Intrinsic effects of solvent polarity on enzymic activation energies, Biotechnol. Bioeng. 67, 112–116. 25 B. A. BEDELL, V. V. MOZHAEV, D. S. CLARK, and J. S. DORDICK, Testing for diffusion limitations in salt-activated enzyme catalysts operating in organic solvents, Biotechnol. Bioeng. 1998, 58, 654–657. 26 C. LAANE, S. BOEREN, and K. VOS, On optimizing organic solvents in multi-liquid-phase biocatalysis, Trends Biotechnol. 1985, 3, 251–252. 27 C. LAANE, S. BOEREN, K. VOS, and C. VEEGER, Rules for optimization of biocatalysis in organic solvents, Biotechnol. Bioeng. 1987, 30, 81–87. 28 P. A. FITZPATRICK, D. RINGE, and A. M. KLIBANOV, Computer-assisted modeling of subtilisin enantioselectivity in organic solvents, Biotechnol. Bioeng. 1992, 40, 735–742. 29 F. TERRADAS, M. TESTON-HENRY, P. A. FITZPATRICK, and A. M. KLIBANOV, Marked dependence of enzyme prochiral selectivity on the solvent, J. Am. Chem. Soc. 1993, 115, 390–396. S. M. THOMAS, R. DICOSINO, and V. NAJARAJAN, Biocatalysis: applications and potentials

31

32 Biocatalysis in Non-conventional Media

30

31

32

33

34

35

36

37

38

39

40

41

for chemical industry, Trends Biotechnol. 2002, 20, 238–242. A. M. KLIBANOV, 1995, Enzyme memory. What is remembered and why?, Nature, 374, 596. T. KE and A. M. KLIBANOV, 1998, On enzymatic activity in organic solvents as a function of enzyme history, Biotechnol. Bioeng. 57, 746–750. L. DAI and A. M. KLIBANOV, Striking activation of oxidative enzymes suspended in nonaqueous media, Proc. Natl. Acad. Sci. 1999, USA 96, 9475–9478. A. M. KLIBANOV, 1997, Why are enzymes less active in organic solvents than in water? Trends Biotechnol. 15, 97–101. D. J. van UNEN, J. F. ENGBERSEN, and D. N. REINHOUDT, 1998, Large acceleration of alpha-chymotrypsin-catalyzed dipeptide formation by 18-crown-6 in organic solvents, Biotechnol. Bioeng. 59, 553–556. D. J. van UNEN, J. F. ENGBERSEN, and D. N. REINHOUDT, 2002, Why do crown ethers activate enzymes in organic solvents? Biotechnol. Bioeng. 77, 248–255. D. J. van UNEN, J. F. ENGBERSEN, and D. N. REINHOUDT, 2001, Sol-gel immobilization of serine proteases for application in organic solvents, Biotechnol. Bioeng. 75, 154–158. M. Y. LEE and J. S. DORDICK, 2002, Enzyme activation for nonaqueous media, Curr. Opin. Biotechnol. 13, 376–384. T. KE and A. M. KLIBANOV, Markedly enhancing enzymic enantioselectivity in organic solvents by forming substrate salts, J. Am. Chem. Soc. 1999, 121, 3334–3340. J. S. SHIN, S. LUQUE, and A. M. KLIBANOV, 2000, Improving lipase enantioselectivity in organic solvents by forming substrate salts with chiral agents, Biotechnol. Bioeng. 69, 577–583. P. K. DAS, J. M. CAAVEIRO, S. LUQUE, and A. M. KLIBANOV, Binding of hydrophobic hydroxamic acids enhances peroxidase’s stereoselectivity in nonaqueous sulf-oxidations, J. Am. Chem. Soc. 2002, 124, 782–787. S. KAMAT, J. BARRERA, E. J. BECKMAN, and A. J. RUSSELL, Biocatalytic synthesis of acrylates in organic solvents and supercritical ﬂuids: I. Optimization of

42

43

44

45

46

47

48

49

50

51

enzyme environment, Biotechnol. Bioeng. 1992a, 40, 158–166. U. KRAGL, M. ECKSTEIN, and N. KAFTZIK, 2002, Enzyme catalysis in ionic liquids, Curr. Opin. Biotechnol. 13, 565–571. M. ERBELDINGER, X. NI, and P. J. HALLING, Kinetics of enzymatic solid-to-solid peptide synthesis: synthesis of Z-aspartame and control of acid-base conditions by using inorganic salts, Biotechnol. Bioeng. 2001, 72, 69–76. N. KAFTZIK, P. WASSERSCHEID, and U. KRAGL, Use of ionic liquids to increase the yield and enzyme stability in the β-galactosidase catalyzed synthesis of N-acetyllactosamine, Org. Proc. Res. Dev. 2002, 6, 553–557. S. PARK and R. J. KAZLAUSKAS, Improved preparation and use of room-temperature ionic liquids in lipase-catalyzed enantioand regioselective acylations, J. Org. Chem. 2001, 66, 8395–8401. M. ECKSTEIN, P. WASSERSCHEID, and U. KRAGL, Enhanced enantioselectivity of lipase from Pseudomonas sp. at high temperatures and ﬁxed water activity in the ionic liquid, 1-butyl-3-methylimidazolium bis[(triﬂuoromethyl)sulfonyl]amide, Biotechnol. Lett. 2002b, 24, 763–767. P. LOZANO, T. de DIEGO, J.-P. GUEGAN, M. VAULTIER, and J. L. IBORRA, Stabilization of α-chymotrypsin by ionic liquids in transesteriﬁcation reactions, Biotechnol. Bioeng. 2001, 75, 563–569. J. W. SHIELD, H. D. FERGUSON, A. S. BOMMARIUS, and T. A. HATTON, Enzymes in reversed micelles as catalysts for organic-phase reactions, Ind. Eng. Chem. Fundam. 1986, 25, 603–612. A. S. BOMMARIUS, D. I. C. WANG, and T. A. HATTON, Xanthine oxidase reactivity in reversed micellar systems: a contribution to the prediction of enzymatic activity in organized media, J. Am. Chem. Soc. 1995, 117, 4515–4523. D. JOBE, in: Reactions in Compartmentalized Liquids, W. KNOCHE and R. SCHOMA¨ CKER (eds.), Springer Verlag, Heidelberg, 1989, pp. 39–51. N. L. KLYACHKO, A. V. LEVASHOV, A. V. PSHEZHETSKII, N. G. BOGDANOVA, I. V. BEREZIN, and K. MARTINEK, Catalysis by

References

52

53

54

55

56

57

58

59

60

61

enzymes entrapped into hydrated surfactant aggregates having lamellar or cylindrical (hexagonal) or ball-shaped (cubic) structure in organic solvents, Eur. J. Biochem. 1986, 161, 149–154. N. H. GRANT and H. E. ALBURN, Acceleration of enzyme reactions in ice, Nature 1966, 212, 194. M. SCHUSTER, A. AAVIKSAAR, and H.-D. JAKUBKE, Enzyme-catalyzed peptide synthesis in ice, Tetrahedron 1990, 24, 8093–8102. M. HAENSLER, N. WEHOFSKY, S. GERISCH, J. D. WISSMANN, and H.-D. JAKUBKE, Reverse catalysis of elastase from porcine pancreas in frozen aqueous systems, Biol. Chem. Hoppe Seyler 1998, 379, 71–74. S. GERISCH, G. ULLMANN, K. STUBENRAUCH, and H.-D. JAKUBKE, Enzymatic peptide synthesis in frozen aqueous systems: inﬂuence of modiﬁed reaction conditions on the peptide yield, Biol. Chem. Hoppe Seyler, 1994, 375, 825–828. G. ULLMANN, M. HAENSLER, W. GRUENDER, M. WAGNER, H. J. HOFMANN, and H.-D. JAKUBKE, Inﬂuence of freezeconcentration effect on proteinase-catalysed peptide synthesis in frozen aqueous systems, Biochim. Biophys. Acta 1997, 1338, 253–258. I. GILL and E. VULFSON, Enzymic catalysis in heterogeneous eutectic mixtures of substrates, Trends Biotechnol. 1994, 12, 118–122. A. MILLQVIST-FUREBY, I. S. GILL, and E. N. VULFSON, 1998, Enzymatic transformations in supersaturated substrate solutions: I. A general study with glycosidases, Biotechnol. Bioeng. 60, 190–196. I. GILL and R. VALIVETY, Pilot-scale enzymatic synthesis of bioactive oligopeptides in eutectic-based media, Org. Proc. Res. Dev. 2002, 6, 684–691. M. T. RU, J. S. DORDICK, J. A. REIMER, and D. S. CLARK, 1999, Optimizing the salt-induced activation of enzymes in organic solvents: effects of lyophilization time and water content, Biotechnol. Bioeng. 63, 233–141. P. J. HALLING, U. EICHHORN, P. KUHL, and H. D. JAKUBKE, Thermodynamics of

62

63

64

65

66

67

68

69

solid-to-solid conversion and application to enzymic peptide synthesis, Enzyme Microb. Technol. 1995, 17, 601– 606. U. EICHHORN, A. S. BOMMARIUS, K. DRAUZ, and H.-D. JAKUBKE, Synthesis of dipeptides by suspension-to-suspension conversion via thermolysin catalysis: from analytical to preparative scale, J. Pept. Sci. 1997, 3, 245–251. H. D. JAKUBKE, U. EICHHORN, M. HANSLER, and D. ULLMANN, Non-conventional enzyme catalysis: application of proteases and zymogens in biotransformations, Biol. Chem. Hoppe Seyler 1996, 377, 455–464. M. ERBELDINGER, X. NI, and P. J. HALLING, Effect of water and enzyme concentration on thermolysin-catalyzed solid-to-solid peptide synthesis, Biotechnol. Bioeng. 1998, 59, 68–72. M. ERBELDINGER, X. NI, and P. J. HALLING, Kinetics of enzymatic solid-to-solid peptide synthesis: intersubstrate compound, substrate ratio, and mixing effects, Biotechnol. Bioeng. 1999, 63, 316–321. A. WOLFF, L. ZHU, Y. W. WONG, A. J. STRAATHOF, J. A. JONGEJAN, and J. J. HEIJNEN, Understanding the inﬂuence of temperature change and cosolvent addition on conversion rate of enzymatic suspension reactions based on regime analysis, Biotechnol. Bioeng. 1999a, 62, 125–134. M. B. DIENDER, A. J. STRAATHOF, T. van der DOES, M. ZOMERDIJK, and J. J. HEIJNEN, Course of pH during the formation of amoxicillin by a suspension-to-suspension reaction, Enzyme Microb. Technol. 2000, 27, 576–582. A. WOLFF, V. van ASPEREN, A. J. STRAATHOF, and J. J. HEIJNEN, Potential of enzymatic kinetic resolution using solid substrates suspension: improved yield, productivity, substrate concentration, and recovery, Biotechnol. Prog. 1999b, 15, 216–227. C. R. WESCOTT and A. M. KLIBANOV, Solvent variation inverts substrate speciﬁcity of an enzyme, J. Am. Chem. Soc. 1993, 115, 1629–1631.

33

34 Biocatalysis in Non-conventional Media

70 E. RUBIO, A. FERNANDEZ-MAYORALES, and A. M. KLIBANOV, Effect of the solvent on enzyme regioselectivity, J. Am. Chem. Soc. 1991, 113, 695–696. 71 R. H. H. van den HEUVEL, J. PARTRIDGE, C. LAANE, P. J. HALLING, and W. J. H. van BERKEL Tuning of the product spectrum of vanillyl-alcohol oxidase by medium engineering, FEBS Lett. 2001, 503, 213–216. 72 S. GOLDMANN and J. STOLTEFUSS, 1, 4-Dihydro-pyridines: effect of chirality and conformation on the calcium-antagonistic and -agonistic effects, Angew. Chem. 1991, 103, 1587–605; Angew. Chem. Int. Ed. Engl. 1991, 30, 1559–1578. 73 Y. HIROSE, K. KARIYA, I. SASAKI, Y. KURONO, H. EBIIKE, and K. ACHIWA, Drastic solvent effect on lipase-catalyzed enantioselective hydrolysis of prochiral 1, 4-dihydropyridines, Tetrahedron Lett. 1992, 33, 7157–7160. 74 G. CARREA, G. OTTOLINA, and S. RIVA, Role of solvents in the control of enzyme selectivity in organic media, Trends Biotechnol. 1995, 13, 63–70. 75 G. CARREA and S. RIVA, Properties and synthetic applications of enzymes in organic solvents, Angew. Chem. Int. Ed. 2000, 39, 2226–2254. 76 M. F. CHAPLIN and C. BUCKE, Enzyme Technology, Cambridge University Press, Cambridge, UK, 1990, p. 229. 77 J. S. DEETZ and J. D. ROZZELL, Enzymic catalysis by alcohol dehydrogenases in organic solvents, Ann. N. Y. Acad. Sci. 1988, 542( Enzyme Eng. 9), 230–234. 78 M. ECKSTEIN, M. SESING, U. KRAGL, and P. ADLERCREUTZ, At low water activity α-chymotrypsin is more active in an ionic liquid than in non-ionic organic solvents, Biotechnol. Lett. 2002a, 24, 867–872. 79 M. ERBELDINGER, A. J. MESIANO, and A. J. RUSSELL, Enzymatic catalysis of formation of Z-aspartame in ionic liquid – an alternative to enzymatic catalysis in organic solvents, Biotechnol. Prog. 2000, 16, 1129–1131.

80 P. A. FITZPATRICK and A. M. KLIBANOV, How can the solvent affect enzyme enantioselectivity? J. Am. Chem. Soc. 1991, 113, 3166–3171. 81 A. S. GHATORAE, G. BELL, and P. J. HALLING, Inactivation of enzymes by organic solvents: new technique with well-deﬁned interfacial area, Biotechnol. Bioeng. 1994, 43, 331–336. 82 S. KAMAT, E. J. BECKMAN, and A. J. RUSSELL, Role of diffusion in nonaqueous enzymology. 1. Theory, Enz. Microb. Technol. 1992b, 14, 265–271. 83 K. MARTINEK, A. V. LEVASHOV, N. KLYACHKO, Yu. L. KHMEL’NITSKII, and I. V. BEREZIN, Micellar enzymology, Eur. J. Biochem. 1986, 155, 453–468. 84 A. MARTY, W. CHULALAKSANANUKUL, J. S. CONDORET, R. M. WILLEMOT, and G. DURAND, Comparison of lipase-catalyzed esteriﬁcation in supercritical carbon dioxide and in n-hexane, Biotechnol. Lett. 1990, 12, 11–16. 85 A. J. MESIANO, E. J. BECKMAN, and A. J. RUSSELL, 1999, Supercritical biocatalysis, Chem. Rev. 99, 623–634. 86 K. OYAMA, S. IRINO, T. HARADA, and N. HAGI, Enzymic production of aspartame, Ann. N. Y. Acad. Sci. (Enzyme Eng. 7) 1984, 434, 95–98. 87 D. ROCHE, K. PRASAD, and O. REPIC, Enantio-selective acylation of β-aminoesters using penicillin G acylase in organic solvents, Tetrahedron Lett. 1999, 40, 3665–3668. 88 M. T. RU, K. C. WU, J. P. LINDSAY, J. S. DORDICK, J. A. REIMER, and D. S. CLARK, 2001, Towards more active biocatalysts in organic media: increasing the activity of salt-activated enzymes, Biotechnol. Bioeng. 75, 187–196. 89 H. WAJANT, S. FORSTER, A. SPRAUER, F. EFFENBERGER, and K. PFIZENMAIER, Enantioselective synthesis of aliphatic (S)-cyanohydrins in organic solvents using hydroxynitrile lyase from Manihot esculenta, Ann. N. Y. Acad. Sci. 1996, 799, 771–776.

1

Photoproteins as Reporters in Whole-cell Sensing Jessika Feliciano, Patrizia Pasini, Sapna K. Deo, and Sylvia Daunert University of Kentucky, Lexington, USA

Originally published in: Photoproteins in Bioanalysis. Edited by Sylvia Daunert and Sapna K. Deo. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31016-6

1 Introduction

Reporter genes code for proteins that produce a detectable signal, which can be differentiated in a mixture of endogenous intra- or extracellular proteins [1, 2]. They have found applications in studies of transcription control, gene expression, cell-signaling mechanisms, drug target discovery, cell-based analyte biosensing, and environmental monitoring of specific microorganisms [3], among others. Photoproteins can serve as reporters in two different ways: via genetically engineered systems that are selective toward specific analytes or by using a constitutively bioluminescent/fluorescent cell line that measures overall toxicity. This chapter will focus on the applications of photoproteins as reporters in genetically engineered whole-cell sensing systems.

1.1 Biosensors Using Intact Cells

A biosensor combines a biological component (the sensing element), which is responsible for the selectivity of the device, with a detection system (the transducer) for measuring the reaction of the biological component with the substance (analyte) being monitored. The biological component can be an enzyme, an antibody, a receptor, or whole cells, while the biological reaction that is monitored can be the activity of an enzyme, the binding of an analyte to an antibody or receptor, the induction of gene expression within cells, or even cell death [4, 5]. A number of detection systems have been employed to monitor these biological events, including electrochemical, optical, and piezoelectric systems [6, 7]. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Photoproteins as Reporters in Whole-cell Sensing

There are a number of advantages in using intact microorganisms, rather than isolated biological components, in biosensors. Microorganisms are usually more tolerant of assay conditions that would be detrimental to an isolated enzyme or protein. Because microorganisms have mechanisms that help them regulate their internal environment, they are also more tolerant of suboptimal pH and temperatures than purified enzymes and are less likely to be inhibited by solutes in the sample being assayed. Microorganisms are cheaper to use in biosensors because the active biological component does not have to be isolated and unlimited quantities can be prepared relatively inexpensively [8–10]. Because an analyte, such as a pollutant, must be taken up by the microorganism before a response is produced, using intact microorganisms can provide information about the bioavailability of the analyte. Classical analytical methods of detection cannot distinguish between bioavailable pollutants from those that are unavailable. In that regard, sensing systems using intact cells complement the more conventional sensors where isolated biological components are being used. These systems may provide physiologically relevant data and give an indication of the bioavailability of a pollutant, which is especially important when decisions regarding remediation of contaminated environments have to be made [9]. While only the bioavailable amount can be obtained, cell-based biosensors are preferred over other biological biosensors because one can acquire other types of information such as toxicity, mutagenicity, and pharmacological activity. Because of all these characteristics, cell-based biosensing systems have found applications in biotechnology, pharmacology, molecular biology, and clinical and environmental chemistry. Among the advances over the years is the constant wish to improve the systems. Improved high-throughput assays for gene function, automation to enhance screening and capacity, and miniaturization to reduce costs have been developed. Systems for the correction of changes in signal that are due to the physiological state of the cell have also been developed. For a good review on how the bioreporter design can be improved by mutagenizing the sensing and the regulatory proteins involved, readers are encouraged to review Ref. [9]. Several reviews have been written on cell-based biosensors, including by our group in 2000 [10]. This chapter will present a general view of the photoproteins with an emphasis on their application specifically to bacterial whole-cell sensing systems. The most advanced technology from our 2000 review will be highlighted, as well as advances in reporter gene technology in cell-based assays since then. 1.2 Reporter Genes in Genetically Engineered Whole-cell Sensors

Genetically engineered whole-cell sensing systems can be developed by coupling a sensing element, which recognizes the analyte and confers selectivity to the system, with a reporter gene, which produces the detectable signal and determines the sensitivity of the system. In this chapter whole-cell sensors that employ receptors as sensing elements will not be discussed. Such biosensing systems are usually based

1 Introduction 3

Fig. 1 Schematic representation of a whole-cell sensing system based on reporter genes.

on eukaryotic cells, either yeast or mammalian cells, and are routinely used in drug screening assays and for the environmental detection of dioxin- and estrogen-like compounds [11–13]. Instead, we will focus on inducible operon-based whole-cell sensors, in which the sensing element is composed of regulatory proteins that are capable of molecular recognition and promoter regions (O/P) of DNA. In a whole-cell sensing system, the regulatory protein controls the transcription of the fused genes. When the target analyte is present, the regulatory protein is capable of recognizing the analyte and will unbind itself from the operator/promoter region, causing transcription to occur. Thus, when the induction takes place in the presence of the target analyte, the reporter gene is co-expressed along with the other genes of the operon. Consequently, the concentration of the inducer can be quantified by measuring the signal generated by the reporter protein (Fig. 1). When a reporter is being selected, one has to take into consideration which type of application the reporter is needed for, including how sensitive the assay should be, in what concentration range one needs to detect, and the nature of the target analyte. In the case of sensitivity, when the reporter gene codes for an enzyme, internal amplification cascades can be used to increase the sensitivity of the device. The stability of the reporter plays a very important role as well. For instance, green fluorescent protein variants that are unstable have different halflives and are more appropriate for kinetic studies than the red fluorescent protein, which takes days to maturate. A good reporter should be environmentally safe, easy to use, highly sensitive, relatively small in size (for inclusion in the vector), and nontoxic, so that expression is related only to the gene under study and not to overall toxicity (biologically inert and benign). Additionally, the reporter should produce a wide dynamic response, its signal should be distinguishable over background, and the expressed product should be enough so that it can be detected directly or

4 Photoproteins as Reporters in Whole-cell Sensing Table 1 Advantages and disadvantages of photoproteins in whole-cell sensing applications

Reporter protein

Advantages

Disadvantages

Bacterial luciferase

High sensitivity. Does not require addition of a substrate. No endogenous activity in mammalian cells. No light source requirement.

Heat labile and therefore of limited use in mammalian cells. Narrow linear range.

Firefly luciferase

High sensitivity. Broad linear range. No endogenous activity in mammalian cells. High quantum yield. Spectral variants.

Sea pansy luciferase

No endogenous activity in bacterial and mammalian cells. No light source requirement. Its substrate, coelenterazine, is membrane permeable. High sensitivity. No endogenous activity in mammalian cells. No light source requirement. Detection down to subattomole levels. Autofluorescent and thus does not require addition of a substrate or cofactors. Spectral variants. No endogenous homologues in most systems. Stable at biological pH.

Requires addition of a substrate. Requires an aerobic environment and ATP. Requires addition of solubilizers for the substrate to permeate the cell. Flash-type emission. Requires addition of a substrate.

Aequorin

Green fluorescent protein

Requires addition of a substrate and the presence of Ca2+ . Flash-type emission. Moderate sensitivity (no signal amplification). Requires post-translational modification. Background fluorescence from biological systems may interfere. Potential cytotoxicity in some cell types.

indirectly. Table 1 summarizes the advantages and disadvantages of the reporter photoproteins. Details about specifics on each of the reporter proteins have been the subject of reviews elsewhere and will not be discussed here. The following section covers the specific characteristics of each photoprotein when used as a reporter in a cell-based assay.

2 The Luciferases

Bioluminescence is the term used to describe the light produced by chemical excitation within living organisms. In many cases light emission is based on luciferasemediated oxidation reactions. Luciferase is a generic name for an enzyme that catalyzes a light-emitting reaction [14]. Luciferases can be found in both terrestrial and aquatic organisms, including bacteria, algae, fungi, jellyfish, insects, shrimps, squids, and fireflies [15]. The most widely used luciferase genes in reporter gene

2 The Luciferases Table 2 Chemical reactions involving photoproteins

Reporter protein

Catalyzed reaction

Bacterial luciferase

FMNH2 + R-CHO + O2 → FMN + H2 O + RCOOH + hν (490 nm)

Firefly luciferase

Firefly luciferin + O2 + ATP → oxyluciferin + AMP + Pi + hν (550–575 nm)

Renilla luciferase

Renilla luciferase + Coelenterazine + O2 → coelenteramide + CO2 + hν (482 nm)

Aequorin

Coelenterazine + O2 + Ca2+ → coelenteramide + CO2 + hν (469 nm)

Green fluorescent protein

Post-translational formation of an internal chromophore

assays are the bacterial luciferase genes (lux genes of terrestrial Photorhabdus luminescens and marine Vibrio harveyi bacteria) and the eukaryotic luc and ruc genes from firefly (Photinus pyralis) and sea pansy (Renilla reniformis). 2.1 Bacterial Luciferases

Bacterial luciferases have found applications in gene transfer and expression, promoter analysis, gene delivery, imaging of gene expression [16], drug discovery and signaling pathways, etc. However, because they are heat-labile dimeric proteins, their applications in mammalian cell-based systems have been limited [1]. Bacterial luciferase catalyzes the oxidation of a reduced flavin mononucleotide (FMNH2 ) and a long-chain aldehyde to FMN and the corresponding long-chain carboxylic acid, emitting light at 490 nm (Table 2). The blue-green light produced in this reaction has a quantum yield between 0.05 and 0.15 [17]. The lux genes from bacteria luciferases have been isolated and extensively employed in the construction of bioreporters. They are five genes that are organized in the luxCDABE operon: luxA and luxB code for the α and β subunits of bacterial luciferase, while luxC, luxD, and luxE code for the synthesis enzymes for the long-chain aldehyde substrate. These five genes are conserved in all bacterial species identified to date. Depending on the combination of the genes employed, different types of bioluminescent sensing systems can be developed [18]. 2.1.1 luxAB Bioreporters These bioreporters contain only the luxA and luxB genes, which are sufficient for generating the bioluminescent signal; however, because the genes that code for the production of the substrate are not present, the substrate must be supplied to the cell. Typically, decanal is added at some point during the assay. This system has the advantage that one can control the moment when the reaction is triggered and the measurement be taken right after, dramatically reducing the basal

5

6 Photoproteins as Reporters in Whole-cell Sensing

bioluminescence resulting from “leaky” promoters. Various luxAB sensing systems have been developed in bacterial, yeast, insect, nematode, plant, and mammalian cell sensing systems. Nevertheless, the addition of a substrate makes it impractical for deployable applications, and platforms for biosensors have utilized the entire luxCDABE cassette instead because expression of all five genes has the advantage of not requiring addition of a substrate. 2.1.2 luxCDABE Bioreporters These bioreporters contain the full lux operon cassette, thereby allowing for a completely independent bioluminescent system that requires no external addition of a substrate. In this biosensing system, the bioreporter is exposed to the target analyte and a quantitative change in bioluminescence is detected, often within an hour. The rapidity of the assay, its ease of use, and the substrate-free characteristic of this system make luxCDABE bioreporters ideal for real-time, online, field, and highthroughput applications. For that reason, luxCDABE-based bacterial luciferases are the most widely used photoproteins in whole-cell sensing applications. They have been incorporated into a diverse range of platforms and have found applications in environmental and clinical applications, among others. 2.1.3 Naturally Luminescent Bioreporters One of the earliest applications of bacterial luciferase as a reporter was in the Microtox assay [19, 20]. This assay relies on the changes in light production of the natural luminescence of Photobacterium phosphoreum and is used to measure overall toxicity. Similarly, toxin-sensitive cells can be used in biosensors for the nonspecific detection of toxins. When the cells are exposed to critical amounts of toxins, either the metabolic activity of the cells decreases or the cells die. In either case, as the concentration of toxins in the environment increases there is a corresponding decrease in the signal produced by a reporter protein expressed constitutively within the cells. It should be noted that this type of biosensor will respond to any type of substance that is toxic to the cells. Based on this principle, biosensors have been developed for determining the biotoxicity of a sample [21, 22]. 2.2 Eukaryotic Luciferases 2.2.1 Firefly Luciferase Firefly luciferase, encoded by the luc gene, was first isolated from the North American firefly Photinus pyralis [23, 24]. The structure of this luciferase is different from that of bacterial luciferase, and the resulting bioluminescent reaction exhibits other characteristics as well. The oxygen-dependent bioluminescent reaction converts the substrate luciferin to oxyluciferin based on energy transfer from ATP to the substrate to yield AMP, carbon dioxide, and light in the 550–575 nm range. The bioluminescence produced by firefly luciferase is characterized by a peak after 300 ms, after which it exhibits decay. This decay is produced by the product oxyluciferin, which associates with the enzyme and inhibits light production. This can be

3 Aequorin

overcome by addition of coenzyme A in the reaction mix, which will render dissociation of oxyluciferin from the enzyme and prolong emission over several minutes [23, 24]. The bioluminescence resulting from firefly luciferase exhibits the highest quantum yield known for a bioluminescent reaction (0.88, which is about 10 times larger than that of bacterial luciferase) [1]. Like bacterial luciferase, mammalian organisms do not exhibit endogenous firefly luciferase activity. However, the genes required to synthesize the substrate (luciferin) are not present; thus, addition of an exogenous substrate is always required for bioluminescence to occur. There has been controversy in this matter because the substrate cannot be permeated into the cell [25]. The addition of solubilizers such as DMSO has been required to facilitate entry into the cell. An interesting feature of some eukaryotic luciferases is that they can exhibit light in different wavelengths, ranging from 547 nm to 593 nm [26]. This characteristic has been employed only in colony detection on the same plate, but it can be further exploited in multi-analyte detection by fusing each mutant to a different regulatory gene. Firefly luciferase has found applications in whole-cell sensing systems for the detection of heavy metals and aromatic organics. Its high sensitivity (subattomole level) and broad dynamic range (eight orders of magnitude) make this reporter a favorite in mammalian cell-based sensing systems. 2.2.2 Sea Pansy Luciferase The sea pansy Renilla reniformis is a coelenterate that, unlike other bioluminescent organisms, emits a bright green light. In vivo, Renilla luciferase oxidizes its substrate coelenterazine, and after energy transfer to the green fluorescent protein, light is emitted at 509 nm. In vitro, it is represented by the Rluc gene and produces a blue light of 482 nm [27]. Like other luciferases, Renilla luciferase has no endogenous activity in bacteria and thus can find applications in bacteria-based sensing systems. Studies have shown a sensitivity and detection range similar to the firefly luciferase. Despite its major advantage over firefly luciferase that coelenterazine is membrane permeable, it has limited applications in analytical chemistry. This is probably due to a flash half-life of several seconds. The main application of this eukaryotic luciferase has been in combination with firefly luciferase, in a dual luciferase-based reporter system [28]. Here, both eukaryotic luciferases show a linear response to the target analyte: five orders of magnitude for Renilla luciferase and seven orders of magnitude for firefly luciferase. In both cases, substrate addition makes it impractical for field applications.

3 Aequorin

Aequorin is a calcium-binding photoprotein isolated from the jellyfish Aequorea victoria. This protein is comprised of three components: apoaequorin (the precursor of the photoprotein aequorin), a coelenterazine chromophore, and molecular oxygen [29]. Within the protein, it contains three Ca2+ -binding sites that, once occupied, cause the protein to undergo a conformational change, which in turn triggers the

7

8 Photoproteins as Reporters in Whole-cell Sensing

oxidation of coelenterazine to a highly unstable intermediate, coelenteramide. Like Renilla luciferase, it catalyzes the oxidation of coelenterazine to yield a flash blue light of 469 nm that lasts five seconds. This flash-type light emission is accompanied by the release of CO2 , has a quantum yield between 0.15 and 0.20, and requires a Ca2+ concentration of 0.1–10 µM to induce light emission [30]. To date, aequorin is one of the most extensively studied calcium-binding photoproteins. It can be regenerated by dialyzing with EDTA to remove Ca2+ and adding fresh coelenterazine. Among the advantages of using aequorin in reporter gene assays are its high sensitivity and low background. Aequorin can be detected at subattomole levels and is not toxic to various types of cells [31]. Furthermore, a complete absence of endogenous aequorin in mammalian cells can find applications in this type of cellbased sensing systems. Despite these advantages, aequorin is not a popular reporter in cell-based assays. This is probably due to its flash-type light emission (lasting less than five seconds), which requires a luminometer that can initiate and detect the reaction rapidly. In addition, the exogenous requirement of coelenterazine makes this impractical for out-of-the-laboratory applications. Currently, there are two ways of assessing aequorin bioluminescence: detection of aequorin itself or detection of the luminescence triggered by the addition of Ca2+ as the product of the signaling cascade. The latter is more commonly used. Aequorin is widely used as a label in immunoassays and in nucleic acid probe assays, and, because it is a Ca2+ -binding protein, it is used to monitor intracellular levels of free calcium in both prokaryotes and eukaryotes. However, to our knowledge there is only one application in cell-based sensing assays. Rider et al. developed the CANARY (Cellular Analysis and Notification of Antigen Risks and Yields) assay for the detection of pathogenic bacteria and viruses [32, 33]. In this assay, B cells were genetically engineered to express membrane-form antibodies and apoaequorin. The cells were patterned on glass substrates using photolithography and packed into a flow chamber of a microfabricated device. Once the antigen binds, an intracellular signaling cascade is triggered, causing the release of Ca2+ and the activation of the luminescent reaction of aequorin. Detection of pathogenic bacteria and viruses was achieved in less than 1 min with high specificity and sensitivity. An advantage of this assay is that it can be engineered for any target analyte and can be specific to antigens of different pathogens.

4 Fluorescent Proteins

Fluorescent proteins absorb light at a certain maximum wavelength and emit it at a different maximum wavelength, normally a higher one. The green fluorescent protein (GFP) is by far the most popular fluorescent reporter protein in whole cellbased assays. GFP variants and the red fluorescent protein (DsRed) also have found applications in this field. Spectral properties of fluorescent proteins are reported in Table 3.

4 Fluorescent Proteins Table 3 Excitation and emission maxima of fluorescent photoproteins

Reporter protein

Green fluorescent protein (GFP) Enhanced green fluorescent protein (EGFP) Green fluorescent protein UV (GFPuv) Blue fluorescent protein (BFP/EBFP) Cyan fluorescent protein (CFP/ECFP) Yellow fluorescent protein (YFP/EYFP) Red fluorescent protein (DsRed)

Excitation kmax (nm)

Emission kmax (nm)

395 488 395 380 433 513 558

509 509 509 440 475 527 583

4.1 Green Fluorescent Protein

The green fluorescent protein, encoded by the gfp gene, has been isolated and cloned from the jellyfish Aequorea victoria. In vivo the role of GFP is to shift the blue bioluminescence of aequorin to green fluorescence by means of a radiationless energy transfer from aequorin to GFP [34]. The wild-type GFP has an excitation maximum at 395 nm, with a minor peak at 475 nm, and an emission maximum at 509 nm with a small shoulder at 540 nm [35]. The main advantage of GFP as a reporter protein is that it does not require the addition of exogenous substrates or cofactors to produce light; therefore, its use is not limited by the accessibility of substrates [1]. Detection of GFP requires only irradiation by light, which makes GFP the reporter of choice for in situ detection of ana lytes. Induced bacteria expressing GFP can be detected at a single-cell level by using techniques such as epifluorescence microscopy, flow cytometry, or con focal laser scanning microscopy [36]. GFP is a very stable protein, which is advantageous because long-lasting GFP produced from weak promoters or under conditions of low metabolic activity can accumulate in the biosensor cells over time. GFP can be easily expressed in a wide range of organisms, in particular bacteria. This enabled extensive mutational studies of the native protein and the functions of individual amino acids, with the final goal of altering GFP stability and spectral properties (Fig. 2). Variants with improved fluorescence intensity, thermostability, and chromophore folding have been obtained. Shifts within the excitation and emission spectra have also been achieved, thus making available mutants such as the blue, cyan, and yellow variants that can be used for multi-analyte detection purposes [37]. Different strategies have been used to generate variants with shorter half-lives that are suitable for time-dependent induction studies [38]. Limitations to the use of GFP as a reporter include lower detectability compared to other reporter systems. This is due in part to lack of the amplification step occurring with enzyme reaction-based reporters. Additionally, despite the absence of endogenous GFP homologues in most target organisms, the sensitivity of GFP may be compromised by the presence of other fluorescent molecules found in

9

10 Photoproteins as Reporters in Whole-cell Sensing

Fig. 2 Expression of fluorescent reporter proteins in E. coli. Reprinted in part from Ref. [10] with permission from the American Chemical Society.

biological systems, which increase background signal [2]. The need for O2 for folding and maturation also limits application to aerobic conditions. Overall, GFP has unique characteristics that have led to its successful application as a reporter for various analytes [10, 38]. 4.2 Red Fluorescent Protein

The dsRed gene encodes a red fluorescent protein (DsRed) with homology to GFP. The red fluorescent protein has been cloned from coral Discosoma sp. [39]. It has an excitation peak at 558 nm and an emission peak at 583 nm. These spectral properties are suitable for its use as a reporter protein. In fact, cellular autofluorescence is supposed to be lower at longer wavelengths, which should result in lower background signal with DsRed as compared to GFP [40]. However, the use of DsRed as a reporter is somewhat limited by the fact that it has a maturation period of several hours to days [41]. In one study the performance of dsRed was compared to that of other reporter genes (gfp, luc, and luxCDABE) in identical constructs in which the reporter genes were inserted under the control of mercury-responsive regulatory units (mer) [42]. As expected, both fluorescent proteins gave a slow response, with DsRed being much slower than GFP. DsRed was expected to give better sensitivity than GFP because of lower autofluorescence; however, that did not happen with the mer–dsRed construct used in the study.

5 Multiplexing

Many of these reporters have been used in combination. Multiple reporter genes are commonly utilized to provide multi-analyte detection or as viability control. Various combinations have been employed, including using different spectral variants of the same photoprotein and using a combination of a fluorescent photoprotein

6 Applications

and bioluminescent one either within the same construct in the same cell [43, 44], within separate constructs within the same cell, or in different cell lines [45, 46]. For instance, the click beetle has four different luciferases that produce bioluminescence emission at different emission maxima (548 nm, 560 nm, 578 nm, and 590 nm) [26, 47]. If their emission spectra do not overlap, the genes encoding the 548-nm and 590-nm emission could be used in a dual-detection system. Not only would detection of more than one analyte be feasible, but also the assay would provide a much simpler system because only one substrate would be required. When using bioluminescent photoproteins, the signals are simultaneously generated and independently measured because of either different kinetics (half-life) or different emission spectrum. In contrast, the fluorescent photoproteins have to be independently excited and/or measured because of different excitation and emission spectra. We developed a dual-analyte sensing system for the simultaneous detection of β-lactose and L-arabinose as model analytes. Two variants of the green fluorescent protein, BFP2 and GFPuv, were used as the signal-generating reporter proteins. The cells were excited at a given wavelength, and emission for the two reporters was collected at wavelengths characteristic of each reporter. The fluorescence emission thus obtained could be correlated with the amount of sugars present in the sample. The system proved to be highly selective for the two analytes [48]. Dual-reporter reagents using Renilla and firefly luciferase reporters are commercially available. For viability control, we developed a fluorescent dual-reporter system that utilizes two fluorescent proteins with different emission spectra (GFPuv and YFP) within the same genetic construct. One of the fluorescent proteins was used for the analytical signal, while the other was the internal reference of general cell viability. Consequently, any environmental conditions that resulted in a nonspecific effect on the analytical signal could be corrected. Such a system with internal response correction is useful for environmental samples whose complex matrices can affect cell viability [49].

6 Applications

Luciferases are the most widely used of the reporter proteins in whole-cell sensing systems. Unlike fluorescent photoproteins, there is no light scattering from other components in the mixture, as well as no background emission from the media because the media itself is not luminescent. This low background expression along with the enzymatic nature and high quantum efficiency of bioluminescent reactions account for the higher and faster detectability of bioluminescence when compared to fluorescence. Because luminometers do not contain a light source, whole-cell sensing systems based on bioluminescence, specifically bacterial luciferases, have been more widely used in real-sample analysis and field applications. The small number of instrument components required has made possible the construction

11

12 Photoproteins as Reporters in Whole-cell Sensing Table 4 Whole-cell sensing systems using photoproteins as reporters able to respond to dif-

ferent stress factors Stress inducer

Promoter

Stress response

Reporter genes

References

Protein damage Oxidative damage Oxidative damage Membrane synthesis DNA damage

GrpE KatG MicF FabA RecA

Heat shock Hydrogen peroxide Superoxide Fatty acid synthesis SOS

luxCDABE luxCDABE luxCDABE luxCDABE luxCDABE gfpuv

[44, 51–56] [43, 52–54, 56, 57] [52] [51–57] [43, 51–55, 57–60]

of cheaper and more compact devices that are suitable for portability in contrast to fluorescence devices. 6.1 Stress Factors and Genotoxicants

Recombinant bacterial whole-cell sensing systems for the detection of general stress, oxidative stress, and genotoxic compounds have been developed. In these systems, reporter genes are fused to promoters of different stress-response regulons, which are activated by a broad range of compounds. A panel of these bacterial strains is usually employed in environmental monitoring studies [50]. Numerous bacterial strains have been developed that are able to respond to different stressful conditions with the production of a reporter photoprotein. Selected examples of such strains are reported in Table 4. 6.2 Environmental Pollutants

A variety of bacterial whole-cell sensing systems based on the promoter–reporter gene concept have been developed for the specific or selective detection of single pollutants or classes of pollutants. Many microorganisms have evolved the ability to survive in suboptimal conditions, including contaminated environments. Such ability usually relies on genetically encoded resistance systems. In the presence of toxic compounds such as heavy metals and metalloids, particular bacteria can synthesize specific proteins that confer resistance to those substances. The mechanisms of resistance vary. Some microorganisms develop efflux pumps, loss-of-uptake systems, or chemical detoxification systems [61, 62]. Other bacterial strains living in contaminated sites can degrade organic xenobiotics and utilize them as carbon and energy sources [63]. Many of these resistance pathways are inducible, meaning that protein synthesis occurs only as required by the presence of given compounds, which makes their use possible for analytical purposes. The gene sequences of numerous regulatory systems have been cloned and plasmid reporter vectors have been constructed that feature the regulatory sequence in gene fusion with a reporter gene. The pioneering work of Sayler and coworkers, who constructed a whole cell-based

6 Applications Table 5 Whole-cell sensing systems using photoproteins as reporters for the detection of

environmental inorganic and organic analytes Analyte

Reporter genes

Regulatory unit

References

Arsenite, arsenate, antimonite

luc luxAB gfp gfpuv

ars

[67–71]

Cadmium, lead

luc luxAB

cad

[72]

Cadmium, lead, zinc

luc luxCDABE rs-GFP

znt

[73, 74]

Chromate

luxCDABE

chr

[75]

Copper

luxCDABE

copA

[73, 76, 77]

Iron

gfp

pvd

[78]

Mercury, organomercurials

luc

mer

[79–84]

luxAB luxCDABE gfp Nickel

luxCDABE

cnr

[85]

Silver

luxCDABE

copA

[77]

Alkanes

luxAB

alk

[86]

Aromatic compounds

luxCDABE

sep

[87]

Benzene, toluene, ethylbenzene, xylene (BTEX)

luc gfp luxCDABE

xyl tbu tod

[88–90]

Naphtalene, salicylate

luxCDABE

nah

[91]

Phenolic compounds

luxCDABE luxAB

mop dmp

[92] [93]

Polychlorinated biphenyls

luxCDABE

bph

Nitrate

gfp

nar

[94, 95]

Phosphate

luxCDABE

pho

[96]

sensing system for the detection of naphthalene and salicylate [64], prompted the development of a plethora of constructs responsive to organic and inorganic pollutants and including different reporter genes. A number of these constructs, along with the application of whole cell-based biosensors to different environmental samples, have been presented and discussed in recent reviews [65, 66]. Many whole-cell sensing systems for the detection of metals and metalloids have been developed, while smaller numbers of biosensing systems for organic pollutants are available. In Table 5 we list inorganic and organic analytes for which whole cell-based biosensors have been developed, along with the regulatory units and reporter genes used. Whole-cell sensing systems are suitable for the detection of metabolites that are formed along the pollutants degradation pathways. As an example, a whole-cell

13

14 Photoproteins as Reporters in Whole-cell Sensing

biosensor for the detection of hydroxylated polychlorinated biphenyls in biological and environmental samples is currently under development in our laboratory. Although whole-cell sensing systems have been mainly developed for environmental monitoring purposes, detection of biologically relevant molecules, such as sugars, has benefited from the analytical performance of reporter gene-based whole-cell biosensors [48, 97, 98]. Whole-cell sensing systems for the detection of quorumsensing signaling molecules and antibiotics have been also developed. These are described in more detail in the following sections of this chapter. 6.3 Quorum-sensing Signaling Molecules

Quorum sensing is a phenomenon that enables certain bacteria to communicate with each other by producing and responding to secreted signaling molecules in proportion to cell density [99]. This process allows a population of bacteria to regulate expression of specialized genes in response to changes in its size. When cell population reaches a critical size, certain genes are expressed, hence the term quorum sensing. This phenomenon is responsible for many processes, including expression and secretion of virulence factors, formation of biofilms, competence, conjugation, antibiotic production, motility, sporulation, symbiosis, and bioluminescence [99]. Several quorum-sensing regulatory systems have been identified so far. The best-characterized are those of the LuxI/LuxR type found in most gramnegative bacteria. These quorum-sensing circuits use N-acyl-homoserine lactones (AHLs) as signaling molecules and contain regulatory proteins with homology to the regulatory proteins LuxI and LuxR, which control bioluminescence in the marine bacterium Vibrio fischeri [100]. The gene sequences of some of these regulatory systems have been cloned and inserted in plasmid reporter vectors, thus allowing the development of whole-cell sensing systems for the detection of quorum-sensing signaling molecules. Whole cell-based sensors that contain the luxCDABE cassette as a reporter in gene fusion with quorum-sensing regulatory sequences have been employed to study structure-activity relationships of AHLs [101]. Such biosensing systems have limits of detection in the nanomolar range. They have been applied successfully to the study of AHLs in sputum of cystic fibrosis patients with bacterial lung infections [102]. Whole-cell sensing systems may also represent a tool to identify AHL-producing bacteria by detecting AHLs in their cell-free culture supernatants. Using this approach, it has been shown that Porphyromonas gingivalis, a gramnegative bacterium implicated in the etiology of human periodontal disease, actually regulates the production of virulence factors by quorum sensing but does not utilize AHL-mediated systems [103]. In another study, it has been reported that plant growth-promoting Pseudomonas putida produces cyclic dipeptides that potentially cross talk with the LuxI/LuxR quorum-sensing system [104]. In general, these biosensors can be used as a preliminary screening tool to identify active samples, which can be then subjected to mass spectrometric and NMR analysis in order to determine the chemical identity of individual molecules.

6 Applications

Recently, the molecular mechanisms of bacterial quorum sensing have been proposed as new drug target. In fact, the pharmacological inhibition of quorum sensing for the treatment of bacterial infections may be an alternative to traditional antibiotic treatments, whose therapeutic efficacy is currently decreasing as a result of the occurrence of multiple antibiotic-resistant pathogenic bacteria [105]. Although physical-chemical methods for the detection of AHLs have been developed [106], detailed investigations of quorum-sensing inhibition by methods such as HPLC, GC-MS, or TLC are not practical for routine screening. Whole-cell sensing systems are capable of easily, rapidly, and inexpensively investigating large numbers of compounds and thus are suitable for high-throughput screening of quorum-sensing inhibitors as potential antimicrobial drugs. A GFP-based whole-cell sensing system was developed to perform single-cell analysis and online studies of AHL-mediated communication among bacteria [107]. In this application, a gfp mutant that encodes a protein with a relatively short half-life was used in order to monitor variations in AHL concentrations in real time. Such a biosensor represents a powerful tool for in vivo studies of cell communication and for in vivo efficacy studies of quorum-sensing inhibitors. Indeed, it has been used to detect the production of AHLs from Pseudomonas aeruginosa in a mouse infection model [108]. A similar biosensor allowed detection of AHL production in laboratorybased P. aeruginosa biofilms and showed the ability of a synthetic compound to penetrate microcolonies and block quorum-sensing signaling in most biofilm cells [109]. GFP-based whole-cell sensing systems combined with flow cytometry analysis have been successfully employed for in situ detection of AHLs in soil [110]. 6.4 Antibiotics

Antibiotics belonging to the tetracycline family are extensively used in the therapy and prophylactic control of bacterial infections in human and veterinary medicine and as food additives for growth promotion in the farming industry. Intensive use of tetracyclines has led to widespread antibiotic resistance in bacterial species. The resistance mechanism genes are located in plasmids that can be efficiently transferred from one strain to another. In addition to the development of drug-resistant pathogens, the use of tetracycline has been associated with problems such as unacceptable levels of drug residues in food products for human consumption and release of drugs into the environment. Control of usage in animal farming is possible by monitoring antibiotic residues in different biological samples. Conventional methods for the detection of tetracycline residues include microbial inhibition tests, immunoassays, and chromatographic methods. Recently, whole-cell sensing systems have been proposed as a sensitive, simple, fast, and inexpensive method for measuring tetracyclines in biological fluids and food samples [111]. A bioluminescent bacterial strain containing the bacterial luciferase operon luxCDABE under the control of the tetracycline-responsive element, which is part of the regulation unit of the tetracycline resistance factor, has been developed. It allows for the specific detection of different clinically relevant tetracyclines, with limits of

15

16 Photoproteins as Reporters in Whole-cell Sensing

detection at picomole levels and an induction time of 90 min [112]. Moreover, freeze-dried cells can detect tetracyclines as sensitively as freshly cultivated cells, thus envisioning the possibility of on-site use. The developed whole-cell sensing system has been tested on different biological samples. It has proved to be suitable to screen veterinary serum samples for tetracycline residues in real time [113] and to detect tetracyclines in raw bovine milk samples below the official limits set by the European Union and the U.S. Food and Drug Administration [114]. The biosensing strain was also applied to the detection of traces of tetracyclines in fish, after simple sample preparation. In this case, tetracycline residues were detected below official limits, and the results correlated with those obtained by conventional HPLC [115]. In another study, a GFP-based whole-cell sensor system for in situ detection of tetracyclines was developed. The biosensor was used for qualitative detection of oxytetracycline production by the bacterium Streptomyces rimosus in soil microcosms [116]. The tetracycline-induced GFP-producing biosensor cells were detected by using flow cytometry and fluorescence-activated cell-sorting (FACS) analysis. This study showed the biosensor potential for microbial ecology studies. The same sensor strain was employed in a separate study for in vivo detection and quantification of tetracyclines in rat intestine [117]. Bioavailable tetracycline concentration within the bacterial growth habitat of the intestine proved to be proportional to the intake concentration, but significantly lower. This finding, made possible by the use of the whole-cell sensing system, may help to clarify and optimize antimicrobial therapy in the intestinal environment.

7 Technological Advances

An emerging trend of research in luminescent whole-cell biosensors is moving toward integration of the bacterial reporter strain into the appropriate detection device. This goal is achieved mainly by immobilizing the cells onto optical fiber tips. Different immobilization methods that utilize membranes or alginate, solid agar, and sol-gel matrices have been developed and optimized. The obtained fiber-optic, whole-cell sensors have been proposed as self-contained, disposable biomonitoring devices for environmental analysis [59, 118–121]. When coupled to a portable photon-counting system, they allow performance of measurements in the field. Additionally, the use of multiple fibers enables multiple detection [122]. An interesting application of optical fiber led to the development of a high-density cell assay platform [123]. Here, fluorescent reporter cells were placed into ordered microwell arrays that were fabricated on an optical imaging fiber. Fluorescence signals from individual cells were detected and spatially located using an imaging detector. The main advantage of this cell array technology is the simultaneous analysis of multiple responses from a large number of different strains, thus showing potential for high-throughput screening applications. A different approach to integration is the implementation of chip-based systems. An example of a whole-cell biochip is the bioluminescent bioreporter

References

integrated circuit (BBIC), in which recombinant bioreporter cells are interfaced with an integrated circuit. In this application, the cells are either immobilized or encapsulated to integrate them into the circuit. Alternatively, a flow system is employed to bring the cell bioreporters in proximity with the photodiode-containing chip of a microluminometer optimized for very low-level luminescence detection [124, 125]. Further developments in whole-cell sensing aim toward system miniaturization, with consequent implementation of high-density assay platforms, including 96and 384-well microtiter plates, microarrays, and microfluidics devices. The main advantage of these analytical formats is that they allow for high-throughput screening of samples. Not only can large numbers of samples be analyzed, but also large amounts of information from each sample can be assessed simultaneously and in a relatively short time. Additionally, miniaturized systems allow decreased reagent consumption, sample volume, and analysis time. Several whole cell-based assay systems that use multi-well platforms have been developed. Reporter cells can be suspended in culture media within the wells [126], immobilized in agar matrices at the bottom of single wells [127], or encapsulated in sol-gel matrices that form thick silicate films [128]. The main advantage of immobilized cells is that, while maintaining the analytical characteristics of free cells, they can be used under continuous flow conditions for on-line monitoring and as multiple-use sensing elements. In that regard, the optimization of deposition and immobilization techniques can contribute significantly to the advancement of whole-cell sensing technology [129]. An example of whole-cell microarray is the Lux-Array, which includes hundreds of E. coli reporter strains, high-density printed to membranes on agar plates [130]. The system was developed for genome-wide transcriptional analysis by fusing a collection of E. coli functional promoters to a reporter gene. However, it shows the potential for assembling an array of reportersensing strains to be used for analytical purposes. Recently, we took advantage of microfluidics to develop a miniaturized whole cell-based sensing system [71]. A recombinant reporter strain for the detection of arsenite/antimonite was adapted to a microcentrifugal microfluidics platform in the shape of a compact disk. Centrifugal force controlled valving, reagent flow, and mixing and drove fluids through microfabricated channels and reservoirs. The force was generated by a motor that spun the disk at programmed velocities. Fluorescence from the reporter was detected using a fiber-optic probe. Despite miniaturization, the biosensing system retained its analytical performance and was able to perform quantitative analysis within short response times. Moreover, the developed system has the potential for portability and applicability in the field.

Acknowledgments

This work was supported in part by the National Institute of Environmental Health Sciences (Grant P42 ES 007380) and the National Science Foundation (Grant CHE0416553).

17

18 Photoproteins as Reporters in Whole-cell Sensing

References 1 NAYLOR, L. H. Reporter gene technology: the future looks bright. Biochem. Pharmacol. 1999, 58, 749–757. 2 WOOD, K. V. Marker proteins for gene expression. Curr. Opin. Biotechnol. 1995, 6, 50–58. 3 JANSSON, J. K. Marker and reporter genes: illuminating tools for environmental microbiologists. Curr. Opin. Microbiol. 2003, 6, 310–316. 4 BOUSSE, L. Whole cell biosensors. Sens. Actuators B Chem. 1996, 34, 270–275. 5 PANCRAZIO, J. J., WHELAN, J. P., BORKHOLDER, D. A., MA, W., STENGER, D. A. Development and application of cell-based biosensors. Ann. Biomed. Eng. 1999, 27, 697–711. 6 Biosensors Fundamentals and Applications (TURNER, A. P. F., KARUBE, I., WILSON, G. S., Eds.), Oxford University Press: Oxford, UK, 1987. 7 Biosensor Principles and Applications (BLUM, L. J., COULET, P. R., Eds.), Marcel Dekker: New York, 1991. 8 D’SOUZA, S. F. Microbial biosensors. Biosens. Bioelectron. 2001, 16, 337–353. 9 van der MEER, J. R., TROPEL, D., JASPERS, M. Illuminating the detection chain of bacterial bioreporters. Environ. Microbiol. 2004, 6, 1005–1020. 10 DAUNERT, S., BARRETT, G., FELICIANO, J. S., SHETTY, R. S., SHRESTHA, S., SMITH-SPENCER, W. Genetically engineered whole-cell sensing systems: coupling biological recognition with reporter genes. Chem. Rev. 2000, 100, 2705–2738. 11 DURICK, K., NEGULESCU, P. Cellular biosensors for drug discovery. Biosens. Bioelectron. 2001, 16, 587–592. 12 WINDAL, I., DENISON, M. S., BIRNBAUM, L. S., Van WOUWE, N., BAEYENS, W., GOEYENS, L. Chemically activated luciferase gene expression (CALUX) cell bioassay analysis for the estimation of dioxin-like activity: critical parameters of the CALUX procedure that impact assay results. Environ. Sci. Technol. 2005, 39, 7357–7364. 13 ROGERS, J. M., DENISON, M. S. Recombinant cell bioassays for

14

15

16

17 18

19

20

21

22

23

endocrine disruptors: development of a stably transfected human ovarian cell line for the detection of estrogenic and anti-estrogenic chemicals. In Vitr. Mol. Toxicol. 2000, 13, 67–82. BALDWIN, T. O., CHRISTOPHER, J. A., RAUSHEL, F. M., SINCLAIR, J. F., ZIEGLER, M. M., FISHER, A. J., RAYMENT, I. Structure of bacterial luciferase. Curr. Opin. Struct. Biol. 1995, 5, 798–809. MEIGHEN, E. A. Molecular biology of bacterial bioluminescence. Microbiol. Rev. 1991, 55, 123–142. RUTTER, G. A., KENNEDY, H. J., WOOD, C. D., WHITE, M. R. H., TAVARE, J. M. Real-time imaging of gene expression in single living cells. Chem. Biol. 1998, 5, R285–R290. MEIGHEN, E. Bioluminescence, Bacterial. Encyclo. Microbiol. 1992, 1, 309–319. BILLARD, P., DUBOW, M. S. Bioluminescence-based assays for detection and characterization of bacteria and chemicals in clinical laboratories. Clin. Biochem. 1998, 31, 1–14. BULICH, A. A practical and reliable method for monitoring the toxicity of aquatic samples. Process Biochem. 1982, 17, 45–47. HERMENS, J., BUSSER, F., LEEUWANGH, P., MUSCH, A. Quantitative structure-activity relationships and mixture toxicity of organic chemicals in Photobacterium phosphoreum: the Microtox test. Ecotoxicol. Environ. Saf. 1985, 9, 17–25. GUPTA, R. K., PATTERSON, S. S., RIPP, S., SIMPSON, M. L., SAYLER, G. S. Expression of the Photorhabdus luminescens lux genes (luxA, B, C, D, and E) in Saccharomyces cerevisiae. FEMS Yeast Res. 2003, 4, 305–313. KELLY, C. J., TUMSAROJ, N., LAJOIE, C. A. Assessing wastewater metal toxicity with bacterial bioluminescence in a bench-scale wastewater treatment system. Water Res. 2004, 38, 423–431. DEWET, J. R., WOOD, K. V., DELUCA, M., HELINSKI, D. R., SUBRAMANI, S. Firefly luciferase gene – structure and

References

24

25

26

27

28

29

30

31

32

expression in mammalian cells. Mol. Cell Biol. 1987, 7, 725–737. DEWET, J. R., WOOD, K. V., HELINSKI, D. R., DELUCA, M. Cloning of firefly luciferase cDNA and the expression of active luciferase in Escherichia coli. Proc. Natl. Acad. Sci. USA 1985, 82, 7870–7873. CRAIG, F. F., SIMMONDS, A. C., WATMORE, D., MCCAPRA, F., WHITE, M. R. Membrane-permeable luciferin esters for assay of firefly luciferase in live intact cells. Biochem. J. 1991, 276 (Pt 3), 637–641. WOOD, K. V., LAM, Y. A., SELIGER, H. H., MCELROY, W. D. Complementary DNA coding click beetle luciferases can elicit bioluminescence of different colors. Science 1989, 244, 700–702. LORENZ, W. W., MCCANN, R. O., LONGIARU, M., CORMIER, M. J. Isolation and expression of a cDNA encoding Renilla reniformis luciferase. Proc. Natl. Acad. Sci. U S A 1991, 88, 4438–4442. STABLES, J., SCOTT, S., BROWN, S., ROELANT, C., BURNS, D., LEE, M. G., REES, S. Development of a dual glow-signal firefly and Renilla luciferase assay reagent for the analysis of g-protein coupled receptor signalling. J. Recept. Signal Transduct. Res. 1999, 19, 395–410. INOUYE, S., NOGUCHI, M., SAKAKI, Y., TAKAGI, Y., MIYATA, T., IWANAGA, S., TSUJI, F. I. Cloning and sequence analysis of cDNA for the luminescent protein aequorin. Proc. Natl. Acad. Sci. USA 1985, 82, 3154–3158. OHMIYA, Y., HIRANO, T. Shining the light: the mechanism of the bioluminescence reaction of calcium-binding photo-proteins. Chem. Biol. 1996, 3, 337–347. SHIMOMURA, O., JOHNSON, F. H., SAIGA, Y. Extraction, purification and properties of aequorin, a bioluminescent protein from the luminous hydromedusan, Aequorea. J. Cell Comp. Physiol. 1962, 59, 223–239. RIDER, T., PETROVICK, M., YOUNG, A., SMITH, L., MATHEWS, R., PALMACCI, S., HOLLIS, M., CHEN, J. NASA/NCI Biomedical Imaging Symposium, Bethesda, MD, 1999.

33 RIDER, T. H., PETROVICK, M. S., NARGI, F. E., HARPER, J. D., SCHWOEBEL, E. D., MATHEWS, R. H., BLANCHARD, D. J., BORTOLIN, L. T., YOUNG, A. M., CHEN, J. Z., HOLLIS, M. A. A B cell-based sensor for rapid identification of pathogens. Science 2003, 301, 213–215. 34 PHILLIPS, G. N. Jr. Structure and dynamics of green fluorescent protein. Curr. Opin. Struct. Biol. 1997, 7, 821–827. 35 WELSH, S., KAY, S. A. Reporter gene expression for monitoring gene transfer. Curr. Opin. Biotechnol. 1997, 8, 617–622. 36 HANSEN, L. H., SØRENSEN, S. J. The use of whole-cell biosensors to detect and quantify compounds or conditions affecting biological systems. Microb. Ecol. 2001, 42, 483–494. 37 LEWIS, J. C., DAUNERT, S. Photoproteins as luminescent labels in binding assays. Fresenius J. Anal. Chem. 2000, 366, 760–768. 38 MARCH, J. C., RAO, G., BENTLEY, W. E. Biotechnological applications of green fluorescent protein. Appl. Microbiol. Biotechnol. 2003, 62, 303–315. 39 MATZ, M. V., FRADKOV, A. F., LABAS, Y. A., SAVITSKY, A. P., ZARAISKY, A. G., MARKELOV, M. L., LUKYANOV, S. A. Fluorescent proteins from nonbioluminescent Anthozoa species. Nat Biotechnol. 1999, 17, 969–973. 40 DAVEY, H. M., KELL, D. B. Flow cytometry and cell sorting of heterogeneous microbial populations: the importance of single-cell analyses. Microbiol. Rev. 1996, 60, 641–696. 41 GROSS, L. A., BAIRD, G. S., HOFFMAN, R. C., BALDRIDGE, K. K., TSIEN, R. Y. The structure of the chromophore within DsRed, a red fluorescent protein from coral. Proc. Natl. Acad. Sci. USA 2000, 97, 11990–11995. 42 HAKKILA, K., MAKSIMOW, M., KARP, M., VIRTA, M. Reporter genes lucFF, luxCDABE, gfp, and dsred have different characteristics in whole-cell bacterial sensors. Anal. Biochem. 2002, 301, 235–242. 43 MITCHELL, R. J., GU, M. B. Construction and Characterization of Novel Dual Stress-Responsive Bacterial Biosensors. Biosens. Bioelectron. 2004, 19, 977–985.

19

20 Photoproteins as Reporters in Whole-cell Sensing

44 MOLINA, A., CARPEAUX, R., MARTIAL, J. A., MULLER, M. A transformed fish cell line expressing a green fluorescent protein-luciferase fusion gene responding to cellular stress. Toxicol. In Vitro 2002, 16, 201–207. 45 SAGI, E., HEVER, N., ROSEN, R., BARTOLOME, A. J., PREMKUMAR, J. R., ULBER, R., LEV, O., SCHEPER, T., BELKIN, S. Fluorescence and bioluminescence reporter functions in genetically modified bacterial sensor strains. Sens. Actuators B Chem. 2003, 90, 2–8. 46 BAUMSTARK-KHAN, C., KHAN, R. A., RETTBERG, P., HORNECK, G. Bacterial Lux-Fluoro test for biological assessment of pollutants in water samples from urban and rural origin. Anal. Chim. Acta 2003, 487, 51–60. 47 WOOD, K. V. The chemical mechanism and evolutionary development of beetle bioluminescence. Photochem. Photobiol. 1995, 62, 662–673. 48 SHRESTHA, S., SHETTY, R. S., RAMANATHAN, S., DAUNERT, S. Simultaneous Detection of Analytes Based on Genetically Engineered Whole Cell Sensing Systems. Anal. Chim. Acta. 2001, 444, 251–260. 49 MIRASOLI, M., FELICIANO, J., MICHELINI, E., DAUNERT, S., RODA, A. Internal response correction for fluorescent whole-cell biosensors. Anal. Chem. 2002, 74, 5948–5953. 50 PEDAHZUR, R., POLYAK, B., MARKS, R. S., BELKIN, S. Water toxicity detection by a panel of stress-responsive luminescent bacteria. J. Appl. Toxicol. 2004, 24, 343–348. 51 GU, M. B., GIL, G. C., KIM, J. H. Enhancing the sensitivity of a two-stage continuous toxicity monitoring system through the manipulation of the dilution rate. J. Biotechnol. 2002, 93, 283–288. 52 PREMKUMAR, J. R., LEV, O., MARKS, R. S., POLYAK, B., ROSEN, R., BELKIN, S. Antibody-based immobilization of bioluminescent bacterial sensor cells. Talanta 2001, 55, 1029–1038. 53 KIM, B. C., GU, M. B. A bioluminescent sensor for high throughput toxicity classification. Biosens. Bioelectron. 2003, 18, 1015–1021.

54 CHOI, S. H., GU, M. B. A portable toxicity biosensor using freeze-dried recombinant bioluminescent bacteria. Biosens. Bioelectron. 2002, 17, 433–440. 55 GU, M. B., GIL, G. C. A multi-channel continuous toxicity monitoring system using recombinant bioluminescent bacteria for classification of toxicity. Biosens. Bioelectron. 2001, 16, 661–666. 56 CHOI, S. H., GU, M. B. Toxicity biomonitoring of degradation byproducts using freeze-dried recombinant bioluminescent bacteria. Anal. Chim. Acta 2003, 481, 229–238. 57 KIM, B. C., PARK, K. S., KIM, S. D., GU, M. B. Evaluation of a high throughput toxicity biosensor and comparison with a Daphnia magna bioassay. Biosens. Bioelectron. 2003, 18, 821–826. 58 GU, M. B., MIN, J., LAROSSA, R. A. Bacterial bioluminescent emission from recombinant Escherichia coli harboring a reca::luxcdabe fusion. J. Biochem. Biophys. Methods 2000, 45, 45–56. 59 POLYAK, B., BASSIS, E., NOVODVORETS, A., BELKIN, S., MARKS, R. S. Bioluminescent whole cell optical fiber sensor to genotoxicants: system optimization. Sens. Actuators B Chem. 2001, 74, 18–26. 60 BARTOLOME, A. J., ULBER, R., SCHEPER, T., SAGI, E., BELKIN, S. Genotoxicity monitoring using a 2D-spectroscopic GFP whole cell biosensing system. Sens. Actuators B Chem. 2003, 89, 27–32. 61 SILVER, S. Genes for all metals – a bacterial view of the Periodic Table. The 1996 Thom Award Lecture. J. Ind. Microbiol. Biotechnol. 1998, 20, 1–12. 62 ROSEN, B. P. Transport and detoxification systems for transition metals, heavy metals and metalloids in eukaryotic and prokaryotic microbes. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 2002, 133, 689–693. 63 PARALES, R. E., HADDOCK, J. D. Biocatalytic degradation of pollutants. Curr. Opin. Biotechnol. 2004, 15, 374–379. 64 BURLAGE, R. S., SAYLER, G. S., LARIMER, F. Monitoring of naphthalene catabolism by bioluminescence with nah-lux transcriptional fusions. J. Bacteriol. 1990, 172, 4749–4757.

References 65 BELKIN, S. Microbial whole-cell sensing systems of environmental pollutants. Curr. Opin. Microbiol. 2003, 6, 206–212. 66 GU, M. B., MITCHELL, R. J., KIM, B. C. Whole-cell-based biosensors for environmental biomonitoring and application. Adv. Biochem. Engin./Biotechnol. 2004, 87, 269–305. 67 RAMANATHAN, S., SHI, W., ROSEN, B. P., DAUNERT, S. Sensing antimonite and arsenite at the subattomole level with genetically engineered bioluminescent bacteria. Anal. Chem. 1997, 69, 3380–3384. 68 TAURIAINEN, S., KARP, M., CHANG, W., VIRTA, M. Recombinant luminescent bacteria for measuring bioavailable arsenite and antimonite. Appl. Environ. Microbiol. 1997, 63, 4456–4461. 69 TAURIAINEN, S., VIRTA, M., CHANG, W., KARP, M. Measurement of firefly luciferase reporter gene activity from cells and lysates using Escherichia coli arsenite and mercury sensors. Anal. Biochem. 1999, 272, 191–198. 70 ROBERTO, F. F., BARNES, J. M., BRUHN, D. F. Evaluation of a gfp reporter gene construct for environmental arsenic detection. Talanta 2002, 58, 181–188. 71 ROTHERT, A., DEO, S. K., MILLNER, L., PUCKETT, L. G., MADOU, M. J., DAUNERT, S. Whole-cell-reporter-gene-based biosensing systems on a compact disk microfluidics platform. Anal. Biochem. 2005, 342, 11–19. 72 TAURIAINEN, S., KARP, M., CHANG, W., VIRTA, M. Luminescent bacterial sensor for cadmium and lead. Biosens. Bioelectron. 1998, 13, 931–938. 73 RIETHER, K. B., DOLLARD, M. A., BILLARD, P. Assessment of heavy metal bioavailability using Escherichia coli zntAp::lux and copAp::lux-based biosensors. Appl. Microbiol. Biotechnol. 2001, 57, 712–716. 74 SHETTY, R. S., DEO, S. K., SHAH, P., SUN, Y., ROSEN, B. P., DAUNERT, S. Luminescence-based whole-cell-sensing systems for cadmium and lead using genetically engineered bacteria. Anal. Bioanal. Chem. 2003, 376, 11–17. 75 PEITZSCH, N., EBERZ, G., NIES, D. H. Alcaligenes eutrophus as a bacterial

76

77

78

79

80

81

82

83

84

85

chromate sensor. Appl. Environ. Microbiol. 1998, 64, 453–458. VULKAN, R., ZHAO, F. J., BARBOSA-JEFFERSON, V., PRESTON, S., PATON, G. I., TIPPING, E., MCGRATH, S. P. Copper speciation and impacts on bacterial biosensors in the pore water of copper-contaminated soils. Environ. Sci. Technol. 2000, 34, 5115–5121. STOYANOV, J. V., MAGNANI, D., SOLIOZ, M. Measurement of cytoplasmic copper, silver, and gold with a lux biosensor shows copper and silver, but not gold, efflux by the CopA ATPase of Escherichia coli. FEBS Lett. 2003, 546, 391–394. JOYNER, D. C., LINDOW, S. E. Heterogeneity of iron bioavailability on plants assessed with a whole-cell GFP-based bacterial biosensor. Microbiology 2000, 146(Pt 10) 2435–2445. SELIFONOVA, O., BURLAGE, R., BARKAY, T. Bioluminescent sensors for detection of bioavailable Hg(II) in the environment. Appl. Environ. Microbiol. 1993, 59, 3083–3090. VIRTA, M., LAMPINEN, J., KARP, M. A luminescence-based mercury biosensor. Anal. Chem. 1995, 67, 667–669. LYNGBERG, O. K., STEMKE, D. J., SCHOTTEL, J. L., FLICKINGER, M. C. A single-use luciferase-based mercury biosensor using Escherichia coli HB101 immobilized in a latex copolymer film. J. Ind. Microbiol. Biotechnol. 1999, 23, 668–676. HANSEN, L. H., SØRENSEN, S. J. Versatile biosensor vectors for detection and quantification of mercury. FEMS Microbiol. Lett. 2000, 193, 123–127. RASMUSSEN, L. D., SORENSEN, S. J., TURNER, R. R., BARKAY, T. Application of a mer-lux biosensor for estimating bioavailable mercury in soil. Soil Biol. Biochem. 2000, 32, 639–646. IVASK, A., HAKKILA, K., VIRTA, M. Detection of organomercurials with sensor bacteria. Anal. Chem. 2001, 73, 5168–5171. TIBAZARWA, C., CORBISIER, P., MENCH, M., BOSSUS, A., SOLDA, P., MERGEAY, M., WYNS, L., van der LELIE, D. A Microbial

21

22 Photoproteins as Reporters in Whole-cell Sensing

86

87

88

89

90

91

92

93

biosensor to predict bioavailable nickel in soil and its transfer to plants. Environ. Pollut. 2001, 113, 19–26. STICHER, P., JASPERS, M. C. M., STEMMLER, K., HARMS, H., ZEHNDER, A. J. B., van der MEER, J. R. Development and characterization of a whole-cell bioluminescent sensor for bioavailable middle-chain alkanes in contaminated ground water samples. Appl. Environ. Microbiol. 1997, 63, 4053–4060. PHOENIX, P., KEANE, A., PATEL, A., BERGERON, H., GHOSHAL, S., LAU, P. C. K. Characterization of a new solvent-responsive gene locus in Pseudomonas putida F1 and its functionalization as a versatile biosensor. Environ. Microbiol. 2003, 5, 1309–1327. KOBATAKE, E., NIIMI, T., HARUYAMA, T., IKARIYAMA, Y., AIZAWA, M. Biosensing of benzene derivatives in the environment by luminescent Escherichia coli. Biosens. Bioelectron. 1995, 10, 601–605. APPLEGATE, B. M., KEHRMEYER, S. R., SAYLER, G. S. A chromosomally based tod-luxCDABE whole-cell reporter for benzene, toluene, ethybenzene, and xylene (BTEX) sensing. Appl. Environ. Microbiol. 1998, 64, 2730–2735. STINER, L., HALVERSON, L. J. Development and characterization of a green fluorescent protein-based bacterial biosensor for bioavailable toluene and related compounds. Appl. Environ. Microbiol. 2002, 68, 1962–1971. HEITZER, A., MALACHOWSKY, K., THONNARD, J. E., BIENKOWSKI, P. R., WHITE, D. C., SAYLER, G. S. Optical biosensor for environmental on-line monitoring of naphthalene and salicylate bioavailability with an immobilized bioluminescent catabolic reporter bacterium. Appl. Environ. Microbiol. 1994, 60, 1487–1494. ABD-EL-HALEEM, D., RIPP, S., SCOTT, C., SAYLER, G. S. A luxCDABE-based bioluminescent bioreporter for the detection of phenol. J. Ind. Microbiol. Biotechnol. 2002, 29, 233–237. LAYTON, A. C., MUCCINI, M., GHOSH, M. M., SAYLER, G. S. Construction of a bioluminescent reporter strain to detect

94

95

96

97

98

99

100

101

102

103

polychlorinated biphenyls. Appl. Environ. Microbiol. 1998, 64, 5023–5026. MBEUNKUI, F., RICHAUD, C., ETIENNE, A. L., SCHMID, R. D., BACHMANN, T. T. Bioavailable nitrate detection in water by an immobilized luminescent cyanobacterial reporter strain. Appl. Microbiol. Biotechnol. 2002, 60, 306–312. TAYLOR, C. J., BAIN, L. A., RICHARDSON, D. J., SPIRO, S., RUSSELL, D. A. Construction of a whole-cell gene reporter for the fluorescent bioassay of nitrate. Anal. Biochem. 2004, 328, 60–66. DOLLARD, M. A., BILLARD, P. Whole-cell bacterial sensors for the monitoring of phosphate bioavailability. J. Microbiol. Methods 2003, 55, 221–229. SHETTY, R. S., RAMANATHAN, S., BADR, I. H. A., WOLFORD, J. L., DAUNERT, S. Green fluorescent protein in the design of a living biosensing system for L-arabinose. Anal. Chem. 1999, 71, 763–768. ˜ ONES, MILLER, W. G., BRANDL, M. T., QUIN B., LINDOW, S. E. Biological sensor for sucrose availability: relative sensitivities of various reporter genes. Appl. Environ. Microbiol. 2001, 67, 1308–1317. MILLER, M. B., BASSLER, B. L. Quorum sensing in bacteria. Annu. Rev. Microbiol. 2001, 55, 165–199. FUQUA, C., PARSEK, M. R., GREENBERG, E. P. Regulation of gene expression by cell-to-cell communication: acylhomoserine lactone quorum sensing. Annu. Rev. Genet. 2001, 35, 439–468. WINSON, M. K., SWIFT, S., FISH, L., THROUP, J. P., JØRGENSEN, F., CHHABRA, S. R., BYCROFT, B. W., WILLIAMS, P., STEWART, G. S. A. B. Construction and analysis of luxCDABE-based plasmid sensors for investigating N-acyl homoserine lactone-mediated quorum sensing. FEMS Microbiol. Lett. 1998, 163, 185–192. MIDDLETON, B., RODGERS, H. C., C´AMARA, M., KNOX, A. J., WILLIAMS, P., HARDMAN, A. Direct detection of N-acylhomoserine lactones in cystic fibrosis sputum. FEMS Microbiol. Lett. 2002, 207, 1–7. BURGESS, N. A., KIRKE, D. F., WILLIAMS, P., WINZER, K., HARDIE, K. R., MEYERS, N. L., ADUSE-OPOKU, J., CURTIS, M. A.,

References

104

105

106

107

108

109

110

C´AMARA, M. LuxS-dependent quorum sensing in Porphyromonas gingivalis modulates protease and haemagglutinin activities but is not essential for virulence. Microbiology 2002, 148, 763–772. DEGRASSI, G., AGUILAR, C., BOSCO, M., ZAHARIEV, S., PONGOR, S., VENTURI, V. Plant growth-promoting Pseudomonas putida WCS358 produces and secretes four cyclic dipeptides: cross-talk with quorum sensing bacterial sensors. Curr. Microbiol. 2002, 45, 250–254. HENTZERM, M., GIVSKOV, M. Pharmacological inhibition of quorum sensing for the treatment of chronic bacterial infections. J. Clin. Invest. 2003, 112, 1300–1307. MORIN, D., GRASLAND, B., VALLE´ E-R´EHEL, K., DUFAU, C., HARAS, D. On-line high-performance liquid chromatography-mass spectrometric detection and quantification of N-acyl-homoserine lactones, quorum sensing signal molecules, in the presence of biological matrices. J. Chromatogr. A 2003, 1002, 79–92. ANDERSEN, J. B., HEYDORN, A., HENTZER, M., EBERL, L., GEISENBERGER, O., CHRISTENSEN, B. B., MOLIN, S., GIVSKOV, M. gfp-Based N-acyl homoserine-lactone sensor systems for detection of bacterial communication. Appl. Environ. Microbiol. 2001, 67, 575–585. WU, H., SONG, Z., HENTZER, M., ANDERSEN, J. B., HEYDORN, A., MATHEE, K., MOSER, C., EBERL, L., MOLIN, S., HØIBY, N., GIVSKOV, M. Detection of N-acylhomoserine lactones in lung tissues of mice infected with Pseudomonas aeruginosa. Microbiology 2000, 146, 2481–2493. HENTZER, M., RIEDEL, K., RASMUSSEN, T. B., HEYDORN, A., ANDERSEN, J. B., PARSEK, M. R., RICE, S. A., EBERL, L., MOLIN, S., HØIBY, N., KJELLEBERG, S., GIVSKOV, M. Inhibition of quorum sensing in Pseudomonas aeruginosa biofilm bacteria by a halogenated furanone compound. Microbiology 2002, 148, 87–102. BURMØLLE, M., HANSEN, L. H., OREGAARD, G., SØRENSEN, S. J. Presence of N-acyl homoserine lactones in soil detected by a

111

112

113

114

115

116

117

118

119

whole-cell biosensor and flow cytometry. Microb. Ecol. 2003, 45, 226–236. HANSEN, L. H., SØRENSEN, S. J. Detection and quantification of tetracyclines by whole cell biosensors. FEMS Microbiol. Lett. 2000, 190, 273–278. KORPELA, M. T., KURITTU, J. S., KARVINEN, J. T., KARP, M. T recombinant Escherichia coli sensor strain for the detection of tetracyclines. Anal. Chem. 1998, 70, 4457–4462. KURITTU, J., KARP, M., KORPELA, M. Detection of tetracyclines with luminescent bacterial strains. Luminescence 2000, 15, 291–297. KURITTU, J., L¨ONNBERG, S., VIRTA, M., KARP, M. A group-specific microbiological test for the detection of tetracycline residues in raw milk. J. Agric. Food Chem. 2000, 48, 3372–3377. PELLINEN, T., BYLUND, G., VIRTA, M., NIEMI, A., KARP, M. Detection of traces of tetracyclines from fish with a bioluminescent sensor strain incorporating bacterial luciferase reporter genes. J. Agric. Food Chem. 2002, 50, 4812–4815. HANSEN, L. H., FERRARI, B., SØRENSEN, A. H., VEAL, D., SØRENSEN, S. J. Detection of oxytetracycline production by Streptomyces rimosus in soil microcosms by combining whole-cell biosensors and flow cytometry. Appl. Environ. Microbiol. 2001, 67, 239–244. BAHL, M. I., HANSEN, L. H., LICHT, T. R., SØRENSEN, S. J. In vivo detection and quantification of tetracycline by use of a whole-cell biosensor in the rat intestine. Antimicrob. Agents Chemother. 2004, 48, 1112–1117. IKARIYAMA, Y., NISHIGUCHI, S., KOYAMA, T., KOBATAKE, E., AIZAWA, M., TSUDA, M., NAKAZAWA, T. Fiber-optic-based biomonitoring of benzene derivatives by recombinant E. coli bearing luciferase gene-fused TOL-plasmid immobilized on the fiber-optic end. Anal. Chem. 1997, 69, 2600–2605. POLYAK, B., BASSIS, E., NOVODVORETS, A., BELKIN, S., MARKS, R. S. Optical fiber bioluminescent whole-cell microbial biosensors to genotoxicants. Water Sci. Technol. 2000, 42, 305–311.

23

24 Photoproteins as Reporters in Whole-cell Sensing

120 GIL, G. C., KIM, Y. J., GU, M. B. Enhancement in the sensitivity of a gas biosensor by using an advanced immobilization of a recombinant bioluminescent bacterium. Biosens. Bioelectron. 2002, 17, 427–432. 121 SUN, Y., ZHOU, T., GUO, J., LI, Y. Dark variants of luminous bacteria whole cell bioluminescent optical fiber sensor to genotoxicants. J. Huazhong Univ. Sci. Technolog. Med. Sci. 2004, 24, 507–509. 122 HAKKILA, K., GREEN, T., LESKINEN, P., IVASK, A., MARKS, R., VIRTA, M. Detection of bioavailable heavy metals in EILATox-Oregon samples using whole-cell luminescent bacterial sensors in suspension or immobilized onto fibre-optic tips. J. Appl. Toxicol. 2004, 24, 333–342. 123 BIRAN, I., WALT, D. R. Optical imaging fiber-based single live cell arrays: a high-density cell assay platform. Anal. Chem. 2002, 74, 3046–3054. 124 SIMPSON, M. L., SAYLER, G. S., PATTERSON, G., NIVENS, D. E., BOLTON, E. K., ROCHELLE, J. M., ARNOTT, J. C., APPLEGATE, B. M., RIPP, S., GUILLORN, M. A. An integrated CMOS microluminometer for low-level luminescence sensing in the bioluminescent bioreporter integrated circuit. Sens. Actuators B Chem. 2001, 72, 134–140.

125 BOLTON, E. K., SAYLER, G. S., NIVENS, D. E., ROCHELLE, J. M., RIPP, S., SIMPSON, M. L. Integrated CMOS photodetectors and signal processing for very low-level chemical sensing with the bioluminescent bioreporter integrated circuit. Sens. Actuators B Chem. 2002, 85, 179–185. 126 RODA, A., PASINI, P., MIRASOLI, M., GUARDIGLI, M., RUSSO, C., MUSIANI, M., BARALDINI, M. Sensitive determination of urinary mercury(II) by a bioluminescent transgenic bacteria-based biosensor. Anal. Letters 2001, 34, 29–41. 127 KIM, B. C., GU, M. B. A bioluminescent sensor for high throughput toxicity classification. Biosens. Bioelectron. 2003, 18, 1015–1021. 128 PREMKUMAR, J. R., ROSEN, R., BELKIN, S., LEV, O. Sol-gel luminescence biosensors: encapsulation of recombinant E. coli reporters in thick silicate films. Anal. Chim. Acta 2002, 462, 11–23. 129 BARRON, J. A., ROSEN, R., JONES-MEEHAN, J., SPARGO, B. J., BELKIN, S., RINGEISEN, B. R. Biological laser printing of genetically modified Escherichia coli for biosensor applications. Biosens. Bioelectron. 2004, 20, 246–252. 130 Van DYK, T. K., DEROSE, E. J., GONYE, G. E. LuxArray, a high-density, genomewide transcription analysis of Escherichia coli using bioluminescent reporter strains. J. Bacteriol. 2001, 183, 5496–5505.

1

Catalytic Antibody Technology Donald Hilvert Swiss Federal Institute of Technology, Z¨urich, Switezerland

Originally published in: Catalytic Antibodies. Edited by Ehud Keinan. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30688-6

1 Introduction

Antibody molecules elicited with rationally designed transition-state analogs catalyze numerous reactions, including many that cannot be achieved by standard chemical methods. Although relatively primitive when compared with natural enzymes, these catalysts are valuable tools for probing the origins and evolution of biological catalysis. Mechanistic and structural analyses of representative antibody catalysts, generated with a variety of strategies for several different reaction types, suggest that their modest efficiency is a consequence of imperfect hapten design and indirect selection. Development of improved transition-state analogs, refinements in immunization and screening protocols, and elaboration of general strategies for augmenting the efficiency of first-generation catalytic antibodies are identified as evident, but difficult, challenges for this field. Rising to these challenges and more successfully integrating programmable design with the selective forces of biology will enhance our understanding of enzymatic catalysis. Further, it should yield useful protein catalysts for an enhanced range of practical applications in chemistry and biology.

2 Exploiting Antibodies as Catalysts

Some three decades ago, Jencks suggested that stable molecules resembling the transition state of a reaction might be used as haptens to elicit antibodies with tailored catalytic activities and selectivities [1]. Implementation of this clever idea was made possible by the development of monoclonal antibodies, viable transition-state analogs, and versatile screening assays, and more than 100 reactions have now been Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Catalytic Antibody Technology

successfully accelerated by antibodies [2, 3]. These include pericyclic processes, group transfer reactions, additions and eliminations, oxidations and reductions, aldol condensations, and miscellaneous cofactor-dependent transformations. Because selectivities in these systems generally reflect the structure of the hapten and can rival those of natural enzymes, transformations that cannot be achieved efficiently or selectively via more traditional chemical methods are the subject of much current research [4]. Total synthesis of the natural product epothilone from a chiral intermediate prepared by antibody catalysis [5] and activation of a prodrug in vivo [6] are two recent accomplishments that illustrate the potential of this technology. Nevertheless, practical applications are still the exception rather than the rule. Low catalytic efficiency, in particular, appears to be a significant limitation. While modest rate accelerations are easily achieved, enzyme-like activity remains elusive. The goal of this review is to address this problem in the light of recent structural and mechanistic work. After first considering the intertwined issues of catalytic efficiency and hapten design, a few well-characterized examples of antibodies promoting a diverse set of reactions are used to illustrate how antibody binding energy is exploited for catalysis and to identify factors that limit overall efficiency. Finally, possible strategies for improving these systems are discussed.

3 Catalytic Efficiency

By almost any criterion, natural enzymes are incredibly efficient catalysts. The fastest enzymes are limited by the rate at which they encounter substrate, and even those that have not achieved this level of evolutionary perfection typically have apparent bi-molecular rate constants (kcat /K m ) between 106 and 108 M−1 s−1 , irrespective of the rate of the corresponding uncatalyzed reaction [7]. Rate accelerations over background (kcat /kuncat ) are also very high, in the range 106 to 1012 , and can even reach 1017 in some special cases [7, 8]. These extraordinary effects have been explained by an enzyme’s ability to bind transition states more tightly than ground states [9]. When enzymatic and nonenzymatic reactions occur by the same mechanism and chemistry is rate determining, a simple thermodynamic cycle based on transition-state theory can be used to show that kcat /kuncat = K m /K TS , whereK m andK TS are the dissociation constants for substrate and transition state from the enzyme [10, 11]. The chemical proficiency of an enzyme, as defined by Wolfenden [7], is then given as the ratio (kcat /K m )/kuncat or 1/K TS . This term represents the lower limit of a protein’s affinity for the transition state and varies from 108 to 1023 M−1 for a series of natural enzymes [7, 8]. Although true transition-state affinity may be underestimated if chemistry is not clearly rate limiting or if the enzyme uses a different mechanism than the uncatalyzed reaction, this term provides a useful measure of catalytic power for the purpose of discussion.

3 Catalytic Efficiency

One practical consequence of the application of transition-state theory to enzymes has been the design of potent inhibitor molecules through mimicry of structural and electronic features of otherwise ephemeral transition states (for recent reviews, see [12–14]). Another has been the use of such compounds to generate catalytic antibodies [2, 3]. The latter strategy has been found to be broadly useful, as any chemical transformation compatible with a biological milieu is potentially amenable to antibody catalysis so long as an appropriate transition-state analog can be devised. Broad scope and high, programmable catalyst selectivity certainly make this route to tailored enzymes one of the most promising to emerge in the last two decades. In terms of efficiency, however, catalytic antibodies do not yet match their natural counterparts. Published kcat /K m values for the best catalysts are only in the range 102 to 104 M−1 s−1 , well below the limit for diffusion-controlled processes, and rate accelerations are usually between 103 - to 105 -fold over background [15, 16]. Smaller effects are often found, but higher efficiencies are extremely rare. Differences between antibodies and enzymes are readily apparent in Fig. 1, where rate acceleration is plotted against chemical proficiency for a representative set of reactions. Antibody-catalyzed reactions cluster in the lower left quadrant of the plot, corresponding to the lowest activities, whereas enzymatic reactions lie above and to the right, spanning a much greater range of efficiency. Because K m values are roughly comparable in all of these systems, an excellent correlation between rate acceleration and proficiency is observed. In essence, better transitionstate binding translates directly into higher activity. Although the best antibodies approach the efficiency of the least efficient enzymes, it should be noted that the

Fig. 1 Comparison of chemical proficiency, -log (K Ts ), and rate acceleration, kcat /kuncat , for a series of reactions catalyzed by enzymes (filled symbols) and antibodies (open symbols). Rearrangements (, ), hydrolyses (,), decarboxylations (, ), de-

protonations (•,◦) and retroaldol reactions (♦, ) are included. Enzyme data are from [7, 112, 131, 132] and antibody data were taken from [25, 26, 38, 75, 91, 110, 113, 115, 121, 127, 200–202].

3

4 Catalytic Antibody Technology

corresponding reactions often involve conversions of relatively activated substrates (e.g., the hydrolysis of aryl esters). In contrast, enzymes specialize in accelerating extremely slow reactions like the cleavage of amides (t1/2 = 7.3 years) and phosphate diesters (t1/2 = 130 000 years), few of which have yielded to significant antibody catalysis. In a complementary analysis, Stewart and Benkovic [15] found a weak correlation between rate acceleration and the ratio of the equilibrium binding constants of the reaction substrate and the hapten: kcat /kuncat = K m /K TS ≈ K m /K i . In other words, affinity for the transition-state analog roughly approximates affinity for the true transition state, in accord with the basic premise underlying the production of catalytic antibodies. Practically speaking, this means that the search for highperformance catalysts can often be reduced to a search for the best hapten binders – provided the hapten is a reasonably good transition-state mimic. However, association constants for low molecular weight haptens (1/K i = 106 –1010 M−1 ) are generally much smaller than chemical proficiencies of the most effective enzymes. Given K m values of ca. 10−4 M, these affinities suggest that kcat /kuncat will seldom exceed ca. 106 for first generation catalytic antibodies, as found experimentally. For reactions with modest activation barriers, effects of this magnitude may be useful. If product inhibition and protein production pose no further technical hurdles, kinetic resolutions can be achieved, the fate of reactive intermediates controlled and so on. For truly difficult reactions, however, such effects are almost certainly inadequate.

4 Hapten Design

Before turning to a description of specific antibodies, a few words about hapten design are in order. Transition states themselves have fleeting lifetimes and cannot be isolated. Synthesis of effective analogs must therefore draw on our chemical intuition about the conformational, stereochemical, and electronic properties of the reaction under study. Because no stable molecule can reproduce all the characteristics of an actual transition state, design efforts have tended to focus on salient features distinguishing transition state from ground state [12–14]. For instance, changes in hybridization or charge occurring as a reaction proceeds can be mimicked by incorporating different elements or charged groups at appropriate sites within the analog. Conformational constraint can help approximate the reactive geometry of a flexible substrate, while multisubstrate analogs are used to imitate the relative disposition of reactants in a bimolecular process. For reactions that require catalytic functionality within the antibody pocket, more sophisticated strategies appear to be needed. In the “bait-and-switch” approach, charge complementarity between hapten and antibody is exploited to induce appropriately positioned acids, bases and nucleophiles. Alternatively, catalytic residues can be selected directly by irreversible chemical modification when mechanismbased inhibitors [17, 18] are employed as haptens. The latter strategy, dubbed

5 Representative Catalytic Antibodies

“reactive immunization” [19], has the virtue of allowing rational engineering of covalent catalysis. For each new reaction, hapten design must be optimized to maximize the probability of finding an antibody catalyst. Because subtle differences between even the best transition-state analogs and actual transition states almost certainly contribute to lower efficiencies of antibodies compared with enzymes, it is important to understand how instructions implicit in any given hapten design are realized in the complementary immunoglobulin binding pocket. Characterization of successful antibody catalysts at the atomic level currently provides the most useful insights into how binding energy is exploited for catalysis.

5 Representative Catalytic Antibodies 5.1 Proximity Effects

Utilization of binding energy to constrain flexible molecules into reactive conformations or to preorganize reactants for bimolecular reaction is a potentially powerful strategy for accelerating reactions with unfavorable entropies of activation [20, 21]. To test whether antibodies might serve as entropy traps [21], concerted pericyclic reactions requiring neither nucleophilic nor acid-base catalysis have been investigated. 5.1.1 Sigmatropic Rearrangements The biologically important Claisen rearrangement of chorismate to prephenate (Fig. 2) and the abiological oxy-Cope rearrangement (Fig. 3) are typical [3,3]-sigmatropic processes. They proceed via highly ordered, entropically unfavorable, cyclic transition states involving the simultaneous formation of a carbon–carbon bond and cleavage of either a carbon-oxygen or another carbon-carbon bond. For the chorismate rearrangement, the conformationally locked oxabicyclic dicarboxylic acid 1 [22] proved to be a successful hapten (Fig. 2). It has a chair-like geometry very similar to that of the presumed transition state and is an effective inhibitor of natural chorismate mutases [22, 23]. Antibodies that bind 1 catalyze the chorismate rearrangement enantioselectively with rate accelerations (kcat /kuncat ) of 102 to 104 over background [24–27]. For comparison, enzymes accelerate this reaction by a factor of ca. 106 [28]. Spectroscopic and X-ray studies of 1F7 (which achieves a 200-fold rate enhancement) have provided insights into the origins of catalysis in these systems. Transferred nuclear Overhauser effects show that 1F7 binds the flexible chorismate molecule in the diaxial conformation specified by the transition-state analog [29]. Crystallographic data confirm that the induced binding pocket faithfully reflects hapten design [30]. Compound 1 is deeply buried in the complex, and the active site’s overall shape and charge are complementary to a single hapten

5

6 Catalytic Antibody Technology

Fig. 2 (Top) The Claisen rearrangement of chorismate to prephenate catalyzed by antibody 1F7; the flexible chorismate adopts an extended pseudo-diequatorial conformation in solution and must undergo a conformational change to populate the less

stable pseudodiaxial conformer in order for reaction to proceed. (Bottom) The conformationally restricted dicarboxylic acid 1 [22] mimics the transition state of the Claisen rearrangement and was used to elicit catalytic antibody 1F7 [25].

Fig. 3 Antibody-catalyzed oxy-Cope rearrangement [36]. The aldehyde product is trapped with hydroxylamine to give an oxime to prevent time dependent inactivation of the catalyst. The 2,5-diaryl cyclohexanol derivative 2 was used to imitate the structure of the pericyclic transition state.

5 Representative Catalytic Antibodies

Fig. 4 (a) Active site of antibody 1F7 [30], showing interactions with hapten 1. The light and heavy chains are pink and blue, respectively; the hapten is yellow. Note that ArgH95 forms a salt bridge with the ligand’s secondary carboxylate but is too far away to form a hydrogen bond with the ether oxygen. (b) Active site of E. coli chorismate

mutase [203]. Bound 1 is completely inaccessible to solvent and makes numerous contacts with protein residues; hydrogen bonds from Lys39 and Gln88 to the ligand’s ether oxygen are essential for high activity [28]. Graphics were prepared with the programs BobScript [204] and Raster3D [205].

enantiomer (Fig. 4a). Consequently, only the corresponding (–)-isomer of chorismate binds in a conformation appropriate for reaction. The subsequent rearrangement of bound substrate then occurs by the same concerted mechanism as that deduced for the uncatalyzed reaction and for natural chorismate mutases. Nevertheless, 1F7 is likely to be a much poorer entropy trap than mutase enzymes. It exploits many fewer hydrogen bonds and electrostatic interactions for ligand recognition (Fig. 4a,b). It also appears to accommodate charge separation in the transition state less effectively. Isotope effects show that the transition state for the rearrangement is highly polarized, with C–O bond cleavage preceding C–C bond formation [31, 32]. The enzymes stabilize this species electrostatically by placing a cationic residue (either Arg or Lys) near the partially negatively-charged ether oxygen of the breaking C–O bond [28, 33, 34], but 1F7 lacks an analogous feature (Fig. 4a,b). These differences presumably explain the antibody’s 104 -fold lower efficiency. They can be attributed, in large part, to shortcomings in hapten design. While 1 reproduces the geometry of the actual transition state reasonably well, it mimics the polarized character of this high energy species poorly [35]. Shortcomings in hapten design are also likely to account for the modest activity of antibody AZ-28. This antibody was raised against cyclohexanol derivative 2

7

8 Catalytic Antibody Technology

(K d = 17 nM) and catalyzes the oxy-Cope rearrangement of the corresponding 2,5diaryl-3-hydroxy-1,5-hexadiene with a kcat /kuncat of 5300 (Fig. 3) [36]. In this case, disposition of the aryl substituents in the transition state is imitated imperfectly in the stable hapten. Like the chorismate mutase antibody, AZ-28 has been shown by TRNOE measurements to preorganize the normally extended hexadiene substrate into a cyclic conformation so that its termini are in close proximity [37]. In this case, ligand recognition is mediated by extensive van der Waals contacts, π-stacking interactions with the aromatic rings, and hydrogen bonding interactions with the alcohol, all evident in the X-ray structure of the antibody-hapten complex (Fig. 5) [38]. However, the conformation adopted by the substrate at the AZ-28 active site is unlikely to be optimal for reaction. The two aryl substituents at C2 and C5 are key recognition elements (Fig. 5) but are oriented very differently in the hapten, where they are sp3 hybridized and equatorial to the plane of the cyclohexane ring, and the transition state, where they are sp2 hybridized and conjugated with the reacting olefins (Fig. 3). Thus, even though the hapten-induced pocket brings together the ends of the hexadiene substrate, binding energy directed to the peripheral aryl groups

Fig. 5 (a) Stereoview of the AZ-28 active site showing hapten-contacting residues [38]. The 5-phenyl ring of 2 sits deep in the hydrophobic pocket, while the 2-phenyl substituent and linker are near the surface. The hapten’s alcohol forms a hydrogen bond

with HisH96 . Two water molecules in the pocket are depicted as red spheres. (b) GRASP [206] surface representation of AZ28 showing the high degree of shape complementarity between ligand and protein.

5 Representative Catalytic Antibodies

almost certainly imposes physical constraints that preclude effective alignment of the reacting [4π + 2σ ] orbitals. Alterations in antibody structure leading to improved orbital overlap should therefore result in significant increases in catalytic efficiency. This inference is supported by studies of AZ-28’s germline precursor. This antibody binds hapten 2 with 40-fold lower affinity than the mature AZ-28 but is a substantially better catalyst, achieving a 163 000-fold rate acceleration over background [38]. Mutagenesis experiments showed that replacement of SerL34 in the germline sequence with Asn is responsible for both effects [38]. In the crystal structure of AZ28, AsnL34 interacts directly with the cyclohexyl ring of the hapten and is therefore in a position to influence the conformation of the substrate at the active site. Although designed as entropy traps, and despite evident restriction in the conformational freedom of their substrates, neither 1F7 nor AZ-28 lowers the entropy of activation for its reaction. Both have S‡ values that are 10–20 cal K−1 mol−1 less favorable than the corresponding uncatalyzed reactions [25, 37]. This contrasts with some natural chorismate mutases which do reduce the entropy barrier to reaction significantly [39]. Mechanistic interpretations of activation parameters are necessarily uncertain [40], but the unfavorable S‡ values are consistent with the need for substantial conformational change in the bound substrate as the reaction proceeds. However, other factors, including changes in solvation or conformation associated with the antibody, cannot be excluded. More generally, these two examples show how the chemical instructions implicit in hapten structure, including deficiencies with respect to transition-state mimicry, are accurately imprinted on an antibody binding site. Improved transition-state analogs should therefore yield much better catalysts. To obtain more efficient chorismate mu-tase antibodies, for example, haptens containing additional negative charges might be used to elicit the catalytically essential cation in the vicinity of the substrate’s ether oxygen. Similarly, haptens in which the aryl substituents are coplanar with the cy-clohexyl ring should increase the probability of identifying faster catalysts for the oxy-Cope rearrangement of Fig. 3. The sensitivity of the latter reaction to anionic substituent effects [41] could also be drawn on. Haptens containing an appropriately positioned ammonium group might induce an antibody residue capable of deproto-nating the substrate alcohol. Fine-tuning of the first-generation antibodies is also likely to yield substantially better catalysts. Identification of a second chorismate mutase antibody possessing significantly higher activity than 1F7 [26] supports the feasibility of such an undertaking. Plausible strategies for optimizing activity include site-directed mutagenesis or random mutagenesis coupled with in vivo selection. The 1F7 Fab fragment’s ability to replace the missing enzyme in a chorismate mutase-deficient yeast cell line [42] is a promising indication that it will be amenable to directed evolution in the laboratory [43]. Overall, there are many ways to bind any given transition-state analog, only some of which will be effective for catalysis. Indeed, a significant fraction of hapten binders in any given experiment is usually found to be inactive. Hapten affinity rather than catalytic activity drives maturation of the immune response, so mutations can arise that favor tighter hapten binding but are deleterious for catalysis,

9

10 Catalytic Antibody Technology

as seen for AZ-28. Broad screening of antibodies raised to each hapten is therefore necessary to guarantee a representative sampling of the immune response. In the present instance, antibodies other than 1F7 and AZ-28 might be obtained that ultimately prove to be better starting points for optimization. 5.1.2 Cycloadditions Loss of both translational and rotational degrees of freedom should make bimolecular reactions particularly sensitive to proximity effects [20]. Diels-Alder reactions between dienes and dienophiles have been used to test this notion. They typically have high activation entropies in the range −30 to −40 cal K−1 mol−1 [44], reflecting the low probability of bringing together two substrates in an orientation optimal for reaction. The transition state for these concerted cycloadditions is highly ordered and resembles the boat form of the cyclohexene product more closely than is does the starting materials. Antibodies raised against bicyclic compounds that mimic the transition-state geometry have displayed a range of useful catalytic effects, including control over reaction pathway and absolute stereochemistry [45–49]. The general approach to catalysis is exemplified by antibody 1E9, which promotes the [4+2] cycloaddition cycloadditionand N-ethylmaleimide (Fig. 6) with multiple turnovers [45]. The initially formed adduct 3 spontaneously eliminates sulfur dioxide to give N-ethyl tetrachlorophthalimide as the final product after oxidation in situ. The endo hexachloronorbornene derivative 4, an excellent mimic of the intermediate and its flanking transition states, served as the hapten (K d = 2 nM). Because the planar product is structurally so different from 4, it binds 105 -fold less tightly to the induced antibody, effectively minimizing product inhibition. In this case, catalytic efficiency can be estimated as an effective molarity (EM) [50]. EM is the ratio of the pseudo-first-order rate constant for the antibody reaction (kcat ) to the second-order rate constant for the uncatalyzed process (kuncat ). This ratio

Fig. 6 Diels-Alder condensation of tetrachlorothiophene dioxide (TCTD) and N-ethyl maleimide (NEM) yields a high-energy intermediate (3) which eliminates sulfur dioxide. The initially formed product is subsequently

oxidized in situ. The transition state for cycloaddition and chelotropic elimination of SO2 closely resemble the hexachloronorbornene derivative 4 used as a hapten to elicit antibody 1E9 [45].

5 Representative Catalytic Antibodies

gives the nominal concentration of one reactant needed to convert the spontaneous bimolecular reaction into a pseudo-first-order process with a rate equivalent to that achieved in the antibody ternary complex. It is usually interpreted as the entropic advantage of a unimolecular over a bimolecular process, with an upper limit of about 108 M for 1 M standard states [51]. For 1E9, the EM is ca. 103 M [52]. Although much lower than the theoretical limit, this value is significantly higher than EMs reported for other antibody Diels-Alderases, which rarely exceed 10 M [46–49], making 1E9 the most efficient such catalyst described to date. Recent structural work has shown that the 1E9 active site is exactly what one might expect of an antibody that functions as an entropy trap [52]. Extensive van der Waals contacts, π-stacking interactions, and a strategically-placed hydrogen bond to one of the succinimide carbonyl groups create a pocket that is highly complementary to the hapten (Fig. 7a). When complexed, the ligand (excluding the hexanoate linker) is 86% buried. Its fit to the protein is so snug that no interfacial cavities are detectable, even when a probe of 1.2 Å radius is used. Thus, the 1E9

Fig. 7 The active site of Diels-Alderase 1E9 provides a snug, complementary binding surface for its hexachloronorbornene hapten [52]. With the exception of a hydrogen bond between the deeply buried carbonyl group of the ligand and the side chain of AsnH35 (not shown), most of the contacts are

hydrophobic in nature. (b) Antibody 39A11 (see Fig. 8) binds its hapten much more loosely. Note that the “dienophilelike” N-aryl succinimide is significantly better packed than the bicyclo[2.2.2]octene moiety, which serves as a diene surrogate.

11

12 Catalytic Antibody Technology

binding pocket appears ideally suited to the task of preorganizing its diene and dienophile substrates in a reactive complex that closely approximates the transitionstate geometry. Nevertheless, a simple entropy trap mechanism does not appear to be operative. The temperature dependence of kcat and kuncat shows that catalysis by 1E9 is achieved entirely by reducing the enthalpy of activation; the solution and the antibody processes are equally unfavorable entropically, with S‡ values of −22 cal K−1 mol−1 [52]. Nor is the rate acceleration due to a simple medium effect associated with the apolar binding cavity, since the uncatalyzed reaction is 10 times slower in acetonitrile than in water. Catalysis by 1E9 can be explained by enthalpic stabilization of the transition state through an unusually close fit to the apolar binding surface of the antibody active site and a strong hydrogen bond between the side chain of AsnH35 and the maleimide carbonyl. Quantum mechanical calculations indicate that the numerous, energetically favorable van der Waals interactions provide the driving force for binding [52]. Although these interactions distinguish poorly between ground and transition state, they hold the substrates against a relatively unfavorable electrostatic field that becomes substantially more favorable as the transition state is approached because of the increased strength of the hydrogen bond to the dienophile carbonyl. Comparison of 1E9 with another Diels-Alderase, 39-A11 [47], dramatically illustrates the importance of close packing for high efficiency. Antibody 39-A11 was generated with the substituted bicyclo[2.2.2]octene derivative 5, which contains an ethano bridge locking the cyclohexene ring into the requisite boat conformation (Fig. 8). It catalyzes the Diels-Alder reaction between an electron-rich acyclic diene and an N-aryl maleimide to give a cyclohexene derivative, albeit with a relatively low effective molarity of 0.35 M.

Fig. 8 Antibody 39-A11 catalyzes a Diels-Alder reaction between an electron-rich acylic diene and an N-aryl maleimide. It was elicited with the bicyclo[2.2.2]octene hapten 5. The ethano bridge in 5 locks the cyclohexane ring into a boat conformation but has no counterpart in the substrate itself.

5 Representative Catalytic Antibodies

Considering the low dissociation constant reported for the antibody-hapten complex (K d = 10 nM) [47], the fit of the bicyclo[2.2.2]octene to 39-A11 is surprisingly loose (Fig. 7b) [53]. Only 66% of the hapten surface area is buried in the complex. Poor complementarity is indicated by the large cavity volume of 117 Å3 detected between ligand and antibody. The portion of the hapten corresponding to the reacting [4+2] system is particularly poorly packed, whereas peripheral substituents of 5, especially the aryl side chain, appear to be important recognition elements. Moreover, the hapten’s ethano bridge, which has no counterpart in the substrates or the transition state, carves out additional unwanted space within the pocket. Consequently, the bound substrates – particularly the diene, which must bind in the least complementary region of the pocket – are likely to retain considerable degrees of freedom. Low catalytic efficiency is therefore unsurprising. Consistent with this idea, introducing large aromatic groups at positions L91 and L96 to improve packing interactions with the kinetically favored endo transition state results in 5 to 10-fold higher kcat values [54]. Relatively non-polar as it is, the environment of the 39-A11 active site may further erode catalytic efficiency. Like 1E9, 39-A11 provides a hydrogen bond (also from AsnH35 ) to the dienophile carbonyl, but its reaction involves a strong donor diene and acceptor dienophile rather than two electron-deficient addends. The corresponding transition state should therefore be more polar and hence more sensitive to transfer from water than that of the 1E9-catalyzed reaction. Unfortunately, mutagenesis experiments to augment activity by providing additional hydrogen bonds to the dienophile have not been successful [54]. Structurally distinct haptens notwithstanding, 1E9 and 39-A11 are unexpectedly closely related in primary sequence and tertiary structure [52, 53, 55]. Both belong to a family of polyspecific antibodies that exhibit extensive cross-reactivity for hydrophobic ligands containing one or two polar groups. For example, 39-A11 and its germline precursor accommodate a range of structurally diverse compounds [53], while 1E9 and the related progesterone-binding antibody DB3 [56, 57] bind each other’s ligands with affinities only 25-fold to 50-fold lower than their own [55]. The side chain of AsnH35 and conserved hydrophobic interactions seem to be particularly important for achieving recognition. However, docking experiments [52] with 1E9 and DB3 suggest that non-cognate molecules bind randomly in the apolar cavity, whereas specific ligands adopt a single, well-defined binding mode similar to that seen in crystal structures of the corresponding antibody-hapten complexes. Specificity in these systems appears to be conferred by a small number of mutations to the shared scaffold. In the case of 39A11, for instance, substituting Val for Ser at position L91 in the germline sequence accounts for almost all the 40-fold increase in hapten affinity achieved during affinity maturation [53]. Similarly, a somatic mutation in the L3 CDR loop (SerL89 → Phe) and a rare mutation in the antibody framework region (TrpH47 → Leu) substantially alter the shape of the 1E9 combining pocket compared to that of 39-A11 or DB3 [52]. These changes are primarily responsible for its virtually perfect shape complementarity to the transition-state analog. This complementarity appears crucial for reactivity, since DB3 does not detectably accelerate the 1E9 reaction despite its affinity for hapten 4.

13

14 Catalytic Antibody Technology

Given the immune system’s enormous combinatorial diversity (more than 108 antibodies are available in the primary response [58]), it is surprising that similar antibodies were generated in separate immunization experiments with 4, 5 and progesterone. Nor, as will be shown below, is this finding unique. To what extent does utilization of a few restricted sets of antibody germline genes limit the catalytic potential of the immune system? Do these frequently selected scaffolds represent local minima from which it will be difficult to evolve more active catalysts? Poor shape complementarity between 39-A11 and the bicyclo[2.2.2]octene is a caseinpoint. However, site-directed mutagenesis experiments demonstrate that the mature antibody is not an evolutionary dead end. The same ValL91 Tyr change that improves catalysis 10-fold also increases hapten affinity 2.5 times [54], and it should be possible to find additional mutations that further tighten the structure. Because only ten antibodies were screened for activity, we cannot know whether analogous mutations were present in the antibody population induced in response to hapten 5. It is conceivable that they never emerged: a 10 nM dissociation constant for the 39-A11-hapten complex is more than adequate for the immune system’s purposes, so there may be little or no selection pressure to increase affinity beyond that point. As shown by 1E9, the combining pocket can be molded remarkably well under favorable circumstances to achieve nearly perfect shape complementarity with a ligand. Superior fit is not necessarily manifest in tighter binding, since 1E9 and 39-A11 have similar hapten affinities (K d = 2 nM versus 10 nM). Compound 4 is more highly optimized than 5 with respect to transition-state mimicry, however, and important binding interactions in 1E9 are concentrated where they are needed for catalysis, rather than loosely dispersed as in 39-A11. A high degree of complementarity at the site of reaction thus appears to pay off in this case in terms of a more efficient catalytic outcome. Even in the case of 1E9, though, much higher efficiency should be attainable. An analogous enzyme is unavailable for direct comparison (see [59] for evidence regarding possible Diels-Alderases in Nature), but 1E9’s chemical proficiency, defined as (kcat /K diene K dienophile )/kuncat = 1.4 × 107 , is far from what is expected of a fully evolved catalyst. Given an already excellent fit between protein and ligand, it is unlikely that mutation of residues lining the binding cavity will substantially improve complementarity. To optimize electrostatic interactions with the transition state and reduce any remaining degrees of freedom available to the bound substrates, residues distant from the active site will have to be modified. Because such mutations will be difficult to identify by inspection of the protein structure, combinatorial mutagenesis and an efficient screening assay [60] or selection protocol will be needed if 1E9 variants with enhanced properties are to be developed. 5.2 Strain

Substrate destabilization through strain has been proposed as another mechanism for achieving rate accelerations with enzymes [20, 61]. Binding energy can be used to strain molecules in various ways. Destabilization can involve geometric

5 Representative Catalytic Antibodies

distortion of the substrate, electrostatic repulsion between groups of like charge, or desolvation effects. If substrate destabilization is relieved at the transition state, significant reductions can result in the free energy of activation for reaction. Rate accelerations obtained in this way are potentially quite large, limited only by the amount of binding energy available to force the substrate into the destabilizing environment. As with entropic effects, it was predicted that strain mechanisms might be readily exploited for antibody catalysis. 5.2.1 Ferrochelatase Mimics Ferrochelatase is an example of an enzyme that is believed to exploit geometric distortion for catalysis. As the terminal enzyme in heme biosynthesis, it promotes complexation of Fe2+ by protoporphyrin IX [62, 63]. Early work suggested that fer-rochelatase functions by distorting the substrate porphyrin from its preferred planar conformation into a bent structure [62]. This distortion exposes the nitrogen of one of the pyrroles to solvent, thereby facilitating metal ion complexation. In fact, non-planar N-methylated porphyrins are known to chelate metal ions 3 to 5 orders of magnitude faster than their non-alkylated counterparts [62]. They are also potent inhibitors of ferrochelatase [64]. When used as haptens, they have yielded antibodies [65] that catalyze insertion of divalent metal ions into mesoporphyrins with kcat values approaching those of the enzyme (Fig. 9). Ferrochelatase antibody 7G12 has been characterized in some detail. It promotes incorporation of Zn2+ and Cu2+ into mesoporphyrin IX with kcat values of 0.022 s−1 and 0.0069 s−1 , respectively [65]. By way of comparison, the kcat value for recombinant Bacillus subtilis ferrochelatase is ca. 0.4 s−1 for metalation with Fe2+ , Zn2+ and Cu2+ [66]. K m values for the porphyrin are also comparable (50 µM for the antibody and 8 µM for the enzyme). However, whereas substrate metal ions bind tightly to the enzyme (20–170 µM), no evidence for saturation of the antibody has been observed up to concentrations up to 2.5 mM, suggesting that binding of metal ions to the antibody does not contribute to catalysis. Severe inhibition of the antibodycatalyzed reaction by the metalloporphyrin product is a further point of contrast. Mechanistic and structural studies have clarified how the antibody chelatase functions. The crystal structure of 7G12 complexed with its hapten has been solved [67]. The N-methylated porphyrin binds at the junction of the heavy and light chains

Fig. 9 N-methyl mesoporphyrin and the mesoporphyrin metalation reaction catalyzed by antibody 7G12 [65].

15

16 Catalytic Antibody Technology

Fig. 10 Stereoview of the 7G12 binding site [67], showing how bound porphyrin packs against the heavy chain (blue) and exposes one face to solvent. The side chains of heavy chain residues AspH96 and ArgH96 are visible under the surface behind the porphyrin. Interactions with the light chain

(pink) are largely restricted to contacts with the methyl and ethyl substituents of the A and B pyrrole rings of the porphyrin. Although a mixture of N-alkylated porphyrins was used, only the derivative with the A ring methylated appears to bind to the antibody.

(Fig. 10). One face of the ligand makes extensive contacts with VH , while the other is relatively exposed to solvent. Packing interactions from light chain Tyr residues may reinforce the distortion from planarity of pyrrole ring A, which bears the N-methyl group. Replacement of these residues with Ala caused large increases in K m and 10-fold to 40-fold decreases in kcat /K m [67], suggesting that they may play a similar role with the non-methylated substrate. In analogy with the enhanced reaction rates achieved by porphyrin alkylation, a catalytic mechanism involving binding and distortion of the porphyrin by the protein, followed by direct chelation of metal ions from solution, seems plausible. Although its precise role is still unclear, amino acid AspH96 is evidently required for this process [67]. Its carboxylate side chain is directed from the VH domain toward the center of the porphyrin ring with one oxygen roughly equidistant from the porphyrin’s four pyrrole nitrogens (Fig. 10). The other oxygen is fixed in place through a hydrogen bond to ArgH95 . It is conceivable that the carboxylate acts as a base which shuttles protons from the porphyrin during metal ion exchange. This residue is also probably responsible, at least in part, for product inhibition, since axial coordination to the metal ion will anchor the metaloporphyrin to the antibody. Direct experimental evidence for porphyrin distortion by 7G12 has been obtained with resonance Raman spectroscopy [68]. Spectral data show that the antibody

5 Representative Catalytic Antibodies

induces an alternating up-and-down tilting of the pyrrole rings very similar to the distortion produced by porphyrin alkylation. In contrast, yeast ferrochelatase apparently causes all four pyrrole rings to tilt in the same direction in a domed fashion. The enzymatic reaction is regulated allosterically by a metal-dependent protein conformational change. Since the antibody has no metal binding site, the distortion it induces must be brought about entirely by binding interactions between porphyrin and protein. An atomic-level explanation of this effect will require elucidation of the antibody-substrate complex structure. The broad lesson to be derived from these experiments is that substrate destabilization can be a very successful approach to antibody catalysis. Although 7G12 and ferrochelatase perturb the porphyrin structure in different ways, both kinds of distortion appear to be effective for metal ion chelation, yielding kcat values within a factor of 10 of each other. That said, destabilization mechanisms are expected to have little to no effect on kcat /K m [69], the steady-state parameter generally optimized through evolution. By this criterion, 7G12 is substantially more primitive than its natural counterpart. Because chelatases catalyze a bimolecular reaction, flux through the catalyst is limited by the least favorably processed substrate. For both enzyme and antibody, this is the metal ion. Hence, rate enhancements are given by [(kcat /K M2+ )/kuncat ]. Since K M2+ values for 7G12 are at least 103 times larger than those for natural ferrochelatases (assuming the metal ion binds at all), the antibody suffers an equivalent rate disadvantage under practical operating conditions, despite its favorable kcat . Similar considerations apply to the chemical proficiencies of the two catalysts [(kcat /K M2+ K porphyrin )/kuncat ]. The antibody’s transition-state affinity is at least four orders of magnitude lower than that of the enzyme. To improve 7G12, a suitable binding site for metal ions should be constructed, perhaps by extending light chain CDR loops that are near the exposed face of the bound porphyrin. The challenge will be to bring the metal ion into close proximity with the porphyrin without further increasing the active site’s affinity for product. As for the chorismate mutase antibody discussed above, an in vivo selection strategy is feasible. Ferrochelatase-deficient yeast auxotrophs have been reported [70]. Complementing this metabolic deficiency with the antibody catalyst would provide a means of identifying more efficient variants. 5.2.2 Other Systems Large rate accelerations are also expected when charged reactants are enthalpically destabilized relative to a charge delocalized transition state by desolvation [20]. Such a mechanism may contribute to the efficacy of enzymes that promote biologically relevant decarboxylations [71–74]. Antibody catalysis of model reactions, such as the solvent-sensitive decarboxylation of 3-carboxy-benzisoxazoles to salicylonitriles [75, 76] and the difficult decarboxylation of orotate to uracil [77], has been used to probe the role of medium effects in a tailored binding pocket. Significant rate enhancements have been achieved, and structural work characterizing the properties of successful active sites will be instructive. Given the well-established and often dramatic sensitivity to solvent change of many reaction types, medium effects are likely to be pervasive in antibody and enzymatic catalysis. Reactions that display

17

18 Catalytic Antibody Technology

large changes in charge localization, including decarboxylations, nucleophilic substitutions, and aldol condensations, should be especially amenable to such effects. 5.3 Electrostatic Catalysis 5.3.1 Acyl Transfer Reactions Enzymes frequently utilize hydrogen bonding and charged groups to stabilize polar transition states electrostatically [69]. When haptens containing positive and negative charges are used, electrostatic interactions can also be exploited for antibody catalysis. For example, anionic phosphonates and phosphonamidates, originally designed as potent inhibitors of hydrolytic enzymes [12, 78], have been useful in the production of antibodies that hydrolyze esters, carbonates and (more rarely) amides [79]. Such analogs resemble the transition state for hydrolysis in a number of ways, including tetrahedral geometry, negative charge, and increased bond lengths. How these features are reflected inthe induced binding pockets has been deduced through structural studies of six different esterase families [80–90]. Antibody 48G7 is typical of this class of catalyst [91]. It was generated against p-nitrophenyl phosphonate (6) and accelerates the hydrolysis of the corresponding activated ester 7a and carbonate 7b by factors of 1.6 × 104 and 4 × 104 , respectively (Fig. 11). Both the antibody and its germline precursor, with and without bound hapten, have been characterized to provide insight into the origins and evolution of its catalytic effects [85–87]. The mature antibody has a deep, well-defined combining site rich in aromatic residues (Fig. 12a). It binds the hapten in an extended conformation with the aryl group at the bottom of a hydrophobic cleft formed between CDRs L1 and H3. The negatively charged phosphonate moiety lies near the pocket entrance, where it forms multiple interactions with charged and neutral antibody residues. The pro-R phosphonyl oxygen hydrogen bonds with the TyrH33 phenolic hydroxyl group and forms a salt bridge with the ArgL96 guanidinium group, while the pro-S

Fig. 11 Hydrolysis of activated aryl esters 7a and carbonates 7b proceeds via an anionic and tetrahedral intermediate (in brackets). Hydrolytic antibody 48G7 was elicited with an aryl phosphonate derivative (6) that mimics this high-energy species and its flanking transition states [91].

5 Representative Catalytic Antibodies

oxygen hydrogen bonds to the HisH35 -imino group and the TyrH96 backbone NH. Comparison of the mature antibody with and without the transition-state analog shows no major conformational changes [86], suggesting a simple lock-and-key mechanism for hapten binding (see, however, [92]). Antibody 48G7 has a 30 000-fold higher affinity for the phosphonate hapten and 20-fold greater catalytic efficiency than its germline precursor. Nine somatic mutations, all lying outside the combining site, are responsible for these effects [85, 87, 93]. Although not in direct contact with bound ligand, the mutated residues appear to preorganize the pocket for binding and catalysis. Improved packing and secondary hydrogen-bonding interactions help limit side-chain and backbone flexibility inherent in the germline protein. In fact, germline 48G7 (Fig. 12b) is conformationally much more flexible than the mature antibody and undergoes significant reorganization upon hapten binding [87]. The hapten itself binds quite differently in the two complexes (Fig. 12a,b). The phosphonate moiety occupies essentially the same location in both, but it cannot form a hydrogen bond with TyrH33 in the germline complex because of an altered conformation of the CDR H1 loop. Further, the p-nitrophenyl group is rotated away from the position it adopts in the mature antibody to occupy a hydrophobic cleft constructed from framework residues. This second apolar pocket is present, but empty, in mature 48G7. The alternative binding mode is made possible by removal of an otherwise repulsive interaction with the nitro group of the hapten in the mature antibody by the somatic mutation SerL34 Gly. Concordant with hapten design, the crystallographic data suggest that 48G7 is a relatively simple catalyst which promotes ester hydrolysis by direct attack of hydroxide on the scissile carbonyl. Although the two hapten binding modes seen in the germline and mature 48G7 complexes create some ambiguity about the substrate’s preferred orientation, the side chains of TyrH33 , HisH35 and ArgL96 and the backbone amide of TyrH96 apparently stabilize the tetrahedral and anionic transition states electrostatically through hydrogen-bonding and ionic interactions (Fig. 12). An analogy between this anion binding site and the well-characterized “oxyanion hole” of serine proteases is apparent. Individually, however, the antibody residues are not very effective oxyanion stabilizers. Mutations at positions H33, H35 and L96 cause only 3 to 30-fold reductions in kcat [85]. Loss of the hydrogen bond between TyrH33 and the transition state probably contributes to the germline’s 20fold lower efficiency, as well. In contrast, replacement of a single asparagine in the oxyanion hole of the protease subtilisin results in 102 to 103 -fold losses in specific activity [94, 95]. High solvent accessibility and/or conformational mobility of the antibody residues may account for their limited efficacy. The 48G7 active-site structure and hapten-recognition properties are strikingly similar to those of other, independently-derived anti-phosphonate antibodies [96]. These include three catalysts (CNJ206 [80, 81], 17E8 [89] and 29G11 [90]) for the cleavage of p-nitrophenyl esters and three (D2.3, D2.4 and D2.5 [83, 84]) for the energetically more demanding hydrolysis of p-nitrobenzyl esters. Although many of these esterolytic antibodies derive from different germline sequences, they appear to have in common a deep hydrophobic pocket in the framework region into

19

20 Catalytic Antibody Technology

Fig. 12 Stereoview of the active site of mature 48G7 (a) and its germline precursor (b), showing the different orientation of the hapten in the two complexes [85–87]. The phosphonate moiety of 6 binds in roughly the same location in both, although the network of hydrogen-bonding interactions that

constitute the oxyanion hole is slightly different. The deeply bound aryl group binds in one of two available hydrophobic clefts at the bottom of the pocket depending on whether the residue at position L34 is Ser (germline) or Gly (mature).

which the leaving group’s aryl/benzyl ring binds (Fig. 13). All exploit multiple hydrogen bonds and salt bridges near the mouth of the cavity for recognition of the phosphonate moiety. Consequently, it is likely that all employ the same basic hydrolytic mechanism as 43G7 [96]. Given this, their comparative efficiency, which ranges over two orders of magnitude, must reflect differing abilities to stabilize the hydrolytic transition state relative to the bound ground state. Examination of the hapten complexes suggests that the rate enhancement (kcat /kuncat = 103 – 105 ) roughly parallels the number of hydrogen bonds to the phosphonate. Flexibility in the active site (as seen in the 48G7 germline antibody) also correlates with low efficiency [80, 81, 87]. In some cases, the antibodies may exploit binding energy to

5 Representative Catalytic Antibodies

Fig. 13 Overlay of three hydrolytic antibodies, 48G7 (green), 17E8 (blue), and CNJ206 (red), shows remarkable structural convergence in these independently-generated active sites [96]. Only haptencontacting side chains are illustrated; the purple balls indicate the position of the phosphorus of the respective bound hapten.

distort the substrate ester from its thermodynamically favorable Z-conformation, making it easier to reach the tetrahedral transition state during catalysis [82, 85]. For example, this factor may come into play in 48G7, depending on whether the substrate binds like the hapten in the mature or the germline complex (Fig. 12). The remarkable degree of structural convergence observed in antibodies selected for tight binding to aryl phosphonate transition-state analog finds parallels in a number of other systems. For example, the immune responses to phosphorylcholine [97], p-azophenylarsonates [98], and 2-phenyloxazolones [99] are dominated by specific combinations of heavy and light chain variable regions. Interestingly, when a phenyloxazolone-binding antibody was found with a unique VH domain, its structure showed conservation of important antigen binding residues [100]. These results point to the immune system’s utilizing a relatively limited number of mechanisms to recognize any given type of antigen. Apparently, strong selective pressure reduces the broad diversity initially present in the primary repertoire to a small set of “best” solutions that can be further optimized by somatic mutation. As discussed above for the Diels-Alderases, structural convergence is even evident in antibodies raised against unrelated haptens. Rather than possessing an infinite variety of differently configured active sites, the immune system appears to play with a limited deck. Variations within the general theme do arise. Another esterase that has been structurally characterized, antibody 6D6, catalyzes the hydrolysis of a chloramphenical monoester with relatively low efficiency (kcat /kuncat = 900) [101]. It shows specific

21

22 Catalytic Antibody Technology

differences from the other antibodies [82]. In particular, the hapten is bound more shallowly, although the stacked aromatic rings of the leaving group and the acyl side chain are the most deeply buried portions of the molecule. The tetrahedral phos-phonate is highly solvent exposed, forming only one hydrogen bond to the antibody, presumably explaining 6D6’s relatively low efficiency. At this point,itisnot clear howmuch activity can be obtained from antiphosphonate antibodies. Over 50 esterolytic antibodies have been showntohave properties roughly comparable to those discussed here: simple hydrolytic mechanisms, rate accelerations up to 105 over background, and chemical proficiencies of 107 − 108 M−1 [15, 16]. While notable, particularly considering concomitant high and predictable selectivity [3], such effects are still orders of magnitude lower than those achieved by analogous enzymes. The immunological approach appears to have hit a ceiling in catalytic efficiency. It seems likely that these activities reflect what can be achieved with an “oxyanion hole” mechanism alone. Recent attempts to augment activity by rational mutagenesis or affinity selection with phage-displayed antibody fragments have met with only limited success [102–105]. Negative results should not be overinterpreted, but theydo raise concerns that the aryl phosphonate binding pocket common to these catalysts is not an intermediate that can be further refined but an evolutionary dead end. Alternative antigen presentation strategies and in vitro selection methods may provide access to different subsets of the immune repertoire more amenable to optimization. For example, antibodies that bind tetrahedral anions not at the entrance to the combining site but deep within their active site, as seen for enzymes that promote hydrolyses, would be of interest. Of course, highly evolved hydrolytic enzymes are more than simple oxyanion holes. They exploit arrays of catalytic groups (and, often, metal cofactors) to catalyze energetically demanding reactions such as amide hydrolysis. Induction of several precisely aligned functional groups in a single immunization step is extremely improbable, however. Such arrays are unlikely to be present in the primary repertoire of the immune system, and, depending on the basic immunoglobulin scaffold, may not be accessible through somatic hypermutation. Occasionally, though, serendipitous mutations that open up new opportunities for catalysis can and do occur. Although phosphonate transition-state analogs specify a simple hydrolytic mechanism, some anti-phosphonate antibodies have been identified that exploit more complex mechanisms. For example, the lipase-like antibody 21H3, generated against a typical benzyl phosphonate ester [106], unexpectedly accelerates ester hydrolysis by a two-step mechanism involving transient acylation of an amino acid within the binding pocket [107]. Its efficiency at hydrolyzing esters is no greater than that of the esterolytic antibodies discussed above, but use of covalent catalysis makes possible stereoselective transesterification reactions that cannot be carried out in water without the antibody [107, 108]. Similarly, considerable biochemical evidence has been adduced for a two-step sequence involving an acyl-antibody intermediate in hydrolytic reactions catalyzed by antibody 43C9 [109]. The latter was generated against phosphonamidate 8 [110], rather than a phosphonate, and it is

5 Representative Catalytic Antibodies

Fig. 14 Phosphonamidate 8, which is a transition-state analog for amide hydrolysis, yielded antibody 43C9 [110]. This antibody cleaves structurally related p-nitroanilides 9 and esters (not shown).

unique in its ability to promote the hydrolysis of activated amides as well as esters (Fig. 14). A 2.5 × 105 -fold rate enhancement (chemical proficiency = 5 × 108 M−1 ) achieved in the cleavage of p-nitroanilide 9 makes 43C9 one of the most efficient hydrolytic antibodies known. The crystal structures of free 43C9 and its complex with p-nitrophenol were recently solved [88]. Although detailed understanding of interactions specific to ligand recognition and catalysis await characterization of the hapten complex, likely participants in the reaction are identifiable upon inspection of the binding pocket (Fig. 15). As with other hydrolytic antibodies, several residues are available for stabilizing the negative charge that develops in the transition state, many of which are shared with 48G7. These include HisH35 and ArgL96 , whose importance for catalysis is supported by the results of site-directed mutagenesis experiments [103]. Residue AsnH33 , like 48G7’s TyrH33 , may also play a role in transition-state stabilization. What distinguishes 43C9 from other esterolytic antibodies, though, is the presence of a second histidine at position L91 . HisL91 is directed into the pocket toward the region where substrate must bind (Fig. 15). Docking experiments have suggested

Fig. 15 Stereoview of the active site of amidase 43C9 with bound p-nitrophenol [88]. The imidazole side chain of HisL91 (green) points into the pocket toward the region normally occupied by the phosphonate in

other hydrolytic antibodies. AsnH33 , HisH35 and ArgL96 are potential oxyanion stabilizers. Two active-site water molecules are shown as red spheres.

23

24 Catalytic Antibody Technology

a plausible orientation of the substrate, placing its scissile carbonyl in an excellent position for nucleophilic attack by this residue’s imidazole side chain [88]. Detailed mechanistic inferences should be considered tentative in the light of likely perturbations to the binding pocket caused by packing interactions between the active sites of adjacent molecules in the crystal [88]. Nevertheless, mutagenesis of HisL91 to Gln decreases catalytic efficiency >50-fold with little effect on ligand binding, supporting its role as the nucleophile that is transiently acylated during catalysis [103]. Consistent with this possibility, the acyl intermediate detected by electrospray mass spectrometry at pH 5.9 is not observed with the HisL91 Gln variant [111]. Covalent catalysis by imidazole is well-established in non-enzymatic reactions of carboxylic acid derivatives [1]. Its efficiency is a consequence of imidazole’s high nucleophilicity and the relative instability of the acyl-imidazole intermediate. The presence of HisL91 can thus explain why 43C9, but not 48G7 or other esterolytic antibodies, cleaves an amide. Context is clearly relevant, since lipase 21H3 lacks amidase activity though it also exploits nucleophilic catalysis [106, 107]. The fact that 43C9’s mechanism is unspecified by its hapten’s design also underscores the importance of serendipity in these experiments. The rarity of such occurrences is reflected in many failed experiments to generate amidases with phosphonamidate haptens. Despite its relative mechanistic sophistication, 43C9 is still a primitive amidase when compared with a typical protease. Subtilisin, which also exploits nucleophilic catalysis, catalyzes the cleavage of succinyl-Ala-Ala-Pro-Phe-p-nitroanilide 104 times more efficiently than 43C9. Its rate acceleration is 3.9 × 109 over background and its chemical proficiency is 2.2 × 1013 M−1 [112]. Moreover, subtilisin can cleave unacti-vated amides, whereas 43C9 is restricted to substrates with leaving groups of pK a < ca. 12 [113]. The high efficiency of covalent catalysis in serine proteases like subtilisin derives from cooperative action of several functional groups in addition to the active site nucleophile. Acid-base chemistry, in particular, is used to activate the serine nucleophile, facilitate proton transfers, and stabilize the amide leaving group. Removal of any single component of the protease’s catalytic triad results in 104 to 106 -fold decreases in activity [112]. Unsurprisingly, 43C9 lacks an analogously complex catalytic machinery. Although it has proven difficult to install additional functional groups by mutagenesis [103], more productive changes may become obvious when the structure of the hapten complex is known. Lessons from the immunological evolution of 48G7 [53, 87] suggest that mutagenesis of residues outside the binding pocket will be required to fine-tune critical synergies between participants in catalysis. 5.4 Functional Groups

Functional groups – acids, bases, and even exogenous cofactors — can clearly extend the capabilities of catalytic antibodies. They will certainly be needed if reactions with large activation barriers are to be significantly accelerated. As we have seen in the case of amidase 43C9, unplanned but useful residues can appear by chance in the

5 Representative Catalytic Antibodies

immunoglobulin pocket during affinity maturation. To increase the probability of eliciting such functional groups where they are needed, several strategies have been devised. Charge complementarity between an antibody and its ligand is the easiest principle to exploit in generating functionalized binding pockets. Cationic haptens have been used, for instance, to elicit negatively charged carboxylates that can serve as bases or nucleophiles or as general acids when protonated. Elimination reactions, epoxide ring openings, cationic cyclizations, and hydrolyses of esters, ketals, and enol ethers have been successfully catalyzed by this approach [114]. In favorable cases, very large catalytic effects have been achieved. For example, antibodies raised against a protonated benzimidazolium derivative use an active-site Asp or Glu to deprotonate substituted benzisoxazoles with effective molarities of 40 000 M and rate accelerations in excess of 108 over the acetate-promoted background reaction [115]. Nevertheless, it is unlikely that a single hapten will ever elicit arrays of residues as sophisticated as those present in highly evolved enzymes. Heterologous immunization has been explored as a method of circumventing this limitation [116]. In this procedure, two molecules, each containing different functional groups, are serially used as haptens to elicit the immune response. Ideally, a subset of the resulting antibodies will possess multiple catalytic groups induced in response to both tem-plating molecules and have, as a result, enhanced activities. An important advantage of this strategy is that simplified haptens can be used, reducing the need for laborious synthesis. Although generality must still be established, results from initial studies indicate that antibody esterases generated by heterologous immunization are more efficient than those generated in response to the individual haptens [116, 117]. Mechanism-based enzyme inhibitors [17, 18] are potentially of even greater utility than haptens [19, 118–122]. Such molecules exploit a protein’s ability to initiate a cascade of events, ultimately leading to its own covalent modification. This irreversible chemical reaction thus provides a means of selecting immunoglobulins in vivo and/or in vitro on the basis of their activity. When the selected antibody is challenged with substrate rather than hapten, the same group(s) responsible for protein modification can be used to promote the desired chemical transformation. Covalent catalysis can thus be specified through hapten design. The potential of reactive immunization is perhaps best illustrated by the production of aldolase antibodies [120]. These catalysts not only utilize a complex chemical mechanism but are among the most efficient catalytic antibodies described to date.

5.4.1 Aldolases Aldol condensations are broadly useful carbon-carbon bond-forming reactions in organic synthesis. They pose special difficulties for biocatalysts, however, because they proceed via a series of consecutive transition states, each requiring acid-base catalysis. Class I aldolases solve these problems by using a reactive lysine to activate the ketone donor through Schiff base formation at the active site [123–125]. Deprotonation of the Schiff base yields an enamine that then adds stereoselectively

25

26 Catalytic Antibody Technology

Fig. 16 $-Diketone 10, used as a hapten to raise antibodies 38C2 and 33F12, traps a reactive, active-site amine to form a stable, chromophoric vinylogous amide [120]. These antibodies promote diverse aldol

condensations. In the example shown (bottom), the two products are formed in a ca. 1:1 ratio with the indicated diastereoselectivities.

to the acceptor aldehyde to form a new carbon-carbon bond. Subsequent hydrolysis of the Schiff base releases product and regenerates the active catalyst. β-Diketones inhibit class I adolases by forming a Schiff base with the active-site lysine and then rearranging to a more stable vinylogous amide. When β-diketone 10 was used as a hapten instead of a more conventional transition-state analog [120], two analogously modifiable antibodies (33F12 and 38C2) were obtained (Fig. 16). The reactive group on the antibodies is a lysine with an anomolously low pK a (5.5 for 33F12 and 6.0 for 38C2 [123]). The vinylogous amide it forms with the hapten (λmax 316 nm, 15 000 M−1 cm−1 ) can be irreversibly trapped by reduction with sodium cyanoborohydride. These same antibodies also mimic the activity of class I aldolases [120]. Their reactive lysine reacts with a wide range of ketones to form enamine adducts. These condense with diverse aldehydes to form aldol products. The structure of antibody aldolase 33F12 [123] shows the catalytic lysine (LysH93 ) buried at the bottom of a hydrophobic pocket (Fig. 17). It is not hydrogen bonded with other amino acids and there are no charged residues within 7Å. The absence of such interactions must be responsible for this group’s unusual reactivity: the hydrophobic microenvironment lowers the amine’s pK a by disfavoring the protonated state. In contrast, class I aldolases are believed to increase the acidity of their catalytically essential lysine through proximity to several positively charged residues [126]. Unfortunately, the unliganded antibody provides few clues about the interactions that stabilize the transition states for formation and breakdown of the carbinol amine, deprotonation to afford the enamine, and creation of the new carbon-carbon bond. Sequestered water molecules or the hydroxyl groups of Tyr or Ser residues

5 Representative Catalytic Antibodies

Fig. 17 The binding pocket of antibody 33F12 [123] is seen through a slice in the molecular surface calculated with a sphere of 1.4 Å radius [206]. Only the g-amino group of LysH93 contacts the molecular surface at the bottom of the antigen binding site.

that constitute the pocket walls may be involved. For instance, a simple rotation of the Lys side chain from its position in the unliganded antibody would bring its amino group near SerH100 , which lies on the opposite side of the pocket. Structures of enamine adducts will be important for clarifying these points. An unanticipated feature of the aldolase antibodies is their promiscuity. Over 100 different reactions, including aldehyde-aldehyde, ketone-aldehyde and ketoneketone condensations, are subject to catalysis [123]. Because the 33F12 pocket is relatively hydrophobic, polyhydroxylated aldehydes, such as glyceraldehyde, glucose, and ribose, which are good acceptors for natural aldolases, are not substrates for the antibodies. Aside from this restriction, a wide range of donors and acceptors is tolerated. Non-specific van der Waals interactions likely provide the driving force for sequestering the first substrate. Once bound, it encounters the reactive lysine and forms the nucleophilic enamine. Provided there are no steric clashes, similar interactions should allow the aldehyde acceptor to bind and undergo aldol addition. The binding site is 11 Å deep and quite capacious, much larger in fact than the β-diketone that induced it, accounting for this broad specificity. These properties have been rationalized as a consequence of the reactive immunization process itself [123]: capture of the antibody by a covalent chemical event early in the process of affinity maturation may obviate the need for further refinement of the binding pocket by somatic mutation. Although induced with an achiral hapten and despite their broad substrate specificity, these aldolase antibodies are surprisingly stereoselective (Fig. 16). When acetone is the donor substrate, addition preferentially occurs on the si-face of the aldehyde acceptor; with hydroxy acetone, attack occurs on the re-face. In most cases, enantiomeric excesses >95% are found [127]. Enantioselective Robinson annulations [128], resolution of tertiary aldols [129], and preparation of chiral intermediates for the total synthesis of brevicomins [130] and epothilones [5] illustrate the synthetic utility of these catalysts. Kinetic resolution of the epothilone precursor

27

28 Catalytic Antibody Technology

was carried out on a gram scale using 0.06 mol% of antibody 38C2 (corresponding to 0.5 g of IgG). The reaction proceeded with good yield (37%) and high enantiomeric excess (90%). More recently, antibody 38C2 has been used to activate prodrugs [6]. It catalyzes the selective removal of generic drug-masking groups via sequential retro-aldol and retro-Michael reactions. This advance could prove useful in the development of selective chemotherapeutic strategies. How do these remarkable antibody aldolases compare to natural aldolases? To address this question, it is easiest to consider representative retroaldol reactions. For such transformations, the antibodies achieve turnover numbers ranging from 0.0003 to 0.08 s−1 [127]. One of the best antibody substrates is 4-(4 isobutyramidophenyl)-4-hydroxy-2-butanone. In the presence of 38C2, it undergoes retroaldolization with kcat and kcat /K m values of 0.083 min−1 and 3.3 × 103 M−1 s−1 , respectively. The rate constant for the background reaction is 1.4 × 10−9 s−1 (aqueous buffer, pH 7 and 25 ◦ C). The antibody thus accelerates this reaction by a factor of 2 × 107 -fold; its chemical proficiency is 6 × 1010 M−1 . These are impressive effects for a catalytic antibody, and may be contrasted with activity accruing to FDP aldolase and KDPG aldolase, typical class I enzymes involved in sugar metabolism. FDP aldolase catalyzes the interconversion of fructose-1,6-diphosphate (FDP) to give dihydroxyacetone phosphate and glyceraldehyde-3-phosphate. Its steady state kinetic parameters in the cleavage direction are kcat = 48 s−1 and kcat /K m = 1.6 × 107 M−1 s−1 [131]. KDPG aldolase, which cleaves 2-keto-3-deoxy-6-phosphogluconate (KDPG) into pyruvate and glyceraldehyde-3-phosphate, is even more active, having kcat and kcat /K m values of 290 s−1 and 4.0 × 10 M−1 s−1 , respectively [132]. Although rate constants for the corresponding uncatalyzed reactions are unavailable, taking 1.4 × 10−9 s−1 as a conservative estimate [127] yields rate enhancements of ca. 1010 − 1011 and chemical proficiencies of ca. 1015 –1016 M−1 . By either criterion, the natural enzymes are several orders of magnitude more efficient than the antibody aldolases. Natural aldolases are more restrictive in their substrate requirements than the antibodies, though they could lose considerable activity and still be competitive. For example, removal of a single phosphate in FDP causes a 50-fold loss in rate with FDP aldolase [131]. The extensive interactions that confer such specificity are almost certainly coupled to the enzyme’s high efficiency. By further refining the antibody pocket tothe precise steric and electronic demandsofaparticular aldol reaction, while maintaining the active-site lysine’s high reactivity, enzyme levels of activity may be attainable. Narrowed scope may be the unavoidable cost of truly high efficiency, however. 6 Perspectives 6.1 General Lessons from Comparisons of Enzymes and Antibodies

Structural and mechanistic work reviewed here reveals many notable parallels between antibodies and their more highly evolved counterparts. Not only are the

6 Perspectives

sizes and shapes of their active sites comparable, but antibodies and enzymes utilize the same set of molecular interactions to bind their respective ligands and stabilize transition states. Although antibodies tend to be less extensively functionalized than enzymes, the basic mechanistic strategies they employ to lower kinetic barriers are strikingly similar. As more primitive catalysts, however, they provide an alternative vantage point for examining the relationship between binding energy and catalysis. In this regard, simplicity is a virtue. Rather than working backwards from a fully evolved enzyme, uncomplicated, tailored model systems can be constructed to illuminate specific mechanistic questions. As multiple mechanistic strategies are combined to augment efficiency, valuable insight into the evolution of catalytic function can be gained. Functional analysis of the antibody intermediates that arise during affinity maturation [38, 53, 85, 87] also sheds light on these issues. Currently, antibodies appear less successful than enzymes intheir ability to achieve the fine level of recognition required for optimal discrimination between transition states and ground states. Their modest efficiencies appear to be a direct consequence of the simple strategy used to generate them. While the process of natural selection optimizes enzymes on the basis of their catalytic activity, the immune system’s microevolutionary mechanisms select antibodies for increased affinity to an imperfect transition-state analog. It is unrealistic toexpect that proteins engineered to recognize such haptens will provide an ideal steric and electrostatic environment for chemical transformation. Even with a perfect transition-state analog, the chances of obtaining a fully evolved catalyst through immunization would be low. As noted above, there is generally insufficient selection pressure to attain the high binding energies that characterize complexes between true enzymes and their transition states. On both micro and macro levels, mechanistic improvements arise as a function of time, so differences in time scales for the evolution of enzymes and antibodies – millions of years versus weeks or months – also come in to play. Although Nature uses a wide variety of different protein scaffolds to build enzyme active sites [133], she does not seem to have adopted the immunoglobulin fold. It is therefore conceivable that antibody structure itself places intrinsic limitations on the kind of reactions amenable to catalysis and on attainable efficiencies. In general, though, structural studies show excellent shape and chemical complementarity between antibodies and their ligands. Depending on the hapten, deep pockets, clefts, grooves, and flatter, more undulating surfaces can be created [134, 135]. Because certain classes of haptens tend to be recognized in the same way [52, 53, 83, 96], structural diversity must be considerably more restricted than might have been expected given 108 variants available in the primary immune repertoire [136, 137], but whether these consensus sites significantly restrict the catalytic capabilities of antibodies is still unclear. Conformational flexibility is another potential concern. Protein conformational changes in enzymes provide a means of excluding water from the active site and enable the catalyst to adjust tochanges in substrate as the reaction coordinate is traversed [138, 139]. Antibodies are known to undergo a comparable range of ligandinduced conformational changes, including alterations in side chain rotamers, segmental movements of hypervariable loops and changes in the relative disposition

29

30 Catalytic Antibody Technology

of the VH and VL domains [134]. Without direct selection for activity, however, these dynamic effects will be difficult to exploit deliberately for catalysis. In fact, conformational flexibility in catalytic antibodies, when observed [80, 81, 87], usually results in lower rather than higher efficiency. 6.2 How efficient does catalysis need to be?

Enzymes represent an extraordinarily high standard against which to judge new catalysts that are rationally designed from simple principles. From the perspective of the chemist, one exciting aspect of catalytic antibody technology is its ability to deliver tailored catalysts for reactions which are difficult to carry out selectively using existing methods or for which natural enzymes do not exist. Must such systems attain enzyme-like efficiency to be useful? Because antibodies are biocompatible and have long serum half-lives, many in vivo applications would be conceivable if sufficient activity were available. In fact, existing catalytic antibodies have already achieved significant effects in biological systems. When expressed at high levels, they have been shown to be competent (if inefficient) catalysts in metabolism, replacing essential enzymes in amino acid [42] and pyrimidine biosynthesis [77] pathways. Therapeutically relevant concentrations of the aldolase antibodies discussed earlierhave beenusedtoactivate prodrugsand kill colon and prostate cell lines [6]. Similarly, an esterolytic antibody has been employed as a cocaine antagonist, protecting rats from cocaine-induced seizures and sudden death [140, 141]. Many chemical reactions cannot proceed in the absence of catalysts because competing pathways have lower energies. In several instances, antibody binding energy has been successfully utilized to alter the course of such reactions by selectively stabilizing the less favorable transition state. For example, antibody catalysts have been developed for normally disfavored syn-eliminations [142], exo rather than endo Diels-Alder cycloadditions [46, 48], and 6-endo-tet ring closures of epoxy alcohols [143, 144]. Antibodies have also been used to control the fate of high-energy intermediates, allowing them to partition along only one of several possible pathways, as in the case of conversion of an enol ether to a cyclic ketal in water [145]. Formation of a strained cyclopropane derivative in a cationic olefin cyclization is another such example [146]. Binding energies up to 5.5 kcal/mol are typically available for achieving such discrimination, and even more energy may be available in favorable cases [15]. Such selectivities could be of great utility in organic synthesis. Assuming an antibody is available for any given transformation, its turnover and cost will ultimately determine whether it is used in practice. Presently, the steady state parameters of typical catalysts necessitate high antibody concentrations (≥ 10 µM = 1.6 mg/ml) and long reaction times to achieve useful conversions [12]. Preparative applications of several antibodies show that gram-scale reactions are feasible, particularly if antibody selectivity is high and competing reactions are substantially slower than the desired transformation [5, 147, 148]. Costs associated with high-volume antibody production are certainly an issue, but some antibodies

6 Perspectives

are now produced on a large scale for diagnostic and medical applications. They are readily obtained in good yield through ascites production [149] or by fermentation in hollow fiber reactors [150, 151]. Their Fab and Fv fragments can often be produced efficiently in plants [152, 153] or microorganisms [154]. Technical advances in microbial fermentation can be expected to make antibody production even more economically favorable in the future. In short, current levels of activity may be adequate for some laboratory applications, but higher efficiencies would certainly be beneficial. A 103 -fold increase in turnover would mean that 103 -fold less catalyst is needed to achieve useful levels of performance. Enhanced proficiency will certainly be necessary if energetically more demanding reactions are to be tackled. The creation of antibody equivalents of site-specific proteases, glycosidases, and nucleases, for example, remains a significant yet unrealized goal. The use of antibodies to synthesize or modify structurally complex and biologically important macromolecules will depend on solving this basic problem. 6.3 Strategies for Optimizing Efficiency

If imperfect design and indirect selection for binding rather than function are the primary reasons for low catalytic efficiency, creation of substantially better antibody catalysts will be feasible. Conceptually, two approaches can be envisaged: (1) refining methods for producing first generation catalysts, and (2) developing new strategies to optimize existing active sites. Improved transition-state analogs and more effective screening of the immune response address the first point. Rational reengineering and directed evolution methods are relevant to the second. These strategies have already been discussed in the context of specific antibody catalysts but are summarized in more general terms below. 6.3.1 Better Haptens The rewards of good – and the penalties of deficient – design are evident in the properties of catalytic antibodies characterized to date. In general, however, it is not clear which structural features of a transition state are most important to mimic to elicit maximally effective catalysts. Statistically meaningful correlation of different hapten types and the properties of their complementary active sites are needed to optimize analogs for each type of reaction. Incorporating design features that maximize transition-state affinity while minimizing ground-state stabilization remains a major challenge. For example, aryl groups are constituents of many haptens. Binding energy directed toward them, while increasing hapten affinity, may be useless or even harmful for catalysis, since these elements are common to both ground and transition state. Inefficient utilization of intrinsic binding energy in this way may help to explain the modest activities seen in the oxy-Cope [38] and Diels-Alder reactions [53] discussed above. The finding that catalytic activity for a series of polyclonal esterases correlates inversely with the size and hydrophobicity of the haptenic aryl phosphates [155] also illustrates this

31

32 Catalytic Antibody Technology

problem. When binding energy is used to recognize parts of the substrate distant from the site of reaction, product inhibition becomes another concern. Strategies to facilitate product release must therefore be considered integral to hapten design. Antigen presentation is a further issue. Small molecules are not immunogenic and mustbe coupledto carrier proteins. Variation of the tether site maybe a useful means of focusing immunorecognition to a hapten’s catalytically relevant epitopes [141]. Linkage of chemical reactivity with the immune system’s selection and amplification processes may mitigate some limitations in design. For this reason, the reactive immunization strategy [19, 118–122] merits increased attention. Many additional examples will be needed to establish its true scope and limitations. Given their importance in natural enzymes[156], metal ions and exogenous organic cofactors should considerably extend the properties of antibody catalysts, as well. Although versatile hybrid catalysts that combine the intrinsic reactivity of the metal ion or coenzyme with the tailored binding specificity of the antibody can be readily envisaged, this strategy has been surprisingly underutilized. Metal ion binding sites have been engineered into antibody binding sites [157], creating a sensitive Zn(II) sensor in one case [158], but catalysis has not been realized. An alternative, seemingly promising strategy using metal chelates for peptide cleavage has received little attention since first reported [159]. Modest activities have been described for other miscellaneous cofactor-dependent reactions [158, 160–163], but more work is obviously needed. Such strategies will be very important for promoting reactions with high kinetic barriers and reactions that cannot be carried out with protein residues alone. 6.3.2 Screening Because unusual germline sequences or fortuitous mutations may be necessary for high activity, the best antibody catalysts are also potentially the rarest. For this reason, more extensive screening of the immune response may dramatically increase the probability of finding highly active clones. Usually, small panels of antibodies chosen for their ability to bind the transition-state analog are purified and tested individually for catalytic activity. This procedure is necessarily indirect and slow. Sensitive chemical [164], biological [42, 77, 91, 165] and immunological assays [60, 166, 167] can facilitate the screening of thousands of candidates directly for catalysis and thereby accelerate the preliminary evaluation process. One practical problem associated with broad screening is that some antigens yield many hapten binders, others relatively few. To increase the size and diversity of the antibody population available for testing, multiple fusions can be performed and several mouse strains utilized for immunization. Mice prone to autoimmunity have been shown to yield unusually large numbers of esterolytic antibodies and may prove more generally useful for expanding the repertoire of catalytic clones elicited by a single transition-state analog [168]. Significant progress has also been made in copying the combinatorial processes of the immune system in vitro. Libraries of antibody fragments containing more than 106 members can be constructed and produced in microorganisms or

6 Perspectives

displayed on phage particles [169–172]. These systems are attractive vehicles for exploring the catalytic potential of different subsets of the primary immunological repertoire. Binders can be selected from these libraries on the basis of hapten affinity and subsequently screened for catalytic activity [169]. Clever strategies for capturing active clones based on their activity should be even more effective [173– 177]. Alternatively, catalysts can be obtained directly by selection in vivo using yeast or bacterial auxotrophs [42, 77, 91]. In these approaches, iterative rounds of mutagenesis and reselection replace somatic mutation as a means of refining initial hits [170]. Domain swapping [178] and powerful DNA shuffling methods [179, 180] have been developed to speed up this process. 6.3.3 Engineering The upper limit on activity that can be achieved with antibodies is unknown and may be reaction dependent. It is therefore important to push several test cases as far as possible. Site-directed mutagenesis is an attractive tool for improving catalytic power, particularly given the availability of increasing numbers of high-resolution structures. In general, it will probably be easiest to reengineer the poorest catalysts, since changes that improve packing or provide missing but critical interactions may be relatively obvious. The mutational study leading to an order of magnitude increase in activity of the Diels-Alderase antibody 39-A11 is a case in point [54]. The fact that relatively few changes are needed to tailor the properties of germline structure during affinity maturation [53, 85, 93] is also encouraging. Pinpointing subtle changes needed to optimize more active clones is likely to be more difficult, however. The obstacles encountered in augmenting the activity of hydrolytic antibodies sound a cautionary note [103–105]. Ultimately, our understanding of will determine what can be achieved in this way. 6.3.4 Selection Enzymes have been brought to peak efficiency over millions of years by the process of natural selection. An analogous process in the laboratory, involving recursive cycles of mutagenesis and genetic selection for function, may provide the ultimate test of the capabilities of antibody catalysts. Evolution of antibodies can be accomplished perhaps most directly by complementation of auxotrophic yeast or bacterial strains. The chorismate mutase antibody 1F7 [42] and an orotate decarboxylase antibody [77] have been shown to confer a significant growth advantage under selective conditions to host cells lacking the corresponding enzymes. Though the experiments are technically difficult because of poor antibody expression in microorganisms, preliminary results with the chorismate mutase antibody have demonstrated the feasibility of selecting antibodies with novel properties [43]. Recent work showing that cytoplasmic production of antibody fragments is optimizable by selection [181] augers well for these efforts. Selection systems are available for many transformations, including the ferrochelatase and metabolic reactions already mentioned [42, 70, 77]. A generalized selection scheme for hydrolytic reactions has also been reported [91]; analogous assays could be developed to exploit the ability of a catalyst to synthesize or destroy nutrients, drugs, hormones, or toxins. In addition to

33

34 Catalytic Antibody Technology

providing clues about the perfectibility of catalytic antibodies, such experiments may yield fundamental insights into structure-activity relationships and the evolution of molecular function. 6.3.5 Other Scaffolds The immune system was originally tapped as a source of catalysts as a matter of convenience: it is unrivaled in its ability to fashion high affinity protein receptors – the antibodies – to virtually any natural or synthetic molecule, essentially on demand [58, 136, 137]. However, now that the combinatorial processes of the immune system can be mimicked in vitro and libraries of macromolecules can be generated relatively easily using the tools of molecular biology, there is no compelling need to restrict Jencks’s original strategy to a single protein fold or even to a single class of macromolecule. Indeed, in analogy to catalytic antibody experiments, catalytic RNAs and DNAs have been obtained from large libraries of nucleic acids by selection for binding to transition-state analogs. Catalysts for porphyrin metalation obtained in this way [182–184] have activity comparable to that of the ferrochelatase antibody 7G12 discussed above, but an RNA rotamase is 30 times less effective than its antibody counterpart [185]. The lower activity of the latter largely reflects the RNA’s lower affinity for the hapten used (K d = 7 µM, compared with 0.21 µM for the antibody). A similar explanation has been invoked for RNAs that bind the 1E9 hapten (compound 4, Fig. 6) but fail to catalyze the corresponding Diels-Alder reaction [186]. Nucleic acids are likely to be intrinsically more limited than proteins in their capacity for high affinity molecular recognition of structurally diverse ligands as well as for catalysis [187]. Direct selection for function rather than transition-state analog binding has proven to be a much more powerful approach for obtaining nucleic acids with novel catalytic properties (for a recent comprehensive review, see [188]). RNA and have been prepared in this way for a variety of reactions, including a Diels-Alder cycloaddition [189]. Although the resulting catalysts often have relatively modest efficiency, phosphoryl transfers, which are difficult to achieve with antibodies because of the dearth of stable analogs of the pentacoordinate transition state, have been particularly amenable to catalysis. One of the most impressive accomplishments in this regard is the selection of highly efficient ribozymes capable of a self-ligation reaction from large pools of random sequence [190]. The number of starting molecules in these experiments was huge (ca. 1015 ), dwarfing the diversity of the primary immune repertoire (ca. 108 molecules), allowing even extremely rare catalysts to be found. One of the ribozymes obtained by selection was subsequently reengineered to function as a true catalyst; it promotes an intermolecular ligation with multiple turnovers and a rate acceleration approaching 109 [191]. This impressive activity is higher than any seen for most antibody catalysts, and provides an impressive demonstration of the power of direct selection. In principle, in vitro selection of peptides and proteins from vast combinatorial libraries is now possible as well using ribosome display [192–194], mRNAprotein fusion methods [195, 196] and more established phage display formats [172].

References

Coupled with efficient ways of linking genotype with phenotype [174–176], these methods can be expected to facilitate the production of proteins with novel properties and functions. As such, they powerfully complement other efforts to harness the power of evolution to redesign the structures and activities of existing enzymes [197–199, 207, 208].

7 Conclusions

Catalytic antibody technology combines programmable design with the combinatorial diversity of the immune system. This fusion has allowed the field to progress in relatively short order from simple model reactions to complex multistep processes, but much remains to be learned. Early efforts focused largely on defining the scope and limitations of this technology. Now that the approach is well established, attention must be paid to strategies for optimizing catalytic efficiency and for promoting more demanding transformations. In many ways, these are far greater challenges than identifying first-generation catalysts with modestactivity. Continued mechanistic and structural analysis of these systems will inform such endeavors. In addition, learning how to create, manipulate and evolve large combinatorial libraries of proteins outside the immune system should help to automate the processes of catalyst discovery and optimization.

Acknowledgements

The author is indebted to Kinya Hotta for creating the graphics for this article and to the National Institutes of Health, the ETH Zurich, the Swiss National Science Foundation, and Novartis Pharma for generous support.

References 1 JENCKS, W. P., Catalysis in Chemistry and Enzymology, McGraw Hill, New York 1969 2 SCHULTZ, P. G., LERNER, R. A., Science 269 (1995), p. 1835–1842 3 HILVERT, D., In Topics in Stereochemistry, ed. S. E. DENMARK 22, John Wiley & Sons, Inc., New York 1999, p. 83– 135 4 SCHULTZ, P. G., LERNER, R. A., Acc. Chem. Res. 26 (1993), p. 391–395 5 SINHA, S. C., BARBAS III, C. F., LERNER, R. A., Proc. Natl. Acad. Sci. USA 95 (1998), p. 14603–14608

6 SHABAT, D., RADER, C., LIST, B., LERNER, R. A., BARBAS III, C. F., Proc. Natl. Acad. Sci. USA 96 (1999), p. 6925–6930 7 RADZICKA, A., WOLFENDEN, R., Science 267 (1995), p. 90–93 8 WOLFENDEN, R., LU, X. D., YOUNG, G., J. Am. Chem. Soc. 120 (1998), p. 6814–6815 9 PAULING, L., Am. Sci. 36 (1948), p. 51–58 10 KURZ, J. L., J. Am. Chem. Soc. 85 (1963), p. 987–991 11 WOLFENDEN, R., Nature 223 (1969), p. 704–705 12 MADER, M. M., BARTLETT, P. A., Chem. Rev. 97 (1997), p. 1281–1301

35

36 Catalytic Antibody Technology

13 RADZICKA, A., WOLFENDEN, R., Methods Enzymol. 249 (1995), p. 284–312 14 LOLIS, E., PETSKO, G. A., Annu. Rev. Biochem. 59 (1990), p. 597–630 15 STEWART, J. D., BENKOVIC, S. J., Nature 375 (1995), p. 388–391 16 THOMAS, N. R., Appl. Biochem. Biotechnol. 47: (1994), p. 345–372 17 ABELES, R. H., MAYCOCK, A. L., Acc. Chem. Res. 9 (1976), p. 313–319 18 WALSH, C., Tetrahedron 38 (1982), p. 871–909 19 WIRSCHING, P., ASHLEY, J. A., LO, C.-H. L., JANDA, K. D., LERNER, R. A., Science 270 (1995), p. 1775–1782 20 JENCKS, W. P., Adv. Enzymol. 43 (1975), p. 219–410 21 WESTHEIMER, F. H., Adv. Enzymol. 24 (1962), p. 441–482 22 BARTLETT, P. A., NAKAGAWA, Y., JOHNSON, C. R., REICH, S. H., LUIS, A. J. Org. Chem. 53 (1988), p. 3195–3210 23 GRAY, J. V., EREN, D., KNOWLES, J. R., Biochemistry 29 (1990), p. 8872–8878 24 HILVERT, D., NARED, K. D., J. Am. Chem. Soc. 110 (1988), p. 5593–5594 25 HILVERT, D., CARPENTER, S. H., NARED, K. D., AUDITOR, M.-T. M., Proc. Natl. Acad. Sci. USA 85 (1988), p. 4953–4955 26 JACKSON, D. Y., JACOBS, J. W., SUGASAWARA, R., REICH, S. H., BARTLETT, P. A., SCHULTZ, P. G., J. Am. Chem. Soc. 110 (1988), p. 4841–4842 27 JACKSON, D. Y., LIANG, M. N., BARTLETT, P. A., SCHULTZ, P. G., Angew. Chem. Int. Ed. Engl. 31 (1992), p. 182–183 28 LEE, A. Y., STEWART, J. D., CLARDY, J., GANEM, B., Chem. Biol. 2 (1995), p. 195–203 29 CAMPBELL, P. A., TARASOW, T. M., MASSEFSKI, W., WRIGHT, P. E., HILVERT, D., Proc. Natl. Acad. Sci. USA 90 (1993), p. 8663–8667 30 HAYNES, M. R., STURA, E. A., HILVERT, D., WILSON, I. A., Science 263 (1994), p. 646–652 31 ADDADI, L., JAFFE, E. K., KNOWLES, J. R., Biochemistry 22 (1983), p. 4494– 4501 32 GUSTIN, D. J., MATTEI, P., KAST, P., WIEST, O., LEE, L., CLELAND, W. W., HILVERT, D., J. Am. Chem. Soc. 121 (1999), p. 1756–1757

33 KAST, P., ASIF-ULLAH, M., JIANG, N., HILVERT, D., Proc. Natl. Acad. Sci. USA 93 (1996), p. 5043–5048 34 LIU, D. R., CLOAD, S. T., PASTOR, R. M., SCHULTZ, P. G., J. Am. Chem. Soc. 118 (1996), p. 1789–1790 35 WIEST, O., HOUK, K. N., J. Org. Chem. 59 (1994), p. 7582–7584 36 BRAISTED, A. C., SCHULTZ, P. G., J. Am. Chem. Soc. 116 (1994), p. 2211–2212 37 DRIGGERS, E. M., CHO, H. S., LIU, C. W., KATZKA, C. W., BRAISTED, A. C., ULRICH, H. D., WEMMER, D. E., SCHULTZ, P. G., J. Am. Chem. Soc. 120 (1998), p. 1945–1958 38 ULRICH, H. D., MUNDORFF, E., SANTARSIERO, B. D., DRIGGERS, E. M., STEVENS, R. C., SCHULTZ, P. G., Nature 389 (1997), p. 271–275 39 G¨ORISCH, J., Biochemistry 17 (1978), p. 3700–3705 40 KAST, P., ASIF-ULLAH, M., HILVERT, D., Tetrahedron Lett. 37 (1996), p. 2691–2694 41 STEIGERWALD, M. J., GODDARD, W. A., EVANS, D. A., J. Am. Chem. Soc. 101 (1979), p. 1994–1997 42 TANG, Y., HICKS, J. B., HILVERT, D., Proc. Natl. Acad. Sci. USA 88 (1991), p. 8784–8786 43 TANG, Y., Evolutionary studies with a catalytic antibody, Ph.D., The Scripps Research Institute 1996 44 SAUER, J., SUSTMAN, R., Angew. Chem. Int. Ed. Engl. 19 (1980), p. 779–807 45 HILVERT, D., HILL, K. W., NARED, K. D., AUDITOR, M.-T. M., J. Am. Chem. Soc. 111 (1989), p. 9261–9262 46 GOUVERNEUR, V. E., HOUK, K. N., PASCUAL-TERESA, B., BENO, B., JANDA, K. D., LERNER, R. A., Science 262 (1993), p. 204–208 47 BRAISTED, A. C., SCHULTZ, P. G., J. Am. Chem. Soc. 112 (1990), p. 7430–7431 48 YLI-KAUHALUOMA, J. T., ASHLEY, J. A., LO, C.-H., TUCKER, L., WOLFE, M. M., JANDA, K. D., J. Am. Chem. Soc. 117 (1995), p. 7041–7047 49 RESMINI, M., MEEKEL, A. A. P., PANDIT, U. K., Pure Appl. Chem. 68 (1996), p. 2025–2028 50 KIRBY, A. J., Adv. Phys. Org. Chem. 17 (1980), p. 183–278 51 PAGE, M. I., JENCKS, W. P., Proc. Natl. Acad. Sci. USA 68 (1971), p. 1678–1683

References 52 XU, J., DENG, Q., CHEN, J., HOUK, K. N., BARTEK, J. D. H., WILSON, I. A., Science (1999), in press 53 ROMESBERG, F. E., SPILLER, B., SCHULTZ, P. G., STEVENS, R. C., Science 279 (1998), p. 1929–1933 54 ROMESBERG, F. E., SCHULTZ, P. G., Bioorg. Med. Chem. Lett. 9 (1999), p. 1741–1744 55 HAYNES, M. R., LENZ, M., TAUSSIG, M. J., WILSON, I. A., HILVERT, D., Israel J. Chem. 36 (1996), p. 151–159 56 AREVALO, J. H., TAUSSIG, M. J., WILSON, I. A., Nature 365 (1993), p. 859–863 57 AREVALO, J. H., HASSIG, C. A., STURA, E. A., SIMS, M. J., TAUSSIG, M. J., WILSON, I. A., J. Mol. Biol. 241 (1994), p. 663–690 58 KABAT, E. A., Structural Concepts in Immunology and Immunochemistry. Holt, Rinehart, and Winston, Inc., New York 1976 59 LASCHAT, S., Angew. Chem. Int. Ed. Engl. 35 (1996), p. 289–291 60 MACBEATH, G., HILVERT, D., J. Am. Chem. Soc. 116 (1994), p. 6101–6106 61 HALDANE, J. B. S., Enzymes. Longmans, Green and Co., London 1930 62 LAVALEE, D. K., Mol. Struct. Energ. 9 (1988), p. 279–314 63 AL-KARADAGHI, S., HANSSON, M., STANISLAV, N., JOHNSON, B., HEDERSTEDT, L., Structure 5 (1997), p. 1501–1510 64 DAILEY, H. A., FLEMING, J. E., J. Biol. Chem. 258 (1983), p. 11453–11459 65 COCHRAN, A. G., SCHULTZ, P. G., Science 249 (1990), p. 781–783 66 HANSSON, M., HEDERSTEDT, L. Eur. J. Biochem. 220 (1994), p. 201–208 67 ROMESBERG, F. E., SANTARSIERO, B. D., SPILLER, B., YIN, J., BARNES, D., SCHULTZ, P. G., STEVENS, R. C., Biochemistry 37 (1998), p. 14404–14409 68 BLACKWOOD Jr., M. E., RUSH III, T. S., ROMESBERG, F., SCHULTZ, P. G., SPIRO, T. G., Biochemistry 37 (1998), p. 779–782 69 WARSHEL, A., J. Biol. Chem. 273 (1998), p. 27035–27038 70 GORA, M., GRZYBOWSKA, E., RYTKA, J., LABBE-BOIS, R., J. Biol. Chem. 271 (1996), p. 11810–11816 71 CROSBY, J., STONE, R., LIENHARD, G. E., J. Am. Chem. Soc. 92 (1970), p. 2891–2900 72 O’LEARY, M. H., PIAZZA, G. J., Biochemistry 20 (1981), p. 2743–2748

73 ALSTON, T. A., ABELES, R. H., Biochemistry 26 (1987), p. 4082–4085 74 SUN, S. X., DUGGLEBY, R. G., SCHOWEN, R. L., J. Am. Chem. Soc. 117 (1995), p. 7317–7322 75 LEWIS, C., KRA¨ MER, T., ROBINSON, S., HILVERT, D., Science 253 (1991), p. 1019–1022 76 TARASOW, T. M., LEWIS, C., HILVERT, D., J. Am. Chem. Soc. 116 (1994), p. 7959– 7963 77 SMILEY, J. A., BENKOVIC, S. J., Proc. Natl. Acad. Sci. USA 91 (1994), p. 8319–8323 78 BARTLETT, P. A., MARLOW, C. K., Biochemistry 22 (1983), p. 4618–4624 79 STEWART, J. D., LIOTTA, L. J., BENKOVIC, S. J., Acc. Chem. Res. 26 (1993), p. 396–404 80 CHARBONNIER, J.-B., CARPENTER, E., GIGANT, B., GOLINELLI-PIMPANEAU, B., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., Proc. Natl. Acad. Sci. USA 92 (1995), p. 11721–11725 81 GOLINELLI-PIMPANEAU, B., GIGANT, B., BIZEBARD, T., NAVAZA, J., SALUDJIAN, P., ZEMEL, R., TAWFIK, D. S., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., Structure 2 (1994), p. 175–183 82 KRISTENSEN, O., VASSYLYEV, D. G., TANAKA, F., MORIKAWA, K., FUJII, I., J. Mol. Biol. 281 (1998), p. 501–511 83 CHARBONNIER, J.-B., GOLINELLI-PIMPANEU, B., GIGANT, B., TAWFIK, D. S., CHAP, R., SCHINDLER, D. G., KIM, S.-H., GREEN, B. S., ESHHAR, Z., KNOSSOW, M., Science 275 (1997), p. 1140–1142 84 GIGANT, B., CHARBONNIER, J.-B., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., Proc. Natl. Acad. Sci. USA 94 (1997), p. 7857–7861 85 PATTEN, P. A., GRAY, N. S., YANG, P. L., MARKS, C. B., WEDEMAYER, G. J., BONIFACE, J. J., STEVENS, R. C., SCHULTZ, P. G., Science 271 (1996), p. 1086–1091 86 WEDEMAYER, G. J., WANG, L. H., PATTEN, P. A., SCHULTZ, P. G., STEVENS, R. C., J. Mol. Biol. 268 (1997), p. 390–400 87 WEDEMAYER, G. J., PATTEN, P. A., WANG, L. H., SCHULTZ, P. G., STEVENS, R. C., Science 276 (1997), p. 1665–1669 88 THAYER, M. M., OLENDER, E. H., ARVAI, A. S., KOIKE, C. K., CANESTRELLI, I. L., STEWART, J. D., BENKOVIC, S. J., GETZOFF, E. D., ROBERTS, V. A., J. Mol. Biol. 291 (1999), p. 329–345

37

38 Catalytic Antibody Technology

89 ZHOU, G. W., GUO, J., HUANG, W., FLETTERICK, R. J., SCANLAN, T. S., Science 265 (1994), p. 1059–1064 90 GUO, J., HUANG, W., ZHOU, G. W., FLETTERICK, R. J., SCANLAN, T. S., Proc. Natl. Acad. Sci. USA 92 (1995), p. 1694–1698 91 LESLEY, S. A., PATTEN, P. A., SCHULTZ, P. G., Proc. Natl. Acad. Sci. USA 90 (1993), p. 1160–1165 92 LINDNER, A. B., ESHHAR, Z., TAWFIK, D. S., J. Mol. Biol. 285 (1999), p. 421–430 93 TOMLINSON, I. M., WALTER, G., JONES, P. T., DEAR, P. H., SONNHAMMER, E. L., WINTER, G., J. Mol. Biol. 256 (1996), p. 813–817 94 WELLS, J. A., CUNNINGHAM, B. C., GRAYCAR, T. P., ESTELL, D. A., Philos. Trans. R. Soc. Lond. (A) 317 (1986), p. 415–423 95 BRYAN, P., PANTOLIANO, M. W., QUILL, S. G., HSIAO, H.-Y., POULOS, T., Proc. Natl. Acad. Sci. USA 83 (1986), p. 3743– 3745 96 MACBEATH, G., HILVERT, D., Chem. Biol. 3 (1996), p. 433–445 97 PADLAN, E. A., COHEN, G. H., DAVIES, D. R., Ann. Inst. Pasteur Immunol. C 136 (1985), p. 271–276 98 STRONG, R. K., PETSKO, G. A., SHARON, J., MARGOLIES, M. N., Biochemistry 30 (1991), p. 3749–3757 99 GRIFFITHS, G. M., BEREK, C., KAARTINEN, M., MILSTEIN, C., Nature 312 (1984), p. 271–275 100 ALZARI, P. M., SPINELLI, S., MARIUZZA, R. A., BOULOT, G., POLJAK, R. J., JARVIS, J. M., MILSTEIN, C., EMBO J. 9 (1990), p. 3807–3814 101 MIYASHITA, H., KARAKI, Y., KIKUCHI, M., FUJII, I., Proc. Natl. Acad. Sci. USA 90 (1993), p. 5337–5340 102 ROBERTS, V. A., STEWART, J., BENKOVIC, S. J., GETZOFF, E. D., J. Mol. Biol. 235 (1994), p. 1098–1116 103 STEWART, J. D., ROBERTS, V. A., THOMAS, N. R., GETZOFF, E. D., BENKOVIC, S. J., Biochemistry 33 (1994), p. 1994–2003 104 BACA, M., SCANLAN, T. S., STEPHENSON, R. C., WELLS, J. A., Proc. Natl. Acad. Sci. USA 94 (1997), p. 10063–10068 105 ARKIN, M. R., WELLS, J. A., J. Mol. Biol. 284 (1998), p. 1083–1094

106 JANDA, K. D., BENKOVIC, S. J., LERNER, R. A., Science 244 (1989), p. 437–440 107 BENKOVIC, S. J., ADAMS, J. A., BORDERS, C. L., JANDA, K. D., LERNER, R. A., Science 250 (1990), p. 1135–1139 108 FERNHOLZ, E., SCHLOEDER, D., LIU, K. K.-C., BRADSHAW, C. W., HUANG, H., JANDA, K. D., LERNER, R. A., WONG, C.-H., J. Org. Chem. 57 (1991), p. 4756–4761 109 STEWART, J. D., KREBS, J. F., SIUZDAK, G., BERDIS, A. J., SMITHRUD, D. B., BENKOVIC, S. J., Proc. Natl. Acad. Sci. USA 91 (1994), p. 7404–7409 110 JANDA, K. D., SCHLOEDER, D., BENKOVIC, S. J., LERNER, R. A., Science 241 (1988), p. 1188–1191 111 KREBS, J. F., SIUZDAK, G., DYSON, H. J., STEWART, J. D., BENKOVIC, S. J., Biochemistry 34 (1995), p. 720–723 112 CARTER, P., WELLS, J. A., Nature 332 (1988), p. 564–568 113 GIBBS, R. A., BENKOVIC, P. A., JANDA, K. D., LERNER, R. A., BENKOVIC, S. J., J. Am. Chem. Soc. 114 (1992), p. 3528–3534 114 LERNER, R. A., BENKOVIC, S. J., SCHULTZ, P. G., Science 252 (1991), p. 659–667 115 THORN, S. N., DANIELS, R. G., AUDITOR, M.-T. M., HILVERT, D., Nature 373 (1995), p. 228–230 116 SUGA, H., ERSOY, O., WILLIAMS, S. F., TSUMURAYA, T., MARGOLIES, M. N., SINSKEY, A. J., MASAMUNE, S., J. Am. Chem. Soc. 116 (1994), p. 6025–6026 117 TSUMURAYA, T., SUGA, H., MEGURO, S., TSUNAKAWA, A., MASAMUNE, S., J. Am. Chem. Soc. 117 (1995), p. 11390–11396 118 JANDA, K. D., LO, C.-H. L., LI, T., BARBAS, C. F., WIRSCHING, P., LERNER, R. A. Proc. Natl. Acad. Sci. USA 91 (1994), p. 2532–2536 119 JANDA, K. D., LO, L.-C., LO, C.-H. L., SIM, M.-M., WANG, R., WONG, C.-H., LERNER, R. A., Science 275 (1997), p. 945–948 120 WAGNER, J., LERNER, R. A., BARBAS, C. F., Science 270 (1995), p. 1797–1800 121 LO, C.-H. L., WENTWORTH Jr., P., JUNG, K. W., YOON, J., ASHLEY, J. A., JANDA, K. D., J. Am. Chem. Soc. 119 (1997), p. 10251–10252 122 GAO, Cs., LAVEY, B. J., LO, C.-H. L., DATTA, A., WENTWORTH Jr., P., JANDA, K. D., J. Am. Chem. Soc. 120 (1998), p. 2211–2217

References 123 BARBAS III, C. F., HEINE, A., ZHONG, G., HOFFMANN, T., GRAMATIKOVA, S., BJO¨ RNESTEDT, R., LIST, B., ANDERSON, J., STURA, E. A., WILSON, I. A., LERNER, R. A., Science 278 (1997), p. 2085–2092 124 HORECKER, B. L., TSOLAS, O., LAI, C.-Y., In The Enzymes, ed. P. D. BOYER, 7, Academic Press, New York 1975, p. 213–258 125 MARSH, J. J., LEBHERZ, H. G., Trends Biochem. Sci. 17 (1992), p. 110–113 126 HIGHBARGER, L. A., GERLT, J. A., KENYON, G. L., Biochemistry 35 (1996), p. 41–46 127 HOFFMANN, T., ZHONG, G., LIST, B., SHABAT, D., ANDERSON, J., GRAMATIKOVA, S., LERNER, R. A., BARBAS III, C. F., J. Am. Chem. Soc. 120 (1998), p. 2768–2779 128 ZHONG, G., HOFFMANN, T., LERNER, R. A., DANISHEFSKY, S., BARBAS, III C. F., J. Am. Chem. Soc. 119 (1997), p. 8131–8132 129 LIST, B., SHABAT, D., ZHONG, G., TURNER, J. M., LI, A., BUI, T., ANDERSON, J., LERNER, R. A., BARBAS III, C. F., J. Am. Chem. Soc. 121 (1999), p. 7283–7291 130 LIST, B., SHABAT, D., BARBAS III, C. F., LERNER, R. A., Chem. Eur. J. 4 (1998), p. 881–885 131 PENTOET E. E., KOCHMAN, M., RUTTER, W. J., Biochemistry 8 (1969), p. 4396–4402 132 HAMMERSTEDT, R. H., M¨OHLER, H., DECKER, K. A., ERSFELD, D. E., WOOD, W. A., Methods Enzymol. 42C (1975), p. 258–264 133 BRANDEN, C., TOOZE, J., Introduction to Protein Structure, Garland Publishing, Inc., New York 1991 134 WILSON, I. A., STANFIELD, R. L., Curr. Opin. Struct. Biol. 3 (1993), p. 113–318 135 DAVIES, D. R., PADLAN, E. A., SHERIFF, S., Annu. Rev. Biochem. 59 (1990), p. 439–473 136 ALT, F. W., BLACKWELL, T. K., YANCOPOULOS, G. D., Science 238 (1987), p. 1079–1087 137 RAJEWSKY, K., F¨ORESTER, I., CUMANG, A., Science 238 (1987), p. 1088–1094 138 TSOU, C. L., Ann. N. Y. Acad. Sci. 864 (1998), p. 1–8 139 POST, C. B., RAY, W. J., Jr., Biochemistry 34 (1995), p. 15881–15885 140 METS, B., WINGER, G., CABRERA, C., SEO, S., JAMDAR, S., YANG, G., ZHAO, K., BRISCOE, R. J., ALMONTE, R., WOODS,

141

142

143 144

145

146 147

148 149

150 151

152

153 154

155 156

157

J. H., LANDRY, D. W., Proc. Natl. Acad. Sci. USA 95 (1998), p. 10176– 10181 YANG, G., CHUN, J., ARAKAWA-URAMOTO, H., WANG, X., GAWINOWICZ, M. A., ZHAO, K., LANDRY, D. W., J. Am. Chem. Soc. 118 (1996), p. 5881–5890 CRAVATT, B. F., ASHLEY, J. A., JANDA, K. D., BOGER, D. L., LERNER, R. A., J. Am. Chem. Soc. 116 (1994), p. 6013–6014 JANDA, K. D., SHEVLIN, C. G., LERNER, R. A., Science 259 (1993), p. 490–493 JANDA, K. D., SHEVLIN, C. G., LERNER, R. A., J. Am. Chem. Soc. 117 (1995), p. 2659–2660 SHABAT, D., ITZHAKY, H., REYMOND, J.-L., KEINAN, E., Nature 374 (1995), p. 143– 146 LI, T., JANDA, K. D., LERNER, R. A., Nature 379 (1996), p. 326–327 REYMOND, J.-L., REBER, J.-L., LERNER, R. A., Angew. Chem. Int. Ed. Engl. 33 (1994), p. 475–477 SINHA, S. C., KEINAN, E., J. Am. Chem. Soc. 117 (1995), p. 3653–3654 HARLOW, E., LANE, D., Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory 1988 LOWRY, D., MURPHY, S., GOFFE, R. A., J. Biotechnol. 36 (1994), p. 35–38 JACKSON, L. R., TRUDEL, L. J., FOX, J. G., LIPMAN, N. S., J. Immunol. Methods 189 (1996), p. 217–231 MA, J. K. C., HIATT, A., HEIN, M., VINE, N. D., WANG, F., STABILA, P., Van DOLLEWEERD, C., MOSTOV, K., LEHNER, T., Science 268 (1995), p. 716–719 HIATT, A., MA, J. K. C., FEBS Lett. 307 (1992), p. 71–75 CARTER, P., KELLEY, R. F., RODRIGUES, M. L., SNEDECOR, B., COVARRUBIAS, M., VELLIGAN, M. D., WONG, W. L. T., ROWLAND, A. M., KOTTS, C. E., CARVER, M. E., YANG, M., BOURELL, J. H., SHEPHARD, H. M., HENNER, D., BioTechnology 10 (1992), p. 163–167 WALLACE, M. B., IVERSON, B. L., J. Am. Chem. Soc. 118 (1996), p. 251–252 WALSH, C., Enzymatic Reaction Mechanisms, W.H. Freeman and Company, New York 1979 IVERSON, B. L., IVERSON, S. A., ROBERTS, V. A., GETZHOFF, E. D., TAINER, J. A.,

39

40 Catalytic Antibody Technology

158

159 160

161 162

163

164

165

166

167

168

169 170

171

172 173

174

BENKOVIC, S. J., LERNER, R. A., Science 249 (1990), p. 659–662 STEWART, J. D., ROBERTS, V. A., CROWDER, M. W., GETZOFF, E. D., BENKOVIC, S. J., J. Am. Chem. Soc. 116 (1994), p. 415–416 IVERSON, B. L., LERNER, R. A., Science 243 (1989), p. 1184–1188 SHOKAT, K. M., LEUMANN, C. J., SUGASAWARA, R., SCHULTZ, P. G., Angew. Chem. Int. Ed. Engl. 27 (1988), p. 1172–1174 NIMRI, S., KEINAN, E., J. Am. Chem. Soc. 121 (1999), in press COCHRAN, A. G., SCHULTZ, P. G., J. Am. Chem. Soc. 112 (1990), p. 9414–9415 COCHRAN, A. G., PHAM, T., SUGASAWARA, R., SCHULTZ, P. G., J. Am. Chem. Soc. 113 (1991), p. 6670–6672 EISENTHAL, R., DANSON, M. J., Enzyme Assays A Practical Approach. IRL Press, Oxford 1993 FENNIRI, H., JANDA, K. D., LERNER, R. A., Proc. Natl. Acad. Sci. USA 92 (1995), p. 2278–2282 LEE, I., STEWART, J. D., ZHONG, W., BENKOVIC, S. J., Anal. Biochem. 230 (1995), p. 62–67 TAWFIK, D. S., GREEN, B. S., CHAP, R., SELA, M., ESHHAR, Z., Proc. Natl. Acad. Sci. USA 90 (1993), p. 373–377 TAWFIK, D. S., CHAP, R., GREEN, B. S., SELA, M., ESHHAR, Z., Proc. Natl. Acad. Sci. USA 92 (1995), p. 2145–2149 CHISWELL, D. J., MCCAFFERTY, J., Trends Biotechnol. 10 (1992), p. 80–84 MARKS, J. D., HOOGENBOOM, H. R., GRIFFITHS, A. D., WINTER, G., J. Biol. Chem. 267 (1992), p. 16007–16010 HUSE, W. D., SASTRY, L., IVERSON, S. A., KANG, A. S., ALTING-MEES, M., BURTON, D. R., BENKOVIC, S. J., LERNER, R. A., Science 246 (1989), p. 1275–1281 SMITH, G. P., PETRENKO, V. A., Chem. Rev. 97 (1997), p. 391–410 JESTIN, J. L., KRISTENSEN, P., WINTER, G., Angew. Chem. Int. Ed. Engl. 38 (1999), p. 1124–1127 GAO, C., LIN, C. H., LO, C.-H. L., MAO, S., WIRSCHING, P., LERNER, R. A., JANDA, K. D., Proc. Natl. Acad. Sci. USA 94 (1997), p. 11777–11782

175 PEDERSEN, H., HOLDER, S., SUTHERLIN, D. P., SCHWITTER, U., KING, D. S., SCHULTZ, P. G., Proc. Natl. Acad. Sci. USA 95 (1998), p. 10523–10528 176 TAWFIK, D. S., GRIFFITHS, A. D., Nat. Biotechnol. 16 (1998), p. 652–656 177 SOUMILLION, P., JESPERS, L., BOUCHET, M., MARCHAND-BRYNAERT, J., WINTER, G., FASTREZ, J., J. Mol. Biol. 237 (1994), p. 415–422 178 MILLER, G. P., POSNER, B. A., BENKOVIC, S. J., Bioorg. Med. Chem. 5 (1997), p. 581–590 179 STEMMER, W. P. C., Nature 370 (1994), p. 389–391 180 CRAMERI, A., CWIRLA, S., STEMMER, W. P. C., Nat. Med. 2 (1996), p. 100– 102 181 MARTINEAU, P., JONES, P., WINTER, G., J. Mol. Biol. 280 (1998), p. 117–127 182 CONN, M. M., PRUDENT, J. R., SCHULTZ, P. G., J. Am. Chem. Soc. 118 (1996), p. 7012–7013 183 LI, Y. F., SEN, D., Nat. Struct. Biol. 3 (1996), p. 743–747 184 LI, Y. F., SEN, D., Biochemistry 36 (1997), p. 5589–5599 185 PRUDENT, J. R., UNO, T., SCHULTZ, P. G., Science 264 (1994), p. 1924–1927. 186 MORRIS, K. N., TARASOW, T. M., JULIN, C. M., SIMONS, S., HILVERT, D., GOLD, L., Proc. Natl. Acad. Sci. USA 91 (1994), p. 13028–13032 187 NARLIKAR, G. J., HERSCHLAG, D., Annu. Rev. Biochem. 66 (1997), p. 19–59 188 WILSON, D. S., SZOSTAK, J. W., Annu. Rev. Biochem. 68 (1999), p. 611–647 189 TARASOW, T. M., TARASOW, S. L., EATON, B. E., Nature 389 (1997), p. 54–57 190 BARTEL, D. P., SZOSTAK, J. W., Science 261 (1993), p. 1411–1418 191 EKLAND, E. H., SZOSTAK, J. W., BARTEL, D. P., Science 269 (1995), p. 364–370 192 HANES, J., PLU¨ CKTHUN, A., Proc. Natl. Acad. Sci. USA 94 (1997), p. 4937– 4942 193 HANES, J., JERMUTUS, L., WEBER-BORNHAUSER, S., BOSSHARD, H. R., PLU¨ CKTHUN, A., Proc. Natl. Acad. Sci. USA 95 (1998), p. 14130– 14135 194 HE, M. Y., TAUSSIG, M. J., Nucleic Acids Res. 25 (1997), p. 5132–5134

References 195 ROBERTS, R. W., SZOSTAK, J. W., Proc. Natl. Acad. Sci. USA 94 (1997), p. 12297–12302 196 NEMOTO, N., MIYAMOTO-SATO, E., HUSIMI, Y., YANAGAWA, H., FEBS Lett. 414 (1997), p. 405–408 197 MACBEATH, G., KAST, P., HILVERT, D., Science 279 (1998), p. 1958–1961 198 MINSHULL, J., STEMMER, W. P. C., Curr. Opin. Chem. Biol. 3 (1999), p. 284–290 199 ARNOLD, F. H., Acc. Chem. Res. 31 (1998), p. 125–131 200 SHOKAT, K. M., LEUMANN, C. J., SUGASAWARA, R., SCHULTZ, P. G., Nature 338 (1989), p. 269–271 201 TAWFIK, D. S., LINDNER, A. B., CHAP, R., ESHHAR, Z., GREEN, B. S., Eur. J. Biochem. 244 (1997), p. 619–626

202 ZEMEL, R., SCHINDLER, D. G., TAWFIK, D. S., ESHHAR, Z., GREEN, B. S., Mol. Immunol. 31 (1994), p. 127–137 203 LEE, A. Y., KARPLUS, A. P., GANEM, B., CLARDY, J., J. Am. Chem. Soc. 117 (1995), p. 3627–3628 204 ESNOUF, R. M., J. Mol. Graph. Model. 15 (1997), p. 132–134, 112–113 205 MERRITT, E. A., MURPHY, M. E. P., Acta Crystallogr. D 50 (1994), p. 869–873 206 NICHOLLS, A., SHARP, K. A., HONIG, B., Proteins Struct. Funct. Genet. 11 (1991), p. 281–296 207 JOO, H., LIN, Z. L., ARNOLD, F. H., Nature 399 (1999), p. 670–673 208 TAYLOR, S. V., KAST, P., HILVERT, D., Angew. Chem. Int. Ed. 40 (2001), 3310–3335

41

1

Structure and Function of Catalytic Antibodies Nicholas A. Larsen Harvard Medical School, Boston, USA

Ian A. Wilson The Scripps Research Institute, La Jolla, USA

Originally published in: Catalytic Antibodies. Edited by Ehud Keinan. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30688-6

1 Introduction

An explosion of antibody structures has emerged in the last 15 years [1, 2]. A subset of these antibodies is catalytic, and the latest version of the Protein Data Bank (PDB) (03/2003) contains 56 such structures representing 27 different antibodies (Table 1). Catalytic antibodies are elicited purposely by immunization with a transition state analog. In principle, antibodies that bind such an analog may also stabilize the true transition state of a chemical reaction relative to the ground state, thereby effecting catalysis [3–7]. Remarkably, the immunoglobulin fold possesses the inherent plasticity to harness the usual combination of electrostatic, van der Waals, hydrophobic, hydrogen bond, and cation-π interactions to generate tailor-made catalysts, which, in rare instances, even rival their natural enzyme counterparts. This general plasticity has been exploited to create model systems that explore a number of common and controversial themes in classical enzymology. Thus, substrate strain, approximation, control of the microenvironment, general acid-base chemistry, induced fit, and covalent catalysis have all been systematically examined in the context of the antibody binding pocket. The main objective of structural analysis is to develop hypotheses about key interactions in the binding pocket for binding substrate, transition state (TS), and product, and to propose plausible mechanisms for catalysis. For some antibodies, these hypotheses have been examined rigorously by biochemistry and mutagenesis (Table 1). Mutagenesis studies allow further quantification of the key interactions and provide an avenue for exploring the routes through the energy landscape that confer catalytic potential to the germline antibody precursor. Structural studies of

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

∗∗)

for diene and dienophile, respectively, equivalent to the so-called effective molarity. ∗∗∗) for substrate and cofactor (periodate), respectively.

∗)

0.28 0.30 0.34 0.08 0.64 0.46 0.05 0.39 0.05 0.004 0.22 3, 10*) 2, 30*) 1, .7* 0.17 0.26 0.27 .04, .25 ***) 0.4 0.36 0.32 0.23 0.21 0.15 0.074 0.051 1.3

3.6 1.0 0.07 0.4 60 220 1500 5.5 0.12 0.11 2.2 0.001 13 40 0.24 1.5 1.4 490 19.2 1.7 0.02 0.02 0.003 0.41 0.02 0.07 0.0016

D2.3 D2.4 D2.5 CNJ206 29G11 17E8 43C9 48G7 6D9 7C8 15A10 13G5 1E9 39-A11 9D9 10F11 33F12 28B4 21D8 5C8 19A4 4C6 1D4 7G12 AZ-28 1F7 4B2

Ester hydrol. pH = 8.3 Ester hydrol. pH = 8.3 Ester hydrol. pH = 8.3 Ester hydrol. pH = 8.0 Ester hydrol. pH = 9.5 Ester hydrol. pH = 9.5 Ester hydrol. pH = 9.3 Ester hydrol. pH = 8.2 Ester hydrol. pH = 8.0 Ester hydrol. pH = 8.0 Ester hydrol. pH = 8.0 Diels-Alder Diels-Alder Diels-Alder Retro-Diels-Alder Retro-Diels-Alder Retro-aldol condensation Sulfide oxidation Decarboxylation Cyclization Cationic cyclization Cationic cyclization Syn-elimination Metal chelatase Oxy-Cope Chorismate mutase Allylic isomerization

kcat (min-1 ) KM(mM)

Antibody Reaction

Table 1 Summary of catalytic antibody structures

2.00 – 2.40 2.15 1.90–2.80 1.61 – 2.10 2.00 – 2.50 2.70 1.30–2.45 1.80–1.85 1.60–2.60 2.00 – 2.80 3.00 1.87

1.85 – 2.10 3.10 2.20 3.20 2.30 2.50 2.20–2.30 1.95–2.70 1.80 2.20 2.35 1.95 1.90 2.10–2.40

x9 1YEC x1 1YED x1 1YEE x1 1KNO x1 1A0Q x1 1EAP x2 43CA x4 1GAF x2 1HYX x1 1CT8 x1 1NJ9 x1 1A3L x1 1C1E x2 1A4K x1 x4 1LO0 x1 1AXT x4 1KEL x2 1C5C x3 25C8 x1 1CF8 x2 1ND0 x2 1JGV x5 3FCT x4 1AXS x1 1FIG x1 1F3D

Yes – – – – Yes Yes Yes – – Yes – – Yes – – Yes Yes Yes – – – – Yes Yes – –

1.3 × 105 3.6 × 104 2.5 × 103 1.6 × 103 2.2 × 103 8.3 × 103 2.7 × 104 1.6 × 104 9.0 × 102 7.1 × 102 2.3 × 104 7 M**) 103M**) 0.4 M**) 4.0 × 102 2.5 × 103 1.7 × 107 9 M**) 2.0 × 104 ND ND ND ND – 5.3 × 103 2.5 × 102 1.5 × 103

1.1 × 105 3.3 × 104 1.3 × 103 0.7 × 102 2.4 × 104 9.0 × 102 6.8 × 104 8.7 × 104 9.0 × 102 1.2 × 10 2.2 × 104 – – – 4.1 × 102 2.5 × 103 – – 7.5 × 103 – – – – – 4.4 × 103 8.5 × 101 4.3 × 102

PDB ID

kcat kuncat -1 KM KDTSA −1 Mutant analysis Res. (Å)

[28] [28] [28] [26] [23] [23, 24] [27] [130] [20, 131] [131] [132, 133] [91] [94] [92, 93] [88, 134] [88, 134] [115–117, 135] [66, 68, 69] [14, 62, 65, 136, 137] [138] [16, 109] [17, 107] [100, 101] [70, 111, 113] [80, 81] [74, 77] [71, 72]

Refs.

2 Structure and Function of Catalytic Antibodies

2 Electrostatic Complementarity

non-catalytic germline precursors further illustrate how the antibodies evolve their catalytic potential by affinity maturation. Such a thorough understanding of these interactions is necessary if improvements are to be made to the antibody catalyst by either mutation or redesign of the immunizing hapten. Importantly, comparison of antibody structures has revealed common structural motifs that are elicited by different haptens during the immune response. Thus, convergent and divergent evolution has been observed directly within the immune system. Most of the catalytic antibody structures found in the are complexes with their respective transition state analog, and provide detailed views of the interactions that contribute to catalysis. In nearly all cases, a high degree of electrostatic and shape complementarity is observed for the corresponding transition state analog (TSA), and, in rarer instances, putative reactive residues that initiate covalent catalysis have been proposed. Consequently, this chapter summarizes antibody-catalyzed reactions in the general context of these three broad headings and concludes with a section on future challenges in the abzyme field for structural biologists and chemists alike.

2 Electrostatic Complementarity

Many antibody-TSA complexes exhibit a high degree of electrostatic complementarity. Recurring binding motifs for anions (e.g., phosphate, arsenate, sulfate) have been identified, indicative of structural convergence from a variety of germline sequences [8–14]. In contrast, general motifs for binding cations (e.g., primary, secondary, tertiary amines) have not yet been identified. Eliciting an Asp/Glu carboxylate using hapten baiting strategies has met with mixed success. Frequently, cations are buried preferentially in hydrophobic pockets but surrounded by aromatic side chains, which are believed to form stabilizing cation-π interactions [15–17].

2.1 Anionic Binding Motifs and Ester Hydrolysis

Nearly half of the catalytic antibody structures found in the PDB correspond to various hydrolytic antibodies in complex with substrate analogs and products (Table 1). Thus, hydrolytic antibodies are by far the most thoroughly studied catalytic antibodies from both a structural and a biochemical vantage, and have already been reviewed extensively in the literature [18, 19]. These hydrolytic antibodies have been raised generally against either phosphonate or phosphonamidate transition state analogs (Scheme 1). The phosphonate moiety is tetrahedral and carries a negative charge, thereby mimicking the transition state for solvent-mediated hydrolysis that involves nucleophilic attack by a water molecule. With the exception of 6D9 [20] 7C8, and 15A10, these hydrolytic antibodies catalyze the hydrolysis of benzyl or

3

4 Structure and Function of Catalytic Antibodies

Scheme 1 Hydrolytic Antibodies 48G7, 17E8, 29G11, CNJ206, 43C9, D2.3, D2.4, D2.5

activated 4-nitrophenol or 4-nitrobenzyl esters (Scheme 1), which are particularly convenient model substrates for both synthetic manipulation and sensitivity during hybridoma screening. 2.2 Structural Motifs

The first structural evidence that antibodies recognize antigens by electrostatic complementarity originates from the pioneering study of mAb McPC603, which binds phosphocholine [8, 9]. Further data have indicated that electronically-related pheny-larsonate compounds elicit a conserved anionic binding motif from the immune repertoire [10–13]. Likewise, crystal structures of catalytic antibodies 48G7, 17E8, 29G11, CNJ206, and 43C9 reveal a recurring motif that coordinates the phosphonate moiety of the hapten (Fig. 1a, b) [18, 19]. In general, the constellation of side-chains derived from L96 (CDR3), H33, H35 (CDR1), and the backbone amide of H96 (CDR3) constitute an anionic binding site, arising from structural convergence within the antibody binding pocket (Fig. 1a, b) [18, 19]. For example, antibody 48G7 utilizes ArgL96, TyrH33, HisH35, and the backbone amide of H96 for phosphonate binding (Fig. 1a) [21, 22]. Antibodies 17E8 and 29G11 are related to each other (92% sequence identity [23]) and use the same residues for phosphonate binding [24, 25] as 48G7, except that a hydrogen bond from the LysH93-Nz amine substitutes for the observed TyrH33 hydroxyl in 48G7. Antibody CNJ206 also similarly uses hydrogen bonds from HisH35 and the backbone amides of H96 and H97 to bind the phosphonate [26]. Although a crystal structure of 43C9 in complex with the transition state analog is not available, it is proposed that HisL91, ArgL96, AsnH33, and HisH35 likewise contribute to the anionic binding site (Fig. 1b) [27]. 2.3 Mechanistic Considerations

Clearly, aryl phosphonate/arsonate/sulfonate-like haptens preferentially elicit electrostatic complementarity from the immune response and may explain the commonly observed binding motif observed in 48G7, 17E8, 29G11, CNJ206, 43C9 and other non-catalytic antibodies. For the hydrolytic antibodies, the significance of the

2 Electrostatic Complementarity Fig. 1. Top view of hydrolytic antibodies. The light chain is represented in pink and the heavy chain in blue; side chains are brown and the ligand is yellow. Hydrogen bonds are represented as dashed green lines. A The hapten complex of 48G7 illustrates the prototypic anionic binding motif composed, for the most part, of heavy chain residues. B 43C9 likely has a similar motif as 48G7. The only available structure is a complex with the product p-nitrophenol. HisL91 is proposed to form an acylated intermediate at the N*1 atom. However, the interpretation of the kinetic and structural data is not fully convincing. C D2.3 has a unique anionic binding motif located on the light chain side of the antibody. The Tyr is proposed to form the key interaction responsible for oxyanion stabilization. Catalytic antibodies D2.3–5, 6D9, and 7C8 differ from this general theme. In D2.3, phosphonate binding arises from hydrogen bonds to AsnL34, TrpH95, and TyrH100D (Fig. 1c) [28]. Thus, the anionic binding site is located on the opposite side of the binding pocket and is composed of different residues. Finally, the phosphonate binding motifs in antibody 6D9 and 7C8 are completely different from those seen in any of the other antibodies and consist of a single, solvent-exposed, hydrogen bond donated from a single HisL27D or TyrH95, respectively [20, 29, 30]. Not surprisingly, these antibodies exhibit lower catalytic activity than any other structurallydetermined hydrolytic antibodies (Table 1). Nevertheless, 6D9 has been recently evolved to have 20-fold higher activity with the addition of a second hydrogen bond to the phosphonate [31].

anionic binding site is underscored by the general agreement between transition state stabilization relative to the ground state, K M K DTS −1 , and the observed rate acceleration, kcat kuncat −1 [32] (Table 1). This agreement implies that hapten-binding interactions observed in the crystal structures correspond to the interactions found in the transition state, and that binding energy is converted into catalysis with near 100% efficiency. Thus, the simplest possible mechanism for ester hydrolysis in this family of antibodies is oxyanion stabilization within the anionic binding site. Oxyanion stabilization has a differential impact on catalysis for various natural esterase and protease enzymes. For example, biochemical studies in a bacterial cocaine esterase have demonstrated that the oxyanion hole is critical, contributing at least 5 kcal mol−1 to catalysis [33]. Oxyanion stabilization is estimated to contribute 2–3 kcal mol−1 to catalysis in papain [34] and cutinase [35], 3-4 kcal mol−1

5

6 Structure and Function of Catalytic Antibodies

in prolyloligopeptidase [36] kcal mol−1 in subtilisin [37–39], and 5–7 kcal mol−1 in acetylcholinesterase [40] and Zn2+ peptidase [41]. These values are within the range ascribed to the oxyanion contribution in the hydrolytic antibodies. For example, a contribution of 5–7 kcal mol−1 from oxyanion stabilization was estimated for the D2.3 hydrolytic antibody [42]. Here, the oxyanion was formally assigned to TyrH100D from pH rate profile and the inactivity of a Tyr → Phe mutant (Fig. 1c) [42]. The alternative candidate, AsnL34, was excluded as the Asn → Gly mutant retained full activity [42]. The activity of this antibody is significantly enhanced by spiking the assay buffer with potent α-nucleophiles, such as peroxide [42]. Thus, the catalytic potential of D2.3 is far from realized, and could be increased by a carefully positioned general acid/base that could more effectively activate water for nucleophilic attack. Even with the more potent nucleophile, the activity of this antibody is apparently limited by product inhibition [42] and conformational isomerization [43]. A largely overlooked ramification of the known anionic binding sites is the asymmetry of the coordinating residues (Fig. 1a–c). This asymmetry means that attack of the substrate carbonyl carbon and oxyanion stabilization has enantiofacial selectivity [44], since the phosphonate oxygens are non-equivalent (prochiral). Thus, the lack of 18 O incorporation in the substrates of hydrolytic antibodies should not be regarded as evidence for an acyl enzyme intermediate [45], but rather that the oxygen atoms are not racemized in the stereoselective binding pocket after hydroxide attack. This enantiofacial selectivity has been noted in other naturally-occurring hydrolytic enzymes [46, 47]. 2.4 Mechanistic Pitfalls

There is an obvious temptation to ascribe complex mechanismsto hydrolytic antibodies to elevate them to the same status as their natural enzyme counterparts. In the case of 17E8, for example, the proximity of a serine and histidine to the phosphonate in the crystal structure led to the proposal that the mechanism involved a catalytic dyad, analogous to the familiar catalytic triad [24]. This structure-based hypothesis was further bolstered by hydroxylamine partitioning data and pH rate profiles that suggested an acyl enzyme intermediate [48]. Initially, this interpretation was criticized because the refined serine rotamer was not within hydrogen-bonding distance to the histidine in the putative dyad [18, 49], but alternative rotamers of a serine have been noted in other serine hydrolases [47, 50, 51]. Eventually, the acyl enzyme mechanism was effectively discounted by a Ser → Ala mutant, which led to twofold improvement in kcat over the wild-type [52]. Likewise, the hydrolytic mechanism for 43C9 was proposed to involve an unprecedented nucleophilic attack by the Nδx atom of HisL91 on the substrate to form an acylated histidine intermediate [53]. In serine hydrolases, the nucleophilicity of histidine N2 atom is enhanced by a buried aspartate that H-bonds to Nδ1 stabilizing the developing positive charge on the imidazole. In the 43C9 pocket, HisL91 N2 H-bonds to TyrL36, but the nearest charged residue is ArgL96 (Fig. 1b). Hence, the available structural data [27] do not

2 Electrostatic Complementarity

yet provide sufficient evidence for such a mechanism. Therefore, the interpretation of the kinetic, mutagenesis, and structural data [45, 53–56] should be carefully reexamined or, at the very least, regarded with skepticism until a direct observation of an acylated histidine intermediate is demonstrated in a crystal structure. 2.5 Structural and Functional Considerations for Hapten Design

The initial use of a phosphonate analog as the transition state for solvent-mediated hydrolysis (Scheme 1) provided the initial breakthrough in the catalytic antibody field [6, 7]. However, the persistent and ubiquitous use of such analogs to elicit hydrolytic antibodies has now become somewhat of a red herring. Phosphonate compounds are not transition state analog inhibitors for serine hydrolases [57]. Thus, it would be impossible to elicit a serine hydrolase-like antibody using a phosphonate analog. Nevertheless, the elicited antibodies are inevitably compared to serine hydrolases [24], or to hydrolase mutants where the entire catalytic triad has been mutated to alanine [58]. Fluorophosphonate compounds (nerve gas reagents, for example) are potent mechanistic inhibitors of serine esterases because of covalent modification of serine with phosphate and concomitant expulsion of fluoride – a good leaving group. Phosphonate and phosphonamidate compounds do not form the same modification in serine esterases, since O− is a poor leaving-group. In contrast, phosphonate compounds are potent transition state analog inhibitors for Zn2+ proteases [59], which utilize a completely different mechanism from that of serine hydrolases and probably do not involve covalent acylation by the enzyme [60]. Rather, Zn2+ is believed to facilitate water-mediated attack of the ester/amide carbonyl carbon with arginine stabilization of the oxyanion [41, 60]. Unfortunately, no design feature in a phosphonate analog would be likely to elicit Zn2+ binding in an antibody combining site. Thus, the likelihood of eliciting hydrolytic antibodies with either serine hydrolase or Zn2+ peptidase-like features would seem unlikely without some innovative new hapten design. 2.6 Anionic Binding Motifs and Control of the Microenvironment in Decarboxylation

Catalytic antibody 21D8 catalyzes a classic decarboxylation reaction (Scheme 2) with a charge-delocalized transition state. This reaction exhibits high sensitivity to the surrounding medium, where the first-order rate constant in aqueous solution is enhanced 106 -fold in aprotic dipolar solvents [61]. Thus, this reaction is a particularly useful model for studying medium effects and control of the microenvironment in a heterogeneous binding pocket [62, 63]. Antibody 21D8 accelerates the decarboxylation reaction by 2 × 104 (Table 1) [62, 64]. The hapten design incorporated two sulfonate moieties to mimic the carboxylate anion of the substrate. Not surprisingly, this antibody has an anionic binding site consisting of HisH35 and ArgL96 [14]. An additional anionic binding site was observed for the second sulfonate group on the opposite side of the binding pocket. However, assigning which of these anionic

7

8 Structure and Function of Catalytic Antibodies

Scheme 2 Decarboxylation Antibody 21D8

binding sites was relevant for binding to the substrate carboxylate was hindered because of the unfortunate symmetry of the hapten. Moreover, attempts at cocrystallization with the substrate were unsuccessful [14]. However, mutagenesis studies support the hypothesis that HisH35 and ArgL96 are indeed key substratebinding residues [65]. It is proposed that the antibody shields the substrate from solvent, but the tetrahedral hydrogen-bonding pattern of the anionic binding site is not fully optimized for binding a planar carboxylate [14, 65]. The pocket is polar with apparently non-optimal hydrogen bonds to the transition state, which may be reminiscent of a dipolar aprotic solvent. Such a model would explain, in part, how the antibody and other enzymes could control the microenvironment to facilitate catalysis. 2.7 Anionic Binding Motifs and the Periodate Cofactor in Sulfide Oxygenation

Catalytic antibody 28B4 catalyzes a bimolecular reaction between sodium periodate (NaIO4 ) and a sulfide to form a sulfoxide (Scheme 3, Table 1) [66]. Oxygenation reactions in naturally-occurring enzymes normally require flavin, heme, or other cofactors as well as NADPH to regenerate the cofactor [67]. Here, periodate is exploited as an artificial chemical cofactor, readily available at high excess and obviating the need for cofactor recycling. The oxygenation reaction is believed to involve nucle-

Scheme 3 Sulfide oxygenation Antibody 28B4

2 Electrostatic Complementarity

ophilic attack of a sulfur lone pair of electrons on a periodate oxygen through a polar transition state (Scheme 3). The amine in the hapten mimics the partial positive charge that develops on the sulfur, while the phosphate mimics the tetrahedral periodate. The crystal structure reveals an anionic binding site similar to that seen in the phosphocholine antibody McPC603 [8, 9] that coordinates periodate (Fig. 2a) [68]. However, the baiting strategy employed to stabilize the developing positive charge on the sulfur was not as effective as the postulated cation-π interaction with a nearby tyrosine does not have optimal geometry (Fig. 2a). Extensive mutagenesis has been performed on this antibody, and the putative germline antibody was also cloned and crystallized in complex with hapten [69]. Here, the p-nitrophenyl portion of the hapten adopts a completely different orientation in the binding pockets of the and affinity-maturated antibodies [69]. In addition, the free and bound forms of the germline antibody structure show significant conformational differences in CDRH3 and CDRL1, while comparison of the free and bound mature antibody shows no significant changes on binding [69]. Thus, the germline is postulated to exhibit wider structural plasticity than the mature antibody [69]. The loss of induced fit binding during the affinity maturation process has been noted in at least two other antibodies [22, 70], but does not rule out the importance of induced fit in affinity-matured antibodies [1].

Fig. 2. Top and side views of sulfide oxygenation and allylic isomerization antibodies, respectively. A Antibody 28B4 has an anion-binding pocket that binds the cofactor periodate in a biomolecular reaction with sulfide to form sulfoxide. This pocket was elicited by a phosphate functionality in the immunizing hapten. B Antibody 4B2 catalyzes an isomerization reaction of a $-( unsaturated ketone. GluL34 acts as general acid/base in the reaction by abstracting the proton from the substrate. Here, GluL34 interacts with the amidinium part of the hapten.

9

10 Structure and Function of Catalytic Antibodies

Scheme 4 Allylic isomerization Antibody 4B2

2.8 Cation Stabilization and Allylic Isomerization

Several antibodies have been elicited that are believed to have mechanistically important, negatively charged carboxylates in their binding pocket. In principle, such carboxylates may be elicited from a baiting strategy that incorporates positivelycharged substituents in the immunizing hapten. Few structures of this type have been determined, however, and for at least one of these antibodies (1D4), the baiting strategy failed. Antibody 4B2 catalyzes an allylic rearrangement of a β–γ unsaturated ketone (Scheme 4, Table 1). Formally, the reaction would be a [1, 3] sigmatropic migration of hydrogen. An amidinium group was incorporated in the hapten design to elicit a carboxylate general acid/base in the binding pocket (Scheme 4) [71]. The crystal structure of 4B2 shows that this baiting strategy was successful, as the two NH groups of the amidinium form H-bonds to each oxygen of the GluL34 carboxylate (Fig. 2b) [72]. The pH rate profile is bell shaped with optimal activity at pH 4.5, the approximate pK a of glutamic acid. An enol intermediate was proposed, based on lack of any antibody-catalyzed 2 H exchange between α and γ carbons in a deuterated substrate, implying that the reaction is not concerted, but involves exchange with solvent [71]. This is a reasonable hypothesis, although it is unlikely that any antibody residue would be positioned to stabilize this transient enol intermediate [72].

3 Shape Complementarity and Approximation

Shape complementarity simply refers to the similarity between the contours of the binding pocket molecular surface and that of its corresponding ligand. Certainly, it is possible to conclude from most of the catalytic antibody structural studies that exquisite shape complementarity is sufficient to explain the stereoselectivity for the majority of these catalysts. Thus, when the catalysts are challenged with chiral (or prochiral) substrates, chiral products are obtained, typically in high enantiomeric excess (ee). In addition, every reaction requires, to some extent, approximation to bring reactive centers together to either break or make new bonds. The antibody molecule is ideally tailored for this role, as it may be programmed to bind flexible substrates in a particular conformation that would otherwise be disfavored in solution, and can bring two substrates together in bimolecular reactions to increase the effective molarity of the substrates relative to one another. In principle,

3 Shape Complementarity and Approximation

approximation lowers the entropy barrier for activation and can effect catalysis. Thus, electrostatic and overall shape complementarity of the binding pocket for the transition state is a key feature for any catalyst and is one of the most important features cited for many catalytic antibody structures. However, approximation alone probably does not fully explain catalysis in all of these systems. Solvent accessibility, control of the microenvironment dielectric, and hydrogen-bonding potential are also important parameters – some reactions are accelerated simply by changing the solvent system. Indeed, not surprisingly, nonspecific interactions with bovine serum albumin (BSA) can accelerate some reactions [73]. In addition, key catalytic residues form specific interactions essential for the reaction to proceed. Catalytic residues have been elicited with mixed success by TSA baiting strategies, reactive immunization, and sometimes by pure chance. Notwithstanding, approximation will be a recurrent theme throughout the remaining sections. 3.1 Unimolecular Rearrangements 3.1.1 Antibody 1F7 Antibody 1F7 catalyzes the [3,3] sigmatropic Claisen rearrangement of chorismate to prephenate (Scheme 5, Table 1) [74]. This reaction is normally catalyzed by chorismate mutase in the biosynthesis of aromatic amino acids. The enzymatic reaction proceeds through an ordered chair-like transition state (S‡ ≈ −13 cal mol−1 K − 1 ) with a rate acceleration of 3 × 106 or four orders of magnitude better than the antibody [75]. Remarkably, even with this modest activity, over-expression of 1F7 complements growth of yeast auxotrophs [76]. The original structure was determined at 3.0 Å resolution from data merged from two crystals collected at room temperature [77]. More recent (unpublished) data at 2.4 Å show some differences in the detailed interactions between the TSA and Fab [77]. However, the general placement of the hapten on the heavy chain side of the antibody is maintained and illustrates a clear and interesting difference in the orientation of the TSA in the antibody pocket as opposed to the enzyme active site [78]. For the antibody, the hapten-binding orientation is determined by the placement of the linker used for covalent attachment to the immunogen, while the enzyme has no such restriction (Scheme 5). The enzyme also utilizes more H-bond interactions than the antibody, which are important for stabilizing the dipolar transition state [77]. Thus, the reaction is probably being catalyzed by preferentially binding the substrate in the

Scheme 5 Chorismate mutase Antibody 1F7

11

12 Structure and Function of Catalytic Antibodies

requisite reactive conformation, while poorly optimized hydrogen bond interactions may account for the inefficiency of the antibody catalyst. This theme recurs in which the substrate in an antibody is inserted upside down compared to that in the natural enzyme [47, 79]. 3.1.2 Antibody AZ-28 AZ-28 catalyzes a related [3,3]sigmatropic oxy-Cope rearrangement (Scheme 6, Table 1) [80]. The chair-like transition state is highly ordered, requiring overlap of the rearranging 4π+ 2σ orbitals of the diene (S‡ ≈ −15 cal mol−1 K−1 ). Antibodies raised against the appropriate, conformationally-restricted TSA would be expected to catalyze the reaction by acting as an “entropy trap.” In addition, appropriate alignment of the 2- and 5-phenyl rings are believed to contribute stereoelectronically to the reaction through hyperconjugation with the 3-hydroxyl (Scheme 6) [81]. This hyperconjugation may accelerate the oxy-Cope rearrangement by an anionic substituent effect. Indeed, the crystal structure shows that packing interactions with the 2- and 5-phenyl rings constrain the cyclohexyl group in the required chairlike transition state (Fig. 3). However, the cyclohexyl ring of the TSA is rotated ca. 80◦ out of plane of the 2- and 5-phenyl rings, so that the phenyl π-orbitals do not align appreciably with the 3-hydroxyl (Fig. 3) [81]. The orientation of the cyclohexyl group is determined by packing interactions, as well as by hydrogen bonds to the 3-hydroxyl (Fig. 3). Surprisingly, biochemical analysis revealed that six mutations occurred during affinity maturation, one of which dramatically decreases the catalytic efficiency [81]. Thus, the evolution of an antibody binding pocket with increased affinity to the TSA does not always correlate with improved catalysis. The germline-encoded antibody structure was determined and revealed an induced-fit mode of binding, as opposed to a lock-and-key type binding in the mature antibody. This apparent conformational flexibility and corresponding decrease in affinity is interpreted as evidence for possible torsional flexibility in the 2-phenyl substituent [82]. This flexibility, in turn, is postulated to facilitate π-overlap and explain the

Scheme 6 Oxy-Cope Antibody AZ-28

3 Shape Complementarity and Approximation Fig. 3. Side view of oxy-Cope antibody AZ-28. The cyclohexyl ring of the TSA is rotated out of plane with the 2- and 5-phenyl rings because of packing interactions and hydrogen bonds to the 3-hydroxyl. The consequent lack of hyperconjugation in the rearranging B-orbitals may explain the decreased rate acceleration in the affinity mature antibody, as opposed to the germline Fab.

improved catalysis. Studies to characterize the dynamics of the antibody catalyzed reaction would provide further evidence to support this hypothesis. 3.2 Bimolecular Rearrangements and the Diels-Alder Reaction

The D-A reaction is one of the most important carbon-carbon bond-forming reactions that is available for organic syntheses of complex natural products and therapeutic agents [83, 84]. Surprisingly, the D-A reaction has not been exploited widely in nature as only a handful of natural D-A enzymes are known [85]. The reaction is a pericyclic [4π+ 2π] cycloaddition of a diene and dienophile. An optimal D-A reaction occurs between an electron-rich diene (HOMO - highest occupied molecular orbital) and an electron-deficient dienophile (LUMO - lowest unoccupied molecular orbital). These abzyme-catalyzed bimolecular reactions are typically much slower and have much higher K M s than esterolytic antibodies (Table 1). The transition state for the D-A reaction is highly ordered for which S‡ is typically -30 to -40 entropy units [86]. Consequently, the recurring theme for these D-A antibodies is shape complementarity; in general, the antibodies with higher shape complementarity to the transition state are better catalysts. Thus, these antibodies may be considered as model systems for studying the effects of approximation in catalysis. 3.2.1 Retro-Diels-Alder Reaction Antibody 10 F11 catalyzes a unimolecular retro-D-A reaction that expels the dieneophile nitroxyl and diene anthracene (Scheme 7, Table 1). The dihedral angles between the phenyl rings in the substrate (110◦ ) and product (180◦ ) differ by 70◦ [87]. A TSA was designed with an intermediate dihedral angle of 140◦ [87]. Four crystal structures were determined including substrate analog (SA), transition state analog (TSA), and product analog (PA) [88]. The kinetics of the reaction suggest that binding energy is converted to catalysis with high efficiency (Table 1), with a clear hierarchy in the shape complementarity index between the antibody and ligands, ranked as TS > SA > PA [88]. This ranking supports the hypothesis that the antibody exerts strain on the substrate to preferentially stabilize the transition state.

13

14 Structure and Function of Catalytic Antibodies

Scheme 7 Retro Diels-Alder Antibody 10F11

Antibody 9D9 catalyzes the same reaction with slower kinetics, which could result from poorer overall shape complementarity [88]. The CDRH3 loop also undergoes a slight induced-fit con-formational change from binding the substrate to the transition state [88]. Finally, a water-mediated hydrogen bond was observed between the antibody backbone and the oxygen that is released from the substrate in the nitroxyl dieneophile product [88]. A second hydrogen bond is formed directly between the antibody backbone and the same oxygen of the substrate. These hydrogen bonds are postulated to act as Lewis acids that further stabilize the LUMO energy of the dieneophile and contribute to the regioselectivity of the retro-D-A reaction [88]. 3.2.2 Disfavored exo Diels-Alder Reaction For antibody 13G5, a novel hapten design was examined (Scheme 8, Table 1). Rather than using a conformationally-restricted TSA, an unrestrained ferrocene derivative was used to test whether the immune system would select a conformer that mimics one of the Diels-Alder transition states (Scheme 8) [89]. In principle, a panel of catalysts would be generated, each of which would selectively catalyze the formation of one of the four possible ortho-diastereomers. In fact, of all the antibodies examined, only one enantiomer was formed, indicative of possible structural bias in the immune response [90]. The catalyzed reaction proceeds with high regio-, diastereo-, and enantioselectivity to form the ortho-exo (S,S) product with ee > 95% after the background exo D-A reaction is subtracted [90]. The background endo:exo product distribution is 85:15 in PBS and 66:34 in toluene, so the exo-transition state is slightly disfavored by ca. 1 kcal mol−1 . The antibody rate acceleration is modest with a kcat(exo) kuncat(exo) −1 of 6.9 M and a kcat(exo) kuncat(endo) −1 of only 1.7 M [89, 91].

Scheme 8 Diels-Alder Antibody 13G5

3 Shape Complementarity and Approximation

Thus, the background reaction to form unwanted ortho-endo product will dominate the final product distribution of the antibody-catalyzed reaction unless sufficiently high concentrations of antibody catalyst are used [90]. The crystal structure of 13G5 shows the cyclopentadienyl ferrocene deeply buried in the antibody combining site [91]. Three H-bonds are observed between the TSA and TyrL36, AspH50, and AsnL91. TyrL36 was initially hypothesized to act as a Lewis acid, activating the dienophile for nucleophilic attack, while AsnL91 and AspH50 form H-bonds to the carboxylate side chain that substitutes for the diene substrate [91]. However, the limited rate accelerations indicate that catalytic residues in the binding pocket have a modest affect, if any, on the kinetics. Rather, the observed diastereo- and enantioselectivity of the antibody could be explained simply by exclusion of solvent (e.g., note toluene product distribution) and specific van der Waals interactions and hydrogen bonds that anchor the substrates in a particular orientation that favors ortho-exo (S,S) product formation due to approximation [86]. The performance of this catalyst may be explained by the relatively poor shape complementarity between the antibody and the true transition state of the reaction, which differs substantially from the hapten as the distance between the cyclopentadienyl rings in the ferrocene structure is ca. 3.3 Å. But what is truly remarkable is that a specific antibody could be generated against such a flexible hapten. 3.2.3 Endo Diels-Alder Reactions Antibody 39-A11 was elicited from immunization with a constrained bicyclic hapten (Scheme 9), and selectively accelerates formation of the kinetically-favored endo product [92]. The acceleration over the background or effective molarity is modest (Table 1). The X-ray structure of the hapten complex indicates that the antibody would bind the diene and dienophile in a reactive conformation, as preprogrammed in the hapten design strategy. The structure also suggests that the catalyst is enantioselective, although an ee has not been determined [93]. AsnH35 forms a hydrogen bond with the dienophile, which could make it more electron deficient and, thereby, more reactive [93]. Interestingly, any evident shape complementarity is lacking between antibody and the endo transition state; the binding pocket can accommodate either the endo or exo transition states (Fig. 4a). Mutations were designed to improve the packing interactions around the kinetically-favored endo transition state, and three of six mutants led to a 5- to 10-fold improvement in kcat [93]. This successful structure-based engineering is a significant and a rare accomplishment in this field. Also, only two somatic mutations were required to generate this catalytic antibody from the germline precursor. The most relevant of these two mutations is SerL91Val (CDR3), located in the binding pocket. The K DTSA for the germline antibody is 380 nM compared to 130 nM in the mature 39-A11. As expected, the catalytic proficiency of the germline antibody is also slightly decreased [93]. In order to circumvent substrate or product inhibition, more sophisticated hapten design strategies incorporate extraneous chemical groups as a negative design, while still promoting high affinity interactions to the transition state (Scheme 10) [94]. 1E9 was originally developed to evaluate the role of proximity

15

16 Structure and Function of Catalytic Antibodies

Scheme 9 Diels-Alder Antibody 39-A11

effects in catalysis, and is the most efficient D-A catalytic antibody yet developed (Table 1) with an effective molarity (EM) of 103 M, where the theoretical limit is 108 M [95]. Two structural features can explain the mechanism of the observed antibody catalysis. First is the conserved H-bond formed between AsnH35 and the succinimide carbonyl oxygen of the hapten, which represents the carbonyl oxygen of the dienophile substrate (Scheme 10). Since an H-bond to the carbonyl oxygen should cause the dieneophile to be electron deficient, it is more susceptible to the attack by the diene. Second, the binding pocket features almost perfect van der

Fig. 4 Molecular surface representation of Diels-Alder antibodies. Antibody 39-A11 (A) and 1E9 (B) are derived from the same germline precursor, yet exhibit a large difference in activity. This difference is attributed to the poor shape complementarity of the

antibody for the transition state in 39-A11 compared to 1E9. Mutations were designed in 39-A11 to improve the shape complementarity, and several improved catalysts were identified.

4 Shape Complementarity and Control of the Reaction Coordinate

Scheme 10 Diels-Alder Antibody 1E9

Waals complementarity to the hapten, providing a clear structural demonstration of approximation in bimolecular catalysis (Fig. 4b) [96]. Interestingly, 1E9, antiprogesterone antibody DB3, and the D-A antibody 39A11 are derived from the same germline gene [93, 97–99]. Structures of these antibodies thus represent possible snapshots in the evolution of substrate binding and catalytic efficiency [94]. Subtle differences in the evolution of their binding pockets through somatic mutations have resulted in antibodies that specifically bind progesterone or catalyze a D-A reaction. The exceptional efficiency of 1E9 is hypothesized to arise from a rare somatic mutation that significantly deepens the active site, creating exquisite complementarity to the transition state [94].

4 Shape Complementarity and Control of the Reaction Coordinate

In many reactions, several products may be formed, but one product predominates because it is favored kinetically. The energy of activation is much lower for the favored product, which reduces the likelihood of forming the disfavored product. If the activation barrier for the reverse reaction is sufficiently large, then the reaction is essentially irreversible, or under kinetic control. Similarly, for some reactions, many possible products may result (e.g., the D-A reaction or cationic cyclization), and complex mixtures result. The antibody catalyst can steer a reaction through a complex reaction coordinate to alter the final reaction profile, often forming exclusively a single product, and, in some cases, a kinetically-disfavored one. 4.1 Syn-Elimination

Antibody 1D4 is a model system for studying the selective catalysis of a disfavored syn-elimination reaction toform a cis product (Scheme 11, Table 1) [100]. The TSA design incorporated a rigid bicyclic ring structure to constrain the two phenyl groups of the substrate in the eclipsed conformation that characterizes the transition state (Scheme 11). In addition, the α-keto proton that is abstracted during elimination was replaced with a positively-charged amine in the hapten to elicit a complementary general base in the antibody combining site. Although

17

18 Structure and Function of Catalytic Antibodies

Scheme 11 Syn-elimination Antibody 1D4

the catalyzed reaction proceeds slowly (Table 1) [100], the background elimination reaction to the cis-product is undetectable in the absence of antibody; the competing anti-elimination reaction (k = 2.5 × 10−4 min−1 ) dominates the background reaction, such that the product distribution is entirely trans. Crystal structures of free and bound 1D4 show a conformational change in CDR H2, indicative of induced fit [101]. The pocket exhibits high shape complementarity to the TSA, which accounts for the selectivity for the eclipsed transition-state of the reaction (Fig. 5) [101]. Hence, the antibody combining site likely utilizes the majority of its binding energy toward overcoming the extreme steric barriers, and, hence, stabilizing the eclipsed transition state. The key mechanistic issue for any βelimination process is whether the reaction is E1, E2, or E1cB [102]. Unfortunately, a crystal structure alone cannot resolve such a question. A water molecule was modeled into electron density that is near the hapten amine [101] and within

Fig. 5 Electrostatic potential mapped to the molecular surface of 1D4. The exquisite shape complementarity for the phenyl rings that characterize the eclipsed transition state of the reaction accounts for the selectivity of the catalyst.

4 Shape Complementarity and Control of the Reaction Coordinate

hydrogen-bonding distance to a histidine. Thus, HisH58 is proposed to promote catalysis through interaction with the water [101]. 4.2 Disfavored Ring Closure

Antibody 5C8 catalyzes the disfavored endo-tet cyclization reaction inviolation of Baldwin’s rules for ring closure (Scheme 12) [15]. The N-methylpiperidinium hapten was anticipated to elicit negative charge in the binding pocket that would stabilize the developing positive charge at the oxirane carbon of the substrate. Several changes were observed between the bound and unbound structures, indicative of induced fit. The largest rearrangement was observed in the backbone and side chain orientations of CDR H3 with the largest differences seen for the two tyrosine residues at the tip of the loop, which fold into the binding site, creating a solvent-inaccessible cavity. AspH95 and AspH101 bind the positive charge of the quaternary amine like a pair of tweezers (Fig. 6) [15]. In addition, the positive charge appears to be stabilized by a cation-π interaction with TyrL91. AspH95 and HisL89 were hypothesized to effect rudimentary general acid/base catalysis (Fig. 6). In the proposed mechanism, AspH95 forms an H-bond to the epoxide-oxygen, favoring the oxirane ring opening, while HisL89 forms an H-bond with the alcohol to promote nucleophilic SN 2-like attack to form the disfavored six-membered ring [15]. This stabilization of the positive charge that develops along the reaction coordinate appears to be an important factor for the rate enhancement and for directing the reaction along the otherwise disfavored pathway [15]. 4.3 Selective Control of a Reactive Carbocation: Antibodies 4C6 and 19A4

Cationic cyclization proceeds via formation of a reactive carbocation, and may be separated into three steps: initiation, propagation, and termination. A carbocation may be formed either by ionization or by electrophilic addition to an unsaturated olefin, while cyclization is terminated by nucleophilic addition or elimination of a proton. The stereochemistry of such transformations should follow the StorkEschenmoser hypothesis [103, 104]. Natural enzymes catalyze polycyclization of squalene through a reactive carbocation intermediate to form a variety of cycloisoprenoids (e.g., cholesterol). For such enzymes, initiation occurs either by protonation of a double bond or by release of pyrophosphate [105]. Structural studies have shown that the enzyme active sites are typically lined with aromatic residues that

Scheme 12 Disfavored cyclization Antibody 5C8

19

20 Structure and Function of Catalytic Antibodies

Fig. 6 Disfavored ring closure antibody 5C8. AspH95 is proposed to form an Hbond with the epoxide-oxygen, favoring oxirane ring opening, with HL89 forming an H-bond with the alcohol to promote SN2-

like attack to form the disfavored ring. The piperidinium moiety was used as bait to elicit the desired negative charge in the binding pocket, which should stabilize the developing positive charge in the actual ring closure reaction.

stabilize the carbocation through cation-π interactions. Catalytic antibodies 4C6 and 19A4 also contain a number of aromatic residues in the binding pocket, which are postulated to play a similar role. Thus, the enzymes and antibodies exhibit similar features. Catalytic antibody 4C6 catalyzes a model reaction that had been shown under other conditions (98% formic acid) to proceed through a carbocation [106]. This reactivity was elicited with an N-oxide hapten [107] that is structurally related to the 5C8 hapten. Structurally analogous N-oxide TSAs are known inhibitors for natural squalene cyclases, validating this design strategy [108]. The oxide was included to promote stabilization of the negative charge that develops in the sulfonic acid leaving group, while the nitrogen would elicit residues that increase the electrophilicity at C1 (Scheme 13, Table 1). The hapten also includes a silane moiety that could stabilize the carbocation via hyperconjugation [107]. In addition, conjugation with silane makes the olefin more electron rich, which facilitates attack of C1 . Carbocations are highly reactive species, yet the antibody selectively catalyzes the formation of only one product in mild conditions where no background reaction is detectable (Scheme 13) [107]. The crystal structure shows several aromatic residues in the active site that could stabilize the reactive carbocation and shield it from solution [17]. In the proposed mechanism, TyrL96 and/or TyrH50 initiate carbocation formation at C1 by promoting SN 1-like departure of the acetamidobenzenesulfonic acid leaving group. Electrophilic attack of the C5 -C6 olefin by the developing C1 carbocation closes the ring and shifts the carbocation to C5 , where the reaction terminates by nucleophilic addition of water [17]. This mechanism, however, does not explain why a related compound (Si exchanged for C) has a ca.100-fold reduction in

4 Shape Complementarity and Control of the Reaction Coordinate

Scheme 13 Cationic cyclization Antibody 4C6

kcat , but similar K M for the reaction, nor why the related substrate that lacks the C5 C6 double bond exhibits no reaction (kinetics monitor formation of leaving group). Together, these data suggest that anchimeric assistance of the electron-rich double bond is the central feature of the mechanism [107]. Thus, the reaction could be initiated by backside attack of C1 by the electron-rich olefin, followed by termination by nucleophilic addition of water at C5 . This attack is facilitated by approximation due to the shape complementarity of the binding pocket. Additional kinetics and mutagenesis could help resolve the details of how the reaction is initiated. Catalytic antibody 19A4 catalyzes a more complicated, tandem-cationic cyclization to form bridge-methylated decalins (Scheme 14, Table 1) [109]. The hapten design was similar to that for 4C6, with the same N-oxide functionality to elicit residues that would promote initiation of the cyclization cascade by expulsion of the sulfonic acid leaving group. In contrast to 4C6, this reaction is terminated by elimination. The antibody does not possess the exquisite control over the reaction that was observed for 4C6, as three different decalins are formed accounting for only 50% of the overall products [109]. Nevertheless, the catalysis is remarkable, as no bicyclic products are formed in the uncatalyzed reaction. The crystal structure shows a hydrophobic binding pocket lined with several aromatic residues that may stabilize the carbocation [16]. In addition, the binding pocket has high shape complementarity to the productive chair-chair conformation of the substrate that is required for effective cyclization (Scheme 14). Here, a hydrogen bond from

Scheme 14 Cationic cyclization Antibody 19A4

21

22 Structure and Function of Catalytic Antibodies

AsnH35a to the sulfonic acid leaving group is believed to facilitate SN 1-like departure with concerted anchimeric assistance from the C5-C6 π bond, yielding the first ring with the carbonium ion at the tertiary C5 carbon [16]. In the next step, the C9-C10 π bond undergoes electrophilic attack by the C5 carbonium to form the second ring. Although an epoxide was included in the original hapten to elicit a catalytic residue in the antibody, there is no catalytic base to direct the regiochemistry of the final termination step of the reaction [16]. Thus, the reaction results in a mixture of elimination products. Structure-based engineering would afford a unique opportunity to control the product distribution for this particular reaction.

5 Shape Complementarity and Substrate Strain

Many catalytic antibodies impose varying degrees of strain on the substrate to achieve reactive conformations that would otherwise be disfavored in solution (e.g., 1D4, 4C6, 19A4, 13G5, 10F11, AZ-28). The enzyme ferrochelatase, however, is a particularly attractive model system for studying substrate strain. Ferrochelatase inserts Fe2+ into porphyrin in heme biosynthesis. In this reaction, the enzyme is proposed to distort the porphyrin ring system to expose the pyrrole nitrogen lone pair electrons, facilitating metal ion complexation. Indeed, N-methyl protoporphyrin – a distorted porphyrin analog – is a potent inhibitor of ferrochelatase [110]. Antibodies elicited against this compound catalyze insertion of Cu2+ or Zn2+ into mesoprotoporphyrin IX, supporting the hypothesis that the hapten is a true transition state analog for the enzyme-catalyzed reaction (Scheme 15, Table 1) [111]. Thus, this reaction is a model system for studying the effects of substrate strain in catalysis [4]. In the antibody-catalyzed reaction, no saturation kinetics were observed for the metal ion, suggesting that metal binding to the antibody is not a prerequisite of catalysis, in contrast to ferrochelatase, which has a K M of 32 µM for iron [112]. The crystal structure, however, reveals that an Asp carboxylate is directed toward

Scheme 15 Metal chelation antibody 7G12

6 Reactive Amino Acids and the Possibility of Covalent Catalysis

Fig. 7 Superposition of free (green) and TSA bound (blue) 7G12. (A) The germline 7G12 shows an induced-fit mode of binding, as demonstrated by the conformation change in CDRH3. (B) The affinity mature

antibody, in contrast, has a lock-and-key mode of binding. The maturation of a preformed binding pocket is associated with higher affinity for the transition state and overall improved catalytic proficiency.

the center of the protoporphyrin (Fig. 7) and is proposed to either guide metal complexation or act as a proton shuttle [113]. The residue is clearly important as Asp → Asn and Asp → His mutations are inactive [113]. In addition, the crystal structure shows that packing interactions between the antibody and substrate induce strain in the Michaelis complex [70]. Although accurate modeling and exact refinement of any distortions in the Michaelis complex at 2.6 Å resolution are not reasonable, the electron density does provide clear qualitative indications of deviation from ideal planarity [70]. The free and hapten-bound structures of the germline and mature antibodies were compared and nicely illustrate the evolution from an induced fit mode of binding to lock-and-key (Fig. 7a,b) [70]. The evolution of this lock-and-key binding results in tighter binding to the TSA (due to faster association and slower dissociation rates), and correlates with a ca. 100-fold improvement in kcat K M −1 [70]. Together, these structural and biochemical results demonstrate the importance of substrate strain in antibody-mediated catalysis.

6 Reactive Amino Acids and the Possibility of Covalent Catalysis

The recently developed reactive immunization strategy [114, 115] has shown that it is possible to create antibodies that catalyze aldol reactions, one of the most important carbon-carbon bond-forming reactions in chemistry and biology. The mechanism for the natural class I aldolase enzyme involves the key formation of a Schiff base between the substrate ketone (aldol donor) and a reactive lysine, followed by condensation with an aldehyde (aldol acceptor) and subsequent hydrolysis to release product (Scheme 16). The binding pocket of aldolase antibody 33F12 is an elongated cleft more than 11 Å deep [116]. The only lysine residue in the active site

23

24 Structure and Function of Catalytic Antibodies

Scheme 16 Previously proposed retro-aldol mechanism for aldolase antibody 33F12

is located at the bottom of the hydrophobic pocket (Fig. 8a). This environment was hypothesized to perturb the pK a of the lysine, making it amore potent nucleophile [116]. The antibody can then use the reactive -amino group to form an enamine with the corresponding substrates, analogous to the natural class I aldolase. The LysH93 → Ala mutant is inactive, which confirms the importance of this residue [117]. An additional residue that may be of mechanistic interest is TyrL36, which forms a water-mediated hydrogen bond with the -amino group of lysine H93 (Fig. 8a). Importantly, a water molecule was postulated to play a key role shuttling protons in the mechanism of a class I aldolase [118]. Interestingly, the antibody exhibits exceptionally broad substrate specificity, implying that the binding pocket is unrefined for a particular substrate presumably due to the lack of further affinity maturation due to the proposed covalent nature of the hapten complex [116]. Antibodies 40F12, 84G3, 93F3 and others were generated with a sulfone β-diketone hapten [119]. Notably, antibodies 93F3 and 84G3 show reversed enantioselectivity compared to 33F12 [119]. Sequence analysis and our unpublished structural data [120] reveal a lysine at position L89 on the opposite side of the binding pocket, which could explain the reversed selectivity of this antibody. The relatively unrefined binding pocket of 33F12 contrasts with that of class I aldolase enzymes, which catalyze the reversible condensation of aldehydes and ketones with high stereospecificity (121,122). The bacterial 2-deoxyribose-5-phosphate aldolase (DERA) (123,124) catalyzes the reversible aldol reaction of acetaldehyde and D-glyceraldehyde-3-phosphate to form 2-deoxyribose-5-phosphate. The ultra high resolution structure of DERA reveals a β-barrel with two lysines in the active site, one of which forms the covalent Schiff base intermediate (Fig. 8b) [118]. Compared to the hydrophobic antibody binding pocket, a number of charged/polar residues form a sophisticated hydrogen-bonding network within the active site, which is likely important for tuning the reactivity of the lysine and the substrate specificity [118]. The mechanism by which Lys167 is rendered nucleophilic in the enzyme is unclear – the pK a was proposed to be depressed by a neighboring Lys201 (Fig. 8b) [118], but mutation of Lys201 → Leu clearly does not hinder formation of the Schiff

6 Reactive Amino Acids and the Possibility of Covalent Catalysis

base [118]. Surprisingly, inhibition of natural aldolases by diketones has not been fully characterized. A lingering challenge is to determine the mode of binding of the aldolase family of antibodies to substrate-related inhibitors, as well as the diketone compound originally used as a hapten to elicit the antibody response (Scheme 16). Despite our best efforts, we have obtained no structural evidence demonstrating the formation

Fig. 8 Comparison of aldolase antibody and enzyme active sites. Red spheres represent water molecules. A The aldolase antibody contains a lysine buried at the bottom of hydrophobic gorge, which has been postulated to form covalent intermediates with an aldol donor (ketone) followed by condensation with an aldol acceptor (aldehyde). TyrL36 may form a water-mediated hydrogen bond with the reactive lysine ,amino group, which may be of importance in the reaction mechanism. B The bacte-

rial 2-deoxyribose-5-phosphate aldolase (DERA) likewise contains a reactive Lys167, which was observed to form a covalent intermediate with the substrate in the crystal structure. Lys167 is probably rendered nucleophilic by the sophisticated hydrogenbonding network in the active site, which includes nearby Asp102 and Lys201. A nearby water molecule forms key stabilizing interactions to the substrate, and is believed to play a key role in the catalytic mechanism.

25

26 Structure and Function of Catalytic Antibodies

of a covalent bond in the antibody binding pocket for the aldolase antibody. This contrasts with the relative ease by which we and others have obtained covalent adducts in enzyme systems [118]. Surprisingly, the Lys167Leu DERA mutant catalyzes the retro aldol reaction for its optimal substrate (3.6 min−1 ) [118] three times faster than the antibody for its optimal substrate (1.4 min−1 ) [125]. Clearly, catalysis of the retro-aldol condensation may occur in the absence of covalent intermediates. Alternative mechanisms for 33F12 and 38C2 were never seriously considered [116], but could involve non-covalent stabilization of an enolate or aldol donor that subsequently attacks the aldol acceptor (analogous to a class II aldolase). Such a mechanism would be more consistent with our structural observations. In any event, the mechanistic supposition that a lysine could be rendered nucleophilic in the hydrophobic environment of the antibody binding pocket led, in part, to the appreciation that secondary amines, e.g., proline, could catalyze the aldol reaction in organic and, to lesser extent, aqueous solvent [126]. Thus, structural and biochemical characterization of aldolase antibodies has led to the development of important new synthetic methodologies.

7 Conclusions

Classically, antibodies are effector molecules that link the recognition elements of the Fab to the destructive elements of the immune system. Recently, this paradigm has been challenged by the discovery that antibodies also possess the intrinsic capability to catalyze the formation of hydrogen peroxide from singlet oxygen and water in the absence of any other metal ions or cofactors [127, 128]. This catalytic capability would potentially link recognition and oxidative killing within the same molecule. This remarkable chemistry is likewise catalyzed by the T-cell receptor (TCR), but not β 2 -microglobulin, which shares the immunoglobulin fold [128]. Very little is understood about where this reaction occurs on the antibody or TCR, and docking studies have not resolved this issue [129]. Assaying light chain dimers and camel antibodies (heavy chain only) would provide some indication of where the chemistry occurs. We have grappled with this problem indirectly by pressurizing numerous antibody crystals under a xenon atmosphere [128]. Our data reveal several hydrophobic pockets in the core of the immunoglobulin fold. Since xenon is larger than oxygen (O2 ), oxygen could potentially diffuse into the same hydrophobic cavities. However, the mechanistic relevance of these hydrophobic cavities, if any, is not yet resolved. Unfortunately, a more direct experimental approach is extremely challenging; distinguishing unambiguously between water, singlet oxygen, hydrogen peroxide, ozone, and other reactive oxygen intermediates (which are believed to be short lived) from an electron density map is not feasible, even at ultra high resolution. Moreover, discriminating between radical chemistry derived from the antibody or synchrotron radiation is non-trivial and requires numerous control experiments. However, we have observed generally that soaking antibody crystals in hydrogen

References

peroxide and exposing them to UV light (source of singlet oxygen) has little effect on the integrity of the antibody [128], although oxidative modifications are observed on TrpLIG3 in the interfacial region between variable and constant domains [139]. Current endeavors aim to trace the origins of antibody-generated oxidants within the immunoglobulin fold.

References 1 WILSON, I. A., STANFIELD, R. L. Curr. Op. Struct. Biol. 4 (1994), p. 857–867 2 PADLAN, E. A., Adv.Prot.Chem. 49 (1996), p. 57–133 3 PAULING, L., Am.Sci. 36 (1948), p. 51–58 4 HALDANE, J. B. S., Enzymes, Longmans Green, London 1930 5 JENCKS, W. P., Catalysis in Chemistry and Enzymology, McGraw-Hill, New York 1969 6 TRAMONTANO, A., JANDA, K. D., LERNER, R. A., Science 234 (1986), p. 1566–1570 7 POLLACK, S. J., JACOBS, J. W., SCHULTZ, P. G., Science 234 (1986), p. 1570–1573 8 SEGAL, D. M., PADLAN, E. A., COHEN, G. H., RUDIKOFF, S., POTTER, M., DAVIES, D. R., Proc. Natl. Acad. Sci. U.S.A. 71 (1974), p. 4298–4302 9 SATOW, Y., COHEN, G. H., PADLAN, E. A., DAVIES, D. R., J. Mol. Biol. 190 (1986), p. 593–604 10 JESKE, D. J., JARVIS, J., MILSTEIN, C., CAPRA, J. D., J. Immunol. 133 (1984), p. 1090–1092 11 ROSE, D. R., STRONG, R. K., MARGOLIES, M. N., GEFTER, M. L., PETSKO, G. A., Proc. Natl. Acad. Sci. U.S.A. 87 (1990), p. 338–342 12 STRONG, R. K., CAMPBELL, R., ROSE, D. R., PETSKO, G. A., SHARON, J., MARGOLIES, M. N., Biochemistry 30 (1991), p. 3739–3748 13 STRONG, R. K., PETSKO, G. A., SHARON, J., MARGOLIES, M. N., Biochemistry 30 (1991), p. 3749–3757 14 HOTTA, K., LANGE, H., TANTILLO, D. J., HOUK, K. N., HILVERT, D., WILSON, I. A., J. Mol. Biol. 302 (2000), p. 1213–1225 15 BALDWIN, J. E., J.Chem.Soc.,Chem. Commun., (1976), p. 734–736 16 PASCHALL, C. M., HASSERODT, J., JONES, T., LERNER, R. A., JANDA, K. D.,

17

18

19 20

21

22

23

24

25

26

27

28

CHRISTIANSON, D. W. Angew. Chem. Int. Ed. 38 (1999), p. 1743–1747 ZHU, X., HEINE, A., MONNAT, F., HOUK, K. N., JANDA, K. D., WILSON, I. A. J. Mol. Biol. 329 (2003), p. 69–83 MACBEATH, G., and HILVERT, D., Chem. Biol. 3 (1996), p. 433–445 TANTILLO, D. J., HOUK, K. N. Chem. Biol. 8 (2001), p. 535–545 KRISTENSEN, O., VASSYLYEV, D. G., TANAKA, F., MORIKAWA, K., FUJII, I., J. Mol. Biol. 281 (1998), p. 501–511 WEDEMAYER, G. J., WANG, L. H., PATTEN, P. A., SCHULTZ, P. G., STEVENS, R. C., J. Mol. Biol. 268 (1997), p. 390–400 WEDEMAYER, G. J., PATTEN, P. A., WANG, L. H., SCHULTZ, P. G., STEVENS, R. C., Science 276 (1997), p. 1665–1669 GUO, J., HUANG, W., ZHOU, G. W., FLETTERICK, R. J., SCANLAN, T. S., Proc. Natl. Acad. Sci. U.S.A. 92 (1995), p. 1694–1698 ZHOU, G. W., GUO, J., HUANG, W., FLETTERICK, R. J., SCANLAN, T. S., Science 265 (1994), p. 1059–1064 BUCHBINDER, J. L., STEPHENSON, R. C., SCANLAN, T. S., FLETTERICK, R. J., J. Mol. Biol. 282 (1998), p. 1033–1041 CHARBONNIER, J. B., CARPENTER, E., GIGANT, B., GOLINELLI-PIMPANEAU, B., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., Proc. Natl. Acad. Sci. U.S.A. 92 (1995), p. 11721–11725 THAYER, M. M., OLENDER, E. H., ARVAI, A. S., KOIKE, C. K., CANESTRELLI, I. L., STEWART, J. D., BENKOVIC, S. J., GETZOFF, E. D., ROBERTS, V. A., J. Mol. Biol. 291 (1999), p. 329–345 CHARBONNIER, J. B., GOLINELLI-PIMPANEAU, B., GIGANT, B.,

27

28 Structure and Function of Catalytic Antibodies

29

30

31

32 33

34

35

36 37

38 39

40

41

42

43 44 45

TAWFIK, D. S., CHAP, R., SCHINDLER, D. G., KIM, S. H., GREEN, B. S., ESHHAR, Z., KNOSSOW, M., Science 275 (1997), p. 1140–1142 MIYASHITA, H., HARA, T., TANIMURA, R., FUKUYAMA, S., CAGNON, C., KOHARA, A., FUJII, I., J. Mol. Biol. 267 (1997), p. 1247–1257 GIGANT, B., TSUMURAYA, T., FUJII, I., KNOSSOW, M., Structure 7 (1999), p. 1385–1393 TAKAHASHI, N., KAKINUMA, H., LIU, L., NISHI, Y., FUJII, I., Nat. Biotech. 19 (2001), p. 563–567 STEWART, J. D., BENKOVIC, S. J., Nature 375 (1995), p. 388–391 TURNER, J. M., LARSEN, N. A., BASRAN, A., BARBAS III, C. F., BRUCE, N. C., WILSON, I. A., LERNER, R. A., Biochemistry 41 (2002), p. 12297–12307 MENARD, R., CARRIERE, J., LAFLAMME, P., PLOUFFE, C., KHOURI, H. E., VERNET, T., Biochemistry 35 (1991), p. 8924–8928 NICOLAS, A., EGMOND, M., VERRIPS, C. T., de VLIEG, J., LONGHI, S., CAMBILLAU, C., MARTINEZ, C., Biochemistry 35 (1996), p. 398–410 SZELTNER, Z., RENNER, V., POLGAR, L., Protein Sci. 9 (2000), p. 353–360 BRYAN, P., PANTOLIANO, M. W., QUILL, S. G., HSIAO, H.-Y., POULOS, T., Proc. Natl. Acad. Sci. U.S.A. 83 (1986), p. 3743–3745 CARTER, P., WELLS, J. A., Proteins: Struct. Funct. Genet. 7 (1990), p. 335–342 O’CONNELL, T. P., DAY, R. M., TORCHILIN, E. V., BACHOVCHIN, W. W., MALTHOUSE, J. G., Biochem. J. 326 (1997), p. 861–866 HAREL, M., QUINN, D. M., NAIR, H. K., SILMAN, I., SUSSMAN, J. L., J. Am. Chem. Soc. 118 (1996), p. 2340–2346 PHILLIPS, M. A., FLETTERICK, R., RUTTER, W. J., J. Biol. Chem. 265 (1990), p. 20692–20698 LINDLER, A. B., KIM, S. H., SCHINDLER, D. G., ESHAR, Z., TAWFIK, D. S., J. Mol. Biol. 329 (2002), p. 559–572 LINDLER, A. B., ESHAR, Z., TAWFIK, D. S., J. Mol. Biol. 285 (1999), p. 421–430 TANTILLO, D. J., HOUK, K. N., J. Comput. Chem. 23 (2002), p. 84–95 JANDA, K. D., ASHLEY, J. A., JONES, T. M., MCLEOD, D. A., SCHLOEDER, D. M.,

46 47

48 49 50

51

52

53

54

55

56

57 58 59 60

61 62

WEINHOUSE, M. I., LERNER, R. A., GIBBS, R. A., BENKOVIC, P. A., HILHORST, R., BENKOVIC, S. J., J. Am. Chem. Soc. 113 (1991), p. 291–297 DEREWENDA, Z. S., WEI, Y., J.Am. Chem. Soc. 117 (1995), p. 2104–2105 LARSEN, N. A., TURNER, J. M., STEVENS, J., ROSSER, S. J., BASRAN, A., LERNER, R. A., BRUCE, N. C., WILSON, I. A., Nat. Struct. Biol. 9 (2002), p. 17–21 GUO, J., HUANG, W., SCANLAN, T. S., J. Am. Chem. Soc. 116 (1994), p. 6062–6069 HAYNES, M. R., HEINE, A., WILSON, I. A., Isr. J. Chem. 36 (1996), p. 133–142 GHOSH, D., SAWICKI, M., LALA, P., ERMAN, M., PANGBORN, W., EYZAGUIRRE, J., GUTIERREZ, R., JORNVALL, H., THIEL, D. J. J. Biol. Chem. 276 (2001), p. 11159–11166 ZHU, X., LARSEN, N. A., BASRAN, A., BRUCE, N. C., WILSON, I. A., J. Biol. Chem. 278 (2003), p. 2008–2014 BACA, M., SCANLAN, T. S., STEPHENSON, R. C., WELLS, J. A., Proc. Natl. Acad. Sci. U.S.A. 94 (1997), p. 10063–10068 STEWART, J. D., KREBS, J. F., SIUZDAK, G., BERDIS, A. J., SMITHRUD, D. B., BENKOVIC, S. J., Proc. Natl. Acad. Sci. U.S.A. 91 (1994), p. 7404–7409 BENKOVIC, S. J., ADAMS, J. A., BORDERS Jr, C. L., JANDA, K. D., LERNER, R. A. Science 250 (1990), p. 1135–1139 GIBBS, R. A., BENKOVIC, P. A., JANDA, K. D., LERNER, R. A., BENKOVIC, S. J., J. Am. Chem. Soc. 114 (1992), p. 3528–3534 KREBS, J. F., SIUZDAK, G., DYSON, H. J., STEWART, J. D., BENKOVIC, S. J., Biochemistry 34 (1995), p. 720–723 SAMPSON, N. S., BARTLETT, P. A., Biochemistry 30 (1991), p. 2255 CARTER, P., WELLS, J. A., Nature 332 (1988), p. 564 BARTLETT, P. A., MARLOWE, C. K., Biochemistry 22 (1983), p. 4618–4624 CHRISTIANSON, D. W., LIPSCOMB, W. N., Acc. Chem. Res. 33 (1989), p. 62–69 KEMP, D. S., PAUL, K. G., J.Am. Chem. Soc. 97 (1975), p. 7305–7312 LEWIS, C., KRAMER, T., ROBINSON, S., HILVERT, D., Science 253 (1991), p. 1019–1022

References 63 TARASOW, T. M., LEWIS, C., HILVERT, D., J. Am. Chem. Soc. 116 (1994), p. 7959–7963 64 HOTTA, K., KIKUCHI, K., HILVERT, D., in Helv. Chim. Acta 83 (2000), p. 2183–2191 65 HOTTA, K., WILSON, I. A., HILVERT, D., Biochemistry 41 (2002), p. 772–779 66 HSIEH, L. C., STEPHANS, J. C., SCHULTZ, P. G., J. Am. Chem. Soc. 116 (1994), p. 2167–2168 67 HOLLAND, H. L., Chem.Rev. 88 (1988), p. 473–485 68 HSIEH-WILSON, L. C., SCHULTZ, P. G., STEVENS, R. C., Proc. Natl. Acad. Sci. U.S.A. 93 (1996), p. 5363–5367 69 YIN, J., MUNDORFF, E. C., YANG, P. L., WENDT, U., HANWAY, D., STEVENS, R. C., SCHULTZ, P. G., Biochemistry 40 (2001), p. 10764–10773 70 YIN, J., ANDRYSKI, S. E., BEUSCHER IV, A. E., STEVENS, R. C., SCHULTZ, P. G., Proc. Natl. Acad. Sci. U.S.A. 100 (2003), p. 856–861 71 GONCALVES, J., DINTINGER, T., LEBRETON, J., BLANCHARD, D., TELLIER, C., Biochem. J. 346 (2000), p. 691–698 72 GOLINELLI-PIMPANEAU, B., GONCALVES, O., DINTINGER, T., BLANCHARD, D., KNOSSOW, M., TELLIER, C., Proc. Natl. Acad. Sci. U.S.A. 97 (2000), p. 9892–9895 73 HOLLFELDER, F., KIRBY, A. J., TAWFIK, D. S., Nature 383 (1996), p. 60–62 74 HILVERT, D., CARPENTER, S. H., NARED, K. D., AUDITOR, M.-T. M., Proc. Natl. Acad. Sci. U.S.A. 85 (1988), p. 4953–4955 75 ANDREWS, P. R., SMITH, G. D., YOUNG, I. G., Biochemistry 12 (1973), p. 3492 76 TANG, Y., HICKS, J. B., HILVERT, D., Proc. Natl. Acad. Sci. U.S.A. 88 (1991), p. 8784–8786 77 HAYNES, M. R., STURA, E. A., HILVERT, D., WILSON, I. A., Science 263 (1994), p. 646–652 78 CHOOK, Y. M., KE, H., LIPSCOMB, W. N. Proc. Natl. Acad. Sci. U.S.A. 90 (1993), p. 8600–8603 79 LARSEN, N. A., ZHOU, B., HEINE, A., WIRSCHING, P., JANDA, K. D., WILSON, I. A., J. Mol. Biol. 311 (2001), p. 9–15 80 BRAISTED, A. C., SCHULTZ, P. G., J. Am. Chem. Soc. 116 (1994), p. 2211–2212

81 ULRICH, H. D., MUNDORFF, E., SANTARSIERO, B. D., DRIGGERS, E. M., STEVENS, R. C., SCHULTZ, P. G., Nature (1997), p. 271–275 82 MUNDORFF, E. C., HANSON, M. A., VARVAK, A., ULRICH, H. D., SCHULTZ, P. G., STEVENS, R. C. Biochemistry 39 (2000), p. 627–632 83 SAUER, J., Angew.Chem.Int.Ed.Engl. 5 (1966), p. 211–230 84 SCHMIDT, R. R., Acc.Chem.Res. 19 (1986), p. 250–259 85 POHNERT, G., Chembiochem 2 (2001), p. 873–875 86 PAGE, M. I., JENCKS, W. P., Proc.Natl. Acad. Sci. U.S.A. 68 (1971), p. 1678-1683 87 BAHR, N., GUELLER, R., REYMOND, J.-L., LERNER, R. A., J. Am. Chem. Soc. 118 (1996), p. 3550–3555 88 HUGOT, M., BENSEL, N., VOGEL, M., REYMOND, M. T., STADLER, B., REYMOND, J.-L., BAUMANN, U., Proc. Natl. Acad. Sci. U.S.A. 99 (2002), p. 9674–9678 89 YLI-KAUHALUOMA, J. T., ASHLEY, J. A., LO, C.-H., TUCKER, L., WOLFE, M. M., JANDA, K. D., J. Am. Chem. Soc. 117 (1995), p. 7041–7047 90 CANNIZZARO, C. E., ASHLEY, J. A., JANDA, K. D., HOUK, K. N., J. Am. Chem. Soc. 125 (2003), p. 2489–2506 91 HEINE, A., STURA, E. A., YLI-KAUHALUOMA, J. T., GAO, C., DENG, Q., BENO, B. R., HOUK, K. N., JANDA, K. D., WILSON, I. A., Science 279 (1998), p. 1934–1940 92 BRAISTED, A. C., SCHULTZ, P. G., J. Am. Chem. Soc. 112 (1990), p. 7430–7431 93 ROMESBERG, F. E., SPILLER, B., SCHULTZ, P. G., STEVENS, R. C., Science 279 (1998), p. 1929–1933 94 XU, J., DENG, D., CHEN, J., HOUK, K. N., BARTEK, J., HILVERT, D., WILSON, I. A., Science 286 (1999), p. 2345–2348 95 PAGE, M. I., JENCKS, W. P., Proc. Natl. Acad. Sci. U.S.A. 68 (1971), p. 1678–1683 96 HILVERT, D., HILL, K. W., Methods Enzymol. 203 (1991), p. 352–369 97 ARE´ VALO, J. H., STURA, E. A., TAUSSIG, M. J., WILSON, I. A., J. Mol. Biol. 231 (1993), p. 103–118

29

30 Structure and Function of Catalytic Antibodies

98 ARE´ VALO, J. H., TAUSSIG, M. J., WILSON, I. A. Nature 365 (1993), p. 859–863 99 ARE´ VALO, J. H., HASSIG, C. A., STURA, E. A., SIMS, M. J., TAUSSIG, M. J., WILSON, I. A. J. Mol. Biol. 241 (1994), p. 663–690 100 CRAVATT, B. F., ASHLEY, J. A., JANDA, K. D., BOGER, D. L., LERNER, R. A., J. Am. Chem. Soc. 116 (1994), p. 6013–6014 101 LARSEN, N. A., HEINE, A., CRANE, L., CRAVATT, B. F., LERNER, R. A., WILSON, I. A., J. Mol. Biol. 314 (2001), p. 93–102 102 MARCH, J. Advanced organic chemistry, John Wiley & Sons, New York 1992 103 STORK, G., BURGSTAHLER, A. W. J.Am. Chem. Soc. 77 (1955), p. 5068 104 ESCHENMOSER, A., RUZICKA, L., JEGER, O., ARIGONI, D. Helv. Chim. Acta 38 (1955), p. 1890 105 LESBURG, C. A., CARUTHERS, J. M., PASCHALL, C. M., CHRISTIANSON, D. W., Curr. Opin. Struct. Biol. 8 (1998), p. 695–703 106 JOHNSON, G., J.Am.Chem.Soc. 86 (1964), p. 1959 107 LI, T., JANDA, K. D., ASHLEY, J. A., LERNER, R. A., Science 264 (1994), p. 1289–1293 108 ABE, I., ROHMER, M., PRESTWICH, G. D., Chem. Rev. 93 (1993), p. 2189–2206 109 HASSERODT, J., JANDA, K. D., LERNER, R. A., J. Am. Chem. Soc. 119 (1997), p. 5993–5998 110 DAILEY, H. A., FLEMING, J. E., J.Biol. Chem. 258 (1983), p. 11453 111 COCHRAN, A. G., SCHULTZ, P. G. Science 249 (1990), p. 781–783 112 TAKETANI, S., TOKUNAGA, R., Eur.J. Biochem. 127 (1982), p. 443–447 113 ROMESBERG, F. E., SANTARSIERO, B. D., SPILLER, B., YIN, J., BARNES, D., SCHULTZ, P. G., STEVENS, R. C. Biochemistry 37 (1998), p. 14404–14409 114 WIRSCHING, P., ASHLEY, J. A., LO, C.-H. L., JANDA, K. D., LERNER, R. A., Science 270 (1995), p. 1775–1782 115 WAGNER, J., LERNER, R. A., BARBAS III, C. F., Science 270 (1995), p. 1797–1800 116 BARBAS III, C. F., HEINE, A., ZHONG, G., HOFFMANN, T., GRAMATIKOVA, S., BJORNESTEDT, R., LIST, B., ANDERSON, J., STURA, E. A., WILSON, I. A., LERNER, R. A. Science 278 (1997), p. 2085–2092

117 KARLSTROM, A., ZHONG, G., RADER, C., LARSEN, N. A., HEINE, A., FULLER, R., LIST, B., TANAKA, F., WILSON, I. A., BARBAS III, C. F., LERNER, R. A., Proc. Natl. Acad. Sci. U.S.A. 97 (2000), p. 3878–3883 118 HEINE, A., DESANTIS, G., LUZ, J. G., MITCHELL, M., WONG, C.-H., WILSON, I. A., Science 294 (2001), p. 369–374 119 ZHONG, G., LERNER, R. A., BARBAS, C. F., III. Angew. Chem. Int. Ed. Engl. submitted (1999) 120 ZHU, X., WILSON, I. A., personal communication (2003) 121 FESSNER, W.-D., Curr.Opin.Chem. Biol. 2 (1998), p. 85–97 122 CHEN, L., DUMAS, D. P., WONG, C.-H., J. Am. Chem. Soc. 114 (1992), p. 741–748 123 RACKER, E., J.Biol.Chem. 196 (1952), p. 347–365 124 BARBAS III, C. F., WANG, Y.-F., WONG, C.-H., J. Am. Chem. Soc. 112 (1990), p. 2013–2014 125 ZHONG, G., SHABAT, D., LIST, B., ANDERSON, J., SINHA, S. C., LERNER, R. A., BARBAS III, C. F., Angew. Chem. Int. Ed. 37 (1998), p. 2481–2484 126 LIST, B., Tetrahedron 58 (2002), p. 5573–5590 127 WENTWORTH, A. D., JONES, L. H., WENTWORTH Jr., P., JANDA, K. D., LERNER, R. A., Proc. Natl. Acad. Sci. U.S.A. 97 (2000), p. 10930–10935 128 WENTWORTH Jr., P., JONES, L. H., WENTWORTH, A. D., ZHU, X., LARSEN, N. A., WILSON, I. A., XU, X., GODDARD, W. A., JANDA, K. D., ESCHENMOSER, A., LERNER, R. A., Science 293 (2001), p. 1806–1811 129 DATTA, D., VAIDEHI, N., XU, X., GODDARD III, W. A., Proc. Natl. Acad. Sci. U.S.A. 99 (2002), p. 2636–2641 130 PATTEN, P. A., GRAY, N. S., YANG, P. L., MARKS, C. B., WEDEMAYER, G. J., BONIFACE, J. J., STEVENS, R. C., SCHULTZ, P. G., Science 271 (1996), p. 1086–1091. 131 FUJII, I., TANAKA, F., MIYASHITA, H., TANIMURA, R., KINOSHITA, K., J. Am. Chem. Soc. 117 (1995), p. 6199–6209 132 YANG, G., CHUN, J., ARAKAWA-URAMOTO, H., WANG, X., GAWINOWICZ, M. A., ZHAO, K., LANDRY, D. W., J. Am. Chem. Soc. 118 (1996), p. 5881–5890

References 133 LARSEN, N. A., HEINE, A., de PRADA, P., REDWAN, R. M., YEATES, T. O., LANDRY, D. W., WILSON, I. A., Acta Crystallogr. D58 (2002), p. 2055–2059 134 BENSEL, N., BAHR, N., REYMOND, M. T., SCHENKELS, C., REYMOND, J.-L., Helv. Chim. Acta 82 (1999), p. 44–52 135 ZHONG, G., SHABAT, D., LIST, B., ANDERSON, J., SINHA, S. C., LERNER, R. A., BARBAS III, C. F., Angew. Chem. Int. Ed. 37 (1998), p. 2481–2484 136 LEWIS, C., PANETH, P., O’LEARY, M. H., HILVERT, D., J. Am. Chem. Soc. 115

(1993), p. 1410–1413 137 TARASOW, T. M., LEWIS, C., HILVERT, D. J. Am. Chem. Soc. 116 (1994), p. 7959–7963 138 GRUBER, K., ZHOU, B., HOUK, K. N., LERNER, R. A., SHEVLIN, C. G., WILSON, I. A., Biochemistry 38 (1999), p. 7062–7074 139 ZHU, X., WENTWORTH, P., WENTWORTH, A. D., ESCHENMOSER, A., LERNER, R. A., WILSON, I. A., Proc. Natl. Acad. Sci. U.S.A. 101 (2004), p. 2247–2252

31

1

N-terminal Ubiquitination Aaron Ciechanover

Technion — Israel Institute of Technology, Haifa, Israel

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-7 Abstract

The ubiquitin–proteasome system (UPS) is involved in selective targeting of innumerable cellular proteins via a complex pathway that plays important roles in a broad array of processes. An important step in the proteolytic cascade is speciﬁc recognition of the substrate by one of many ubiquitin ligases, E3s, that is followed by generation of the polyubiquitin degradation signal. For most substrates, it is believed, though it has not been shown directly, that the ﬁrst ubiquitin moiety is conjugated, via its C-terminal Gly76 residue, to an ε-NH2 group of an internal lysine residue. Recent ﬁndings indicate that for an increasing number of proteins, the ﬁrst ubiquitin moiety is fused linearly to the α -NH2 group of the N-terminal residue. An important biological question relates to the evolutionary requirement for an alternative mode of ubiquitination.

1 Background

Two distinct structural elements play a role in the ubiquitination of a target protein: (i) the E3 recognition site and (ii) the anchoring residue of the polyubiquitin chain. In most cases, it is believed, though it has been shown for only a few proteins, that the ﬁrst ubiquitin moiety is transferred to an ε-NH2 group of an internal lysine residue in the substrate. The N-terminal domain of the target protein has attracted attention both as an E3 recognition domain and, recently, as a ubiquitination site. As for speciﬁc recognition, in certain rare cases, the stability of a protein is a direct function of its N-terminal residue, which serves as a binding site for the ubiquitin ligase E3α (Ubr1 in yeast; ‘N-end-rule’; [1, 2]). Accordingly, two types of N-terminal residues have been deﬁned, “stabilizing” and “destabilizing”. For the

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 N-terminal Ubiquitination

Mos protein, it was found that its stability is governed primarily by the penultimate proline residue and by a phosphorylation/dephosphorylation cycle of serine3 [3]. A mechanistic explanation for the role of the Pro and Ser residues is still missing. As for the lysine residue targeted, there is no consensus as to its speciﬁcity. In some cases distinct lysines are required, while in others there is little or no speciﬁcity. Thus signal-induced degradation of IκBα involves two particular lysine residues, 21 and 22 [4]. In the case of Gcn4, lysine residues in the vicinity of a speciﬁc PEST degradation signal serve as ubiquitin attachment sites [5]. Mapping of ubiquitination sites of the yeast iso-2-cytochrome c has revealed that the polyubiquitin chain is synthesized almost exclusively on a single lysine [6]. In two other examples, that of Mos (see above; [3]) and the model “N-end rule” substrate X-β-gal (where X is a short fused peptide not encoded by the native molecule [7]), one and two lysines, respectively, that reside in proximity to the degradation signal are required for ubiquitination. In striking contrast, ubiquitination of the ζ chain of the T-cell receptor is independent of any particular lysine residue and proceeds as long as one residue is present in the cytosolic tail of the molecule [8]. Similarly, no single speciﬁc lysine residue is required for ubiquitination of either c-Jun [9] or cyclin B [10]: any single lysine residue, even artiﬁcially inserted, can serve as a ubiquitin acceptor. Important in this context is that only in a handful of cases it has been shown directly, via chromatographic or mass spectrometric analyses, that ubiquitin is indeed anchored to a lysine residue (see for example Refs. [11, 12]). In most cases studied, and there are not too many, the assumption that an internal lysine serves as the polyubiquitin chain anchor is indirect and based on mutational analyses. One interesting case involves the artiﬁcial fusion protein ubiquitin-Pro-X-βgalactosidase. In this chimera, the ubiquitin moiety was fused to the N-terminal Pro residue of the protein. Unlike other ubiquitin-B-X-β-galactosidase species (where B is any of the remaining 19 amino acid residues), here ubiquitin is not removed by isopeptidases and serves as a degradation signal following generation of a polyubiquitin chain that is anchored to Lys48 of the artiﬁcially fused ubiquitin moiety [13]. However as noted, in this case the ubiquitin moiety was fused to the N-terminal residue artiﬁcially. The ﬁrst substrate that was identiﬁed in which the N-terminal residue serves as a ubiquitination target was MyoD. The basic helix–loop–helix (bHLH) protein MyoD is a tissue-speciﬁc transcriptional activator that acts as a master switch for muscle development. MyoD forms heterodimers with other proteins belonging to the bHLH group, such as the ubiquitously expressed E2A, E12 and E47. These dimers are probably the transcriptionally active forms of the factor. Association of MyoD with HLH proteins of the Id family (inhibitors of differentiation that lack the basic domain) inhibits its DNA-binding and biological activities. MyoD is a short-lived protein with a half-life of ∼45 min [14, 15]. Degradation of MyoD is mediated by the ubiquitin system both in vitro and in vivo. Furthermore, the process is inhibited by its consensus DNA-binding site. In contrast, addition of Id1 destabilizes the MyoD–E47–DNA complex and renders the protein susceptible to degradation [15].

2 Results

2 Results

To analyze speciﬁc ubiquitination sites in MyoD, we used site-directed mutagenesis to substitute systematically all the lysine residues with arginines [16]. The protein contains nine lysine residues, most of them located within the N-terminal domain of the molecule. The nine residues are in positions 58, 99, 102, 104, 112, 124, 133, 146 and 241. The various proteins were generated either by expression in bacteria followed by puriﬁcation, or by in vitro translation in reticulocyte lysate in the presence of [35 S]methionine. Conjugation and degradation of the proteins were monitored in a reconstituted cell-free system or in cells. Proteins were detected by either Western blot analysis or PhosphorImaging. Surprisingly, even a MyoD species that lacked all lysine residues was still degraded efﬁciently in an ATPdependent manner in vitro. To demonstrate involvement of the ubiquitin system in the process, we followed the degradation of wild-type (WT) and lysine-less (LL) MyoD in the absence and presence of ubiquitin. Similar to the degradation of the WT protein, degradation of the LL MyoD was completely dependent upon the addition of exogenous ubiquitin to an extract that does not contain it (Fraction II). Furthermore, addition of methylated ubiquitin, which cannot form polyubiquitin chains and serves as a chain terminator [17], inhibited the degradation of LL MyoD. The inhibition could be alleviated by the addition of excess of free ubiquitin. These results strongly suggested that polyubiquitination of LL MyoD is necessary for degradation of the protein. Furthermore, they implied that the polyubiquitin chain is synthesized on internal lysine residues of ubiquitin. To demonstrate directly polyubiquitinated LL MyoD, we used in-vitro-translated 35 S-labeled protein in a partially reconstituted system. We demonstrated that LL MyoD generates high molecular mass ubiquitinated adducts. It should be noted, however, that these conjugates are of somewhat lower molecular mass than those of the WT MyoD. This can be attributed to the role that the internal lysine residues also play in the process (see also below). To investigate the physiological relevance of the observations in the cell-free system, we followed the fate of the different MyoD lysine-mutated proteins in vivo, using pulse-chase labeling experiments in COS-7 cells that were transiently transfected with the different MyoD cDNAs. In agreement with our in vitro data, the lysine-less MyoD protein is degraded efﬁciently in cells as well. However, we could observe a progressive increase in the half-life of the proteins of up to ∼2-fold with the gradual substitution of the lysine residues. While the half-life of WT MyoD was ∼50 min, that of LL MyoD was ∼2 h. Interestingly, we found that the stability of MyoD is not affected by the substitution of any speciﬁc lysine residue, and it is the total number of these residues that determines the half-life of the protein. To identify the system involved in the destruction of LL MyoD in vivo, transfected cells were incubated in the presence of inhibitors of proteasomal and lysosomal degradation. Chloroquine, a general inhibitor of lysosomal proteolysis, and E-64, a cysteine protease inhibitor that affects lysosomal, but also certain cytosolic proteases, had no effect on the stability of the LL MyoD. In striking contrast, the proteasomal

3

4 N-terminal Ubiquitination

inhibitors MG132 and lactacystin blocked degradation of the LL protein signiﬁcantly. To demonstrate the intermediacy of ubiquitin conjugates in the degradation of LL MyoD, we incubated COS-7 cells, transiently transfected with either WT or LL MyoD cDNAs, with MG132, and followed generation of ubiquitin-MyoD adducts. Immunoprecipitation with anti-MyoD antibody followed by Western blot analysis with anti-ubiquitin antibody revealed accumulation of high molecular mass compounds in cells transfected with either WT or LL MyoD. A similar analysis of mock-transfected cells clearly demonstrated the speciﬁcity of both the anti-MyoD and anti-ubiquitin antibodies. Based on these results, it was clear that polyubiquitination is essential for targeting MyoD for degradation. The lack of internal lysine residues, the only known targets for ubiquitin modiﬁcation, made it important to identify the functional group that can serve as an attachment site for ubiquitin. Chemically, several groups can generate covalent bonds with ubiquitin. Ser and Thr can participate in ester bond formation, while Cys can generate a thiol ester bond. However, these bonds are unstable and are hydrolyzed in either high pH (Ser and Thr) or high concentration of -SH groups (Cys). The stability of the MyoD-ubiquitin adducts under these conditions made it highly unlikely that any of these modiﬁcations is the one we observed. A likely candidate, however, was the free amino group of the N-terminal residue of the protein, which can generate a stable peptide bond with the C-terminal Gly residue of ubiquitin. Edman degradation of the N-terminal residue of bacterially expressed, in-vitro-translated and cellularly expressed MyoDs, has revealed that the ubiquitin attachment site can be the free, unmodiﬁed initiator methionine: the proteins were not modiﬁed and the N-terminal residue was not acetylated. To demonstrate a role for the free N-terminal amino group in the degradation of MyoD, we chemically modiﬁed this group. Initially, we blocked this group in the LL MyoD protein by reductive methylation. While this procedure blocks all amino groups in a protein in a non-discriminatory manner, in this case, it could have been only the α-NH2 group, which is the only free amino group left in the MyoD molecule. The modiﬁcation stabilized the protein completely. Whereas a free α-NH2 appears to be sufﬁcient for degradation (probably following ubiquitination) of LL MyoD, it is not clear whether it also plays a physiological role in targeting the WT molecule, which has nine available lysine residues. In order to investigate the role and biological relevance of the free α-NH2 group in the targeting of WT MyoD, we selectively blocked it by carbamoylation with potassium cyanate at low pH. This procedure does not modify ε-NH2 groups of internal lysine residues. Automated Edman degradation along with fuorescamine determination of the extent of remaining free NH2 groups conﬁrmed that the modiﬁcation affected only the N-terminal group. The modiﬁed protein was subjected to in vitro degradation and conjugation in cell extract. In contrast to LL MyoD, the N-terminally carbamoylated protein could not be ubiquitinated and was stable. Thus, a free and exposed NH2 terminus of MyoD appears to be an essential site for degradation, most probably because it serves as an attachment site for the ﬁrst ubiquitin moiety. As an additional control, we selectively modiﬁed the internal lysine residues of WT MyoD by guanidination

2 Results

with O-methylisourea. The modiﬁcation, which does not affect the N-terminal group, generates a protein that is essentially the chemically modiﬁed counterpart of the LL MyoD that was generated by site-directed mutagenesis. Similar to the LL protein, this MyoD derivative is degraded efﬁciently in the cell-free system in a ubiquitin-and an ATP-dependent mode. To analyze the role of the N-terminal residue of MyoD as a ubiquitination site, we fused to WT MyoD, upstream to the N-terminal residue, a 6 × Myc tag, and monitored the stability of the tagged protein. We showed that it is stable both in vitro and in vivo. It should be noted that the two ﬁrst N-terminal residues of the Myc tag, methionine and glutamate, are identical to the ﬁrst two N-terminal residues in MyoD. In addition, the Myc tag also contains a lysine residue. Thus, altogether, six additional lysine residues were added to WT MyoD in addition to its own nine native residues. Nevertheless, the tag stabilizes it, probably by blocking access to a speciﬁc N-terminal residue, and, as became clear later (see below), to its neighboring domain. Taken together, these ﬁndings strongly suggested that MyoD is ﬁrst ubiquitinated at its N-terminal residue, and the polyubiquitin chain is synthesized on this ﬁrst conjugated ubiquitin moiety. Internal lysine residues also play a role, probably by serving as additional anchoring sites, whose ubiquitination accelerates degradation. Yet they are not essential for proteolysis to occur. In contrast, ubiquitination of the N-terminal residue plays a critical role in governing the protein’s stability [16] (Figure 1). Using a similar, though not a complete, set of experiments, 12 additional proteins have been identiﬁed recently that appear to undergo N-terminal ubiquitination: (i) the human papilloma virus-16 (HPV-16) E7 oncoprotein (18), (ii) the latent membrane protein 1 (LMP1)(19) and (iii) 2A (LMP2A)(20) of the Epstein Barr virus (EBV), involved in viral activation from latency, (iv) the cell cycle-dependent kinase (CDK) inhibitor p21 (21,22), (v) the extracellular signal-regulated kinase 3, ERK3 (22), the inhibitors of differentiation (vi) Id2 (23) and (vii) Id1 (24), two pro-proliferative Helix-Loop-Helix proteins, (viii) hydroxymethyglutaryl-Coenzyme A reductase (HMG-CoA reductase), the ﬁrst and key regulatory enzyme in the cholesterol biosynthetic pathway (25), (ix) p19ARF , the mouse Mdm2 inhibitor and (x) p14ARF , its human homologue (26), (xi) the HPV-58 E7 oncoprotein, and (xii) the cell cycle regulator p16INK4a (27). As for HMG-CoA reductase, several speciﬁc internal lysine residues have also been shown to be important for its targeting, and therefore the essentiality of the N-terminal residue in the process has to be further substantiated. The case of p21 requires further investigation, as one study reported that its degradation by the proteasome does not require ubiquitination [28], while an independent study has demonstrated a role for Mdm2 in targeting p21, also without a requirement for ubiquitination [29]. As we noted for MyoD, substitution of the internal lysines inhibited slightly (up to twofold) both conjugation and degradation of HPV-16, LMP1, and Id2, suggesting that these residues, probably also by serving as ubiquitin anchors, can modulate the stability of these proteins. It is possible to suppose that N-terminal ubiquitination and modiﬁcation of internal lysines is catalyzed by different ligases that may be even located in different subcellular

5

6 N-terminal Ubiquitination

NH

A

Lys48 - based polyubiquitin chain

K48 G76

C=O NH Ubiquitin K48 G76 C=O NH NH2

X

Kn

COOH

Protein target

B

NH

Lys48 – based polyubiquitin chain

K48 G76

C=O NH NH2

K48 Ubiquitin G76

C NH O

X

Fig. 1 Ubiquitination on an internal lysine and on the N-terminal residue of the target substrate. (A) The ﬁrst ubiquitin moiety is conjugated, via its C-terminal Gly76 residue, to the g-NH2 group of an internal lysine residue of the target substrate (Kn ). (B) The ﬁrst ubiquitin moiety is conjugated to a free

Kn

COOH

Protein target "-NH2 group of the N-terminal residue, X. In both cases, successive addition of activated ubiquitin moieties to internal Lys48 on the previously conjugated ubiquitin moiety leads to the synthesis of a polyubiquitin chain which serves as the degradation signal for the 26S proteasome.

compartments (e.g. the nucleus and cytosol). Because of the role that internal lysines play in modulating the stability of these proteins, and in order to better understand the physiological signiﬁcance of this novel mode of modiﬁcation, it was important to identify proteins whose degradation is completely dependent on N-terminal ubiquitination. An important group of potential substrates for Nterminal ubiquitination is that of naturally occurring lysine-less proteins – NOLLPs. Since these proteins cannot use the “canonical” lysine conjugation pathway, in order to be targeted by the ubiquitin system they must use, an alternative site for their tagging. Searching the database, we were able to identify 177 eukaryotic NOLLPs, 14 of which occur in humans. In addition, we have identiﬁed 111 viral NOLLPs. We have shown that two of the proteins mentioned above, the human tumor supressor p16INK4a and the viral oncoprotein HPV-58 E7 are degraded via

2 Results

the N-terminal ubiquitination pathway [27]. Interestingly, we demonstrated that p16INK4a is ubiquitinated and degraded only in sparse cells, and is stable in dense cells. Similar ﬁndings were reported for the NOLLPs p19ARF and p14ARF (26). For E7-16 [18], LMP1 [19], and MyoD (unpublished), it has been shown that truncation of a short N-terminal segment of 10–20 residues stabilized the proteins, suggesting that the entire domain beyond the single N-terminal residue plays a role in governing the stability of these proteins. Such a segment can allow the mobility/ﬂexibility necessary for the N-terminal residue to serve as a ubiquitin acceptor. It can also serve as a recognition domain for the cognate E3. There is no homology between the N-terminal domains of these three proteins, suggesting that if the three N-terminal domains serve as recognition motifs, they recognize different components of the ubiquitin system. Interestingly, the LMP2A E3 was identiﬁed as a member of the NEDD4 family of HECT domain ligases, AIP4 and/or WWP2 [20]. A PY motif in LMP2A is recognized by the E3. It resides in the Nterminal domain of the molecule, supporting the hypothesis that in these proteins the E3-binding domain may reside in close proximity to the N-terminal residue ubiquitination site. Is there any direct evidence for N-terminal ubiquitination? All the different and independent lines of evidence in the various studies strongly suggest that ubiquiti-nation occurs on the N-terminal residue, and any other scenario is highly unlikely. Yet, the only direct evidence must be demonstration of a fusion peptide between the C-terminal domain of ubiquitin and the N-terminal domain of the target substrate. The study on p21 [21] and E7-58 [27] brought us a little closer. Bloom and colleagues [21] transfected cells with N-terminally His-tagged ubiquitin and N-terminally HA-tagged p21 that contained a Factor X proteolytic site immediately after the HA tag and upstream of the p21 reading frame. They then immunoprecipitated and resolved the cell-generated ubiquitin conjugates of p21 and treated the mono-ubiquitin–p21 adduct with Factor-X protease. This treatment released a smaller species of p21 (lacking His-ubiquitin and the HA-tag-Factor X site) and His-ubiquitin-HA-Factor X site, thus demonstrating that the HA-tag-Factor X site, which was previously part of p21, had now become part of the Factor X-cleaved ubiquitin. A similar experimental evidence was brought for the NOLLP HPV E7-58 [27]. Here, Ben-Saadon and colleagues generated two species of the protein containing the eight amino acid sequence of the Tobacco Etch Virus (TEV) protease cleavage site inserted either 21 amino acid residues after the iMet [E7-58-TEV(21)] or immediately after the iMet [E7-58-TEV(1)]. The prediction from this experiment was that if ubiquitin is indeed attached to the N-terminal residue of E7-58, TEV protease-catalyzed cleavage will generate an extended ubiquitin molecule that will also contain the respective N-terminal domain of E7-58 [21 residues or 1 residue, respectively, dependent upon whether the substrate of the reaction is E7-58-TEV(21) or E7-58-TEV(1), and the six amino acids derived from the TEV cleavage site]. Such extended ubiquitin moieties were indeed generated following incubation of the substrate with labeled methylated ubiquitin (which generated mostly the mono-ubiquitin adduct of the E7-58 protein), followed by cleavage of the adduct with TEV. The only conclusion that can be derived from these experiments

7

8 N-terminal Ubiquitination

is that the ubiquitin moiety was fused to any of the amino acid residues of the HA tag-Factor X site at the N-terminal domain of p21, or to any of the ﬁrst 21 amino acids of E7 or the TEV site (part of it; the protease cleaves after the sixth amino acids out of eight in the complete site). Such internal modiﬁcation is unlikely, however, as it must require a novel chemistry since none of the residues in the tags, the protease sites or the E7 N-terminal fragment, is lysine. Yet, formally, it is still possible that such a modiﬁcation occurs. The HA tag contains, for example, three Tyr residues. Thus, the evidence provided by these two experiments clearly limits an unlikely non-peptide bond ubiquitination, such as esteriﬁcation, to a much smaller zone in the N-terminal domain of the tagged p21 or the TEV-containing E7-58, but does not demonstrate directly that the modiﬁcation occurs indeed on the N-terminal residue. As noted, only identiﬁcation of a fusion peptide between the C-terminal domain of ubiquitin and the N-terminal domain of the target protein will constitute such an evidence. Ben-Saadon and colleagues have recently isolated the long sought after fusion peptide [27]: mass spectrometric analysis of a tryptic digest of the isolated mono-ubiquitin adduct of HPV-58 E7 revealed a peptide of 11 amino acids, GG-MHGNNPTLR which represents the last two C-terminal amino acids of ubiquitin, GlyGly, and the ﬁrst nine residues of E7, MetHisGlyAsnAsnProThrLeuArg (MHGNNPTLR). It should be noted that WT E7-58 contains an Arg residue in position 2. It was necessary to substitute this Arg with His, since otherwise the digesting enzyme, trypsin, would have generated a tetrapeptide, GG-MR, that would have been difﬁcult, if not impossible, to identify in the MS analysis. MS/MS analysis of the 11-mer, veriﬁed its internal sequence. Coulombe and colleagues were also able to isolate and identify the sequence of a fusion peptide between the C-terminal domain of ubiquitin and the N-terminal domain of HA-tagged p21 that also contained, downstream of the tag, a stretch of residues derived from the N-terminal domain of the native substrate, but without the iMet (which was removed during the construction of the tagged protein) [22].

3 Discussion

N-terminal ubiquitination is a novel pathway, clearly distinct from the N-end rule pathway [30]. In the latter, the N-terminal residue serves as a recognition and binding motif to the ubiquitin ligase, E3α; however, ubiquitination occurs on an internal lysine(s). In contrast, in the N-terminal ubiquitination pathway, modiﬁcation occurs on the N-terminal residue, whereas recognition probably involves a downstream motif. It should be mentioned that in yeast, using the model fusion protein ubiquitin-Pro-X-β-galactosidase (where X is a short sequence derived from the λ repressor), a new proteolytic pathway has been described, designated the UFD (ubiquitin fusion degradation) pathway [31]. The stably fused ubiquitin moiety (note that in this exceptional case, with Pro and not any other amino acid residue as the linking residue, the ubiquitin moiety is not cleaved off by ubiquitin C-terminal

3 Discussion

hydrolases), functions as a degradation signal, where its Lys48 serves as an anchor for the synthesized polyubiquitin chain. This pathway involves several enzymes, UFD 1–5, some of which appear to be unique and are not part of the “canonical” UPS. It is possible that N-terminal ubiquitination is the most upstream event in the UFD pathway – which was discovered using an artiﬁcial chimeric ubiquitin–protein model substrate: the N-terminal ubiquitination pathway can function by providing substrates to the UFD pathway. The physiological signiﬁcance of N-terminal ubiquitination is still obscure. Naturally occurring lysine-less proteins, NOLLPs, that are degraded by the ubiquitin system must traverse this pathway. Many such proteins, mostly viral, can be found in the database (see above and in Refs. [26, 27]). We believe that many additional lysine-containing proteins, will be discovered to be targeted via this novel mode of modiﬁcation. Of note is that all the proteins that are N-terminally ubiquitinated must contain a free, unmodiﬁed N-terminal residue. Such proteins constitute approximately 25% of all cellular proteins, while the remaining 75% are Nα-acetylated. Whether a protein will be acetylated is dependent on the structure of the N-terminal domain of the mature protein. This is determined by the combined activities of methionine aminopeptidases (MAPs) and N-terminal acetyltransferases (NATs), which are dependent on the speciﬁc sequence of up to the ﬁrst four N-terminal residues of the target protein substrates (reviewed in Ref. [32]). Thus, it is possible to predict which proteins will be potential substrates of the N-terminal ubiquitination pathway. Internal C-terminal fragments of Nα-acetylated proteins can also be modiﬁed by ubiquitin at their “new” N-terminal residue following limited processing. Many proteins, such as the NF-κB precursors p105 and p100 or caspase substrates are processed initially in a limited manner, generating a C-terminal fragment with a newly exposed N-terminal residue. For all lysine-containing proteins, the intact free N-termini as well as the products of processing, the assumption is that their internal lysines are not easily accessible, for whatever reason, for ubiquitination, and it is only the N-terminal residue that can be modiﬁed. Interestingly, most of the substrates identiﬁed thus far have a few lysine residues that might not be accessible to the E3s: For example, MyoD has nine (out of 319), E7 two (out of 97), LMP1 has a single lysine residue (out of 440), LMP2A has three (out of 497), Id2 nine (out of 134) and p21 six (out of 164). From the random discovery of thirteen N-terminally ubiquitinated proteins, it appears that their number could well be larger and that many more will be discovered, which will help in the unraveling of the unique characteristics that distinguish this group of substrates.

Acknowledgments

Research in the laboratory of A.C. is supported by grants from Prostate Cancer Foundation Israel - Centers of Excellence Program, the Israel Science Foundation – Centers of Excellence Program, a Professorship funded by the Israel Cancer Research Fund (ICRF) and the Foundation for Promotion of Research in the Technion administered by the Vice President of the Technion for Research. Infrastructural

9

10 N-terminal Ubiquitination

equipment was purchased with the support of the Wolfson Charitable Fund Center of Excellence for studies on Turnover of Cellular Proteins and its Implications to Human Diseases.

References 1 BACHMAIR, A., FINLEY, D. and VARSHAVSKY, A. In vivo half-life of a protein is a function of its amino-terminal residue. Science, 1986, 234, 179–186. 2 VARSHAVSKY, A. The N-end rule: functions, mysteries, uses. Proc. Natl Acad. Sci. USA, 1996, 93, 12142–12149. 3 NISHIZAWA, M., FURUNO, N., OKAZAKI, K., TANAKA, H., OGAWA, Y. and SAGATA, N. Degradation of Mos by the N-terminal proline (Pro2)-dependent ubiquitin pathway on fertilization of Xenopus eggs: possible signiﬁcance of natural selection for Pro2 in Mos. EMBO J., 1993, 12, 4021–4027. 4 SCHERER, D. C., BROCKMAN, J. A., CHEN, Z., MANIATIS, T. and BALLARD, D. W. Signal-induced degradation of IkBα requires site-speciﬁc ubiquitination. Proc. Natl Acad. Sci. USA, 1995, 92, 11259–11263. 5 KORNITZER, D., RABOY, B., KULKA, R. G. and FINK, G. R. Regulated degradation of the transcription factor Gcn4. EMBO J., 1994, 13, 6021–6030. 6 SOKOLIK, C. W. and COHEN, R. E. The structures of ubiquitin conjugates of yeast Iso-2-cytochrome c. J. Biol. Chem., 1991, 266, 9100–9107. 7 CHAU, V., TOBIAS, J. W., BACHMAIR, A., MARRIOTT, D., ECKER, D. J., GONDA, D. K. and VARSHAVSKY, A. A multi ubiquitin chain is conﬁned to speciﬁc lysine in a targeted short-lived protein. Science, 1989, 243, 1576–1583. 8 HOU, D., CENCIARELLI, C., JENSEN, J. P., NGUYGEN, H. B. and WEISSMAN, A. M. Activation-dependent ubiquitination of a T cell antigen receptor subunit on multiple intracellular lysines. J. Biol. Chem., 1994, 269, 14244–14247. 9 TREIER, M., STASZEWSKI, L. and BOHMANN, D. Ubiquitin-dependent c-Jun degradation in vivo is mediated by the δ-domain. Cell, 1994, 78, 787–798.

10 KING, R. W., GLOTZER, M. and KIRSCHNER, M. W. Mutagenic analysis of the destruction signal of mitotic cyclins and structural characterization of ubiquitinated intermediates. Mol. Biol. Cell, 1996, 7, 1343–1357. 11 GOLDKNOPF, I. L. and BUSCH, H. Isopeptide linkage between nonhistone and histone 2A polypeptides of chromosomal conjugate-protein A24. Proc. Natl. Acad. USA, 1977, 74, 864–868. 12 GRONROOS, E., HELLMAN, U., HELDIN, C. H. and ERICSSON, J. Control of Smad7 stability by competition between acetylation and ubiquitination. Mol. Cell, 2002, 10, 483–493. 13 JOHNSON, E. S., BARTEL, B., SEUFERT, W. and VARSHAVSKY, A. Ubiquitin as a degradation signal. EMBO J. 1992, 11, 497–505. 14 THAYER, M. J., TAPSCOTT, S. J., DAVIS, R. L., WRIGHT, W. E., LASSAR, A. B. and WEINTRAUB, H. Positive autoregulation of the myogenic determination gene MyoD1. Cell, 1989, 58, 241–248. 15 ABU-HATOUM, O., GROSS-MESILATY, S., BREITSCHOPF, K., HOFFMAN, A., GONEN, H., CIECHANOVER, A. and BENGAL, E. Degradation of the myogenic transcription factor MyoD by the ubiquitin pathway in vivo and in vitro: regulation by speciﬁc DNA binding. Mol. Cell. Biol., 1998, 18, 5670–5677. 16 BREITSCHOPF, K., BENGAL, E., ZIV, T., ADMON, A. and CIECHANOVER, A. A novel site for ubiquitination: The N-terminal residue and not internal lysines of MyoD is essential for conjugation and degradation of the protein. EMBO J., 1998, 17, 5964–5973. 17 HERSHKO, A. and HELLER, H. Occurrence of a polyubiquitin structure in ubiquitin-protein conjugates. Biochem. Biophys. Res. Commun. 1985, 128, 1079–1086.

References 18 REINSTEIN, E., SCHEFFNER, M., OREN, M., SCHWARTZ, A. L. and CIECHANOVER, A. Degradation of the E7 human papillomavirus oncoprotein by the ubiquitin-proteasome system: 2 N-terminal Ubiquitination: No Longer Such a Rare Modiﬁcation targeting via ubiquitination of the N-terminal residue. Oncogene, 2000, 19, 5944–5950. 19 AVIEL, S., WINBERG, G., MASSUCCI, M. and CIECHANOVER, A. Degradation of the Epstein-Barr virus latent membrane protein 1 (LMP1) by the ubiquitin-proteasome pathway: targeting via ubiquitination of the N-terminal residue. J. Biol Chem., 2000, 275, 23491–23499. 20 IKEDA, M., IKEDA, A. and LONGNECKER, R. Lysine-independent ubiquitination of the Epstein-Barr virus LMP2A. Virology, 2002, 300, 153–159. 21 BLOOM, J., AMADOR, V., BARTOLINI, F., DEMARTINO, G. and PAGANO, M. Proteasome-mediated degradation of p21 via N-terminal ubiquitinylation. Cell, 2003, 115, 1–20. 22 COULOMBE, P., RODIER, G., BONNEIL, E., THIBAULT, P. and MELOCHE, S. N-Terminal ubiquitination of extracellular signal-regulated kinase 3 and p21 directs their degradation by the proteasome. Mol. Cell Biol., 2004, 24, 6140–6150. 23 FAJERMAN, I., SCHAWARTZ, A. L. and CIECHANOVER, A. Degradation of the Id2 developmental regulator: Targeting via N-Terminal Ubiquitination. Biochem. Biophys. Res. Commun., 2004, 314, 505–512. 24 AZAR-TRAUSCH, J. S., LINGBECK, J., CIECHANOVER, A. and SCHWARTZ, A. L. Ubiquitin-proteasome-mediated degradation of Id1 is modulated by MyoD. J. Biol. Chem., 2004, 279, 32614–32619. 25 DOOLMAN, R., LEICHNER, G. S., AVNER, R. and ROITELMAN, J. Ubiquitin is conjugated by membrane ubiquitin ligase to three

26

27

28

29

30

31

32

sites, including N-terminus, in transmembrane region of mammalian 3-hydroxy-3- methylglutaryl coenzyme a reductase: Implications for sterol-regulated enzyme degradation. J. Biol. Chem., 2004, 279, 38184–38193. KUO, M. L., DEN BESTEN, W., BERTWISTLE, D., ROUSSEL, M. F., and SHERR, C. J. N-terminal polyubiquitination and degradation of the Arf tumor suppressor. Genes and Dev., 2004, 18, 1862–1874. BEN-SAADON, R., FAJERMAN, I., ZIV, T., HELLMAN, U., SCHWARTZ, A. L., and CIECHANOVER, A. The Tumor Suppressor Protein p16INK4a and the Human Papillomavirus oncoprotein E7-58 are Naturally Occurring Lysine-Less Proteins that are Degraded by the Ubiquitin System: Direct Evidence for Ubiquitination at the N-Terminal Residue. J. Biol. Chem., 2004, 279, 41414–41421. SHEAFF, R. J., SINGER, J. D., SWANGER, J., SMITHERMAN, M., ROBERTS, J. M. and CLURMAN, B. E. Proteasomal turnover of p21Cip1 does not require p21Cip1 ubiquitination. Mol. Cell, 2000, 5, 403–410. JIN, Y., LEE, H., ZENG, S. X., DAI, M. S. and LU, H. Mdm2 promotes p21waf cip1 proteasomal turnover independently of ubiquitylation. EMBO J., 2003, 22, 6365–6377. VARSHAVSKY, A. The N-end rule: Functions, mysteries, uses. Proc. Natl. Acad. Sci. USA, 1996, 93, 12142–12149. JOHNSON, E. S., MA, P. C., OTA, I. M. and VARSHAVSKY, A. A proteolytic pathway that recognizes ubiquitin as a degradation signal. J. Biol. Chem., 1995, 270, 17442–17456. POLEVODA, B. and SHERMAN, F. N-terminal acetyltransferases and sequence requirements for N-terminal acetylation of eukaryotic proteins. J. Mol. Biol., 2003, 325, 595–622.

11

1

The COP9 Signalosome and Its Role in the Ubiquitin System Dawadschargal Bech-Otschir

Western General Hospital, Edinburgh, United Kingdom

Barbara Kapelari Max-Planck-Institute for Biochemistry, Martinsried, Germany

Wolfgang Dubiel Charit´e-Universit¨atsmedizin Berlin, Berlin, Germany

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8

1 Introduction

The COP9 signalosome (CSN) is a multimeric, highly-conserved protein complex [1]. Just like the ubiquitin system it occurs in all studied eukaryotic cells. Following its 1994 discovery in plant cells the complex was postulated to function in signal transduction [2]. Originally described as a regulator of light-dependent growth in plants [3, 4], identiﬁcation and characterization of the CSN from mammalian cells led to the discovery of sequence homologies between CSN subunits and subunits of the 26S proteasome lid complex [5, 6] as well as subunits of the translation-initiation complex eIF3 [7]. Signiﬁcant progress has been made towards understanding its structure and function by analyzing different eukaryotic organisms. The complex is involved in developmental processes of plants [8] and Drosophila [9] and is essential for embryogenesis in mice [10]. It seems to participate in processes such as DNA repair [11], cell-cycle regulation [12] and angiogenesis [13]. At the moment the pleiotropic effects of the CSN can be explained by its regulatory impact on the ubiquitin system. Here we provide a summary of current knowledge of CSN function in the ubiquitin system.

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 The COP9 Signalosome and Its Role in the Ubiquitin System

2 Discovery of the CSN

Deng and co-workers discovered the CSN in Arabidopsis thaliana when they characterized mutants of light-dependent development, and they called it the COP9 complex [2]. Morphogenesis of germinating seedlings is light-dependent. Light triggers a developmental process called photomorphogenesis. A number of mutations in the Arabidopsis COP/DET/FUS loci result in the loss of the COP9 complex accompanied with cop phenotypes in which germinating seedlings exhibit light-independent expression of light-induced genes [3]. Therefore the complex was originally hypothesized to be a repressor of photomorphogenesis [14]. The mammalian CSN complex was independently isolated during preparations of the 26S proteasome and called the JAB1-containing signalosome [15]. The same studies identiﬁed proteins such as JAB1 [15] and TRIP15 [16] as components of the complex and revealed homologies between subunits of the CSN and components of the 26S proteasome lid complex. Puriﬁcation and analysis of the complex from Arabidopsis, pork spleen [6, 17] and human red blood cells [5, 18] led to the conclusion that each subunit of the CSN has its paralog subunit in the 26S proteasome lid complex. These data suggested a common origin for the two complexes during evolution. Because they have similar architectures, the two complexes have been postulated to perform related functions (see below). Unfortunately there is only limited information on the structure or function of the eIF3 complex, and its relationship to the CSN and the lid is not well understood [7]. Studies have revealed that the CSN possesses both intrinsic and extrinsic (associated) activities, which will be reviewed in detail below. Historical gene names of the CSN have been summarized before [1]. In this article we use the uniﬁed nomenclature of the CSN [1].

3 Architecture of the CSN 3.1 CSN Subunit–Subunit Interactions

The CSN is composed of eight subunits called CSN1 to CSN8, which are highly conserved in eukaryotes, although only six of them occur in ﬁssion yeast. Two hybrid screens and biochemical methods such as far westerns, pull downs and co-precipitation deﬁned a number of CSN subunit–subunit interactions. Figure 1 illustrates known subunit–subunit interactions. Initial insight into the architecture of CSN came from the ﬁrst 2D electron microscopic analysis of puriﬁed CSN from human red blood cells [19] (see also Figure 2 below). The CSN architecture shows similarity to that of the lid. Both complexes have an asymmetric arrangement of their subunits and exhibit a central groove structure [19]. Whether the structural similarity of the two complexes is connected with

3 Architecture of the CSN 3

Fig. 1 Subunit–subunit interactions of the CSN and interactions of CSN subunits with other proteins. Subunits are numbered according to the uniﬁed nomenclature [1]. CSN subunit–subunit interactions have been published before [19]. Darker shad-

ing indicates subunits with MPN domains and lighter those with PCI domains. Known phosphorylated subunits are indicated. Details on CSN subunit interactions with other proteins can be found in the text.

similar functions remains unclear. The exact arrangement of CSN and lid subunits within their complexes remains uncertain in the absence of high-resolution crystal structures for the two complexes. Interestingly, the occurrence of smaller CSN sub-complexes apart from the large 500-kDa CSN complex has been described in different species such as Arabidopsis, Drosophila, Schizosaccharomyces pombe and mammalian cells (for a review see Ref. [20]). At the moment the physiological function of CSN sub-complexes is unclear. It can be speculated that a controlled equilibrium exists between the large and small CSN complexes. The small complexes may have a function in shuttling between nucleus and cytoplasma and/or between large multi-subunit complexes such as the 26S proteasome and cullin-based Ub ligases. 3.2 CSN-subunit Interactions With Other Proteins

Apart from subunit–subunit interactions within the CSN, a considerable number of cellular proteins interact with CSN subunits (see Figure 1). Although the

4 The COP9 Signalosome and Its Role in the Ubiquitin System

physiological relevance of many of the identiﬁed interactions is questionable, most of them might be attributed to a role of the CSN complex in signal transduction and ubiquitin-dependent proteolysis. CSN1 formerly called Gps1 was ﬁrst described as a signal transduction repressor in Arabidopsis [21]. Over-expression of CSN1 suppresses the activated JNK signaling pathway and also inhibits UV- and serum-induced c-fos expression as well as MEKKactivated AP1-activity in mammalian cells [21, 22]. It remains unclear whether overexpressed CSN1 plays a role as dominant negative regulator when in the CSN complex. Whereas the N-terminal region of CSN1 is sufﬁcient for repression, the C-terminal region is necessary for its integration into the complex and for the stability of the CSN complex [22]. Curiously, the N-terminal region of CSN1 may be not required for the CSN-associated deneddylation of cullin 1 (CUL1) and cullin 3 (CUL3), components of cullin-based E3 Ub ligases in Arabidopsis, although it appears to be one of the binding sites of the CSN for cullin-based complexes [23]. Moreover, CSN1 is the receptor site for the interaction of the CSN with inositol 1,3,4-trisphosphate 5/6 kinase [24]. In addition, CSN1 represents the interactor for a subunit of the translation-initiation factor 3, eIF3c/NIP1 [25], and for the 26S proteasome non-ATPase subunit Rpn6 [26]. Possible functions of these interactions are discussed later. CSN2 also known as alien [27] is perhaps an important regulatory subunit of the CSN. Firstly, CSN2 was identiﬁed as Trip15 (thyroid hormone receptor-interacting protein) using a yeast-two-hybrid screen [16]. The binding site of CUL1 and CUL2 is located at the N-terminal region of CSN2. This interaction is important for the CSNmediated deneddylation of cullin-based complexes that regulate their Ub-ligase activity [28]. Additionally, CSN2 binds to the transcription factor, ICSBP (interferon consensus sequence binding protein), which modulates interferon-directed gene expression [29]. Moreover it interacts with the nuclear receptors DAX-1, COUP1TF1 and ecdysone receptor [27, 30]. Interestingly, CSN2 is phos-phorylated by the CSN-associated kinases CK2 and PKD [19, 31]. However, the phosphorylation sites and their physiological function remain unclear. The CSN3 subunit interacts with IKKγ , a component of the IκB-kinase complex controlling NF-κB activity [32]. Additionally, it is the binding site for the CSNassociated kinases CK2 and PKD [31]. The subunit of the translation-initiation factor 3 complex, Int6/eIF3e, and the ubiquitin-conjugating enzyme variant, COP10, have been identiﬁed as other cellular interactors [33, 34]. Also the HIV-1 Tat protein interacts with CSN3 (our unpublished data). CSN4 is a poorly studied subunit of the CSN. Only one interactor of CSN4 has been identiﬁed, the ubiquitin-conjugating enzyme COP10 [34]. CSN5 appears to be a most important subunit both in terms of interactions with other cellular proteins and because it is a component with intrinsic metalloprotease activity (see below). The binding of CSN5 to cellular proteins including the transcription factors p53 [35] and c-Jun [15], the cell-cycle regulator protein p27 [36], rLHR (lutropin/choriogonadotropin receptor precursor) [37], Smad4 (TGF-β signaling pathway common effector) [38] and HIF1α (hypoxia-inducible factor 1) [39] appears to regulate their metabolic stability. In many cases the CSN5-interacting

3 Architecture of the CSN 5

proteins are phosphorylated by the CSN-associated kinases, which determines the speed of their destruction [40]. In contrast, Id1 and Id3 binding to the CSN complex via CSN5 leads to their stabilization, not to their phosphorylation [41]. The interaction of CSN5 to the member of the IκB multigene family Bcl3, the progesterone receptor PR, and the steroid receptor co-activator SRC-1, leads to stabilization of Bcl3-p50 and PR-SRC-1 complexes and enhances transcriptional activity [42, 43]. Whereas AP-1 activity is stimulated by interaction of CSN5 with the integrin adhesion receptor LFA-1 [44], the opposite effect was reported in the case of the cytokine migration inhibitor factor, MIF [45]. Additionally, there are other published interactors of CSN5 including the membrane-associated RING-ﬁnger Ub ligase TRC8 [46], hepatopoietin (HPO) [47], germ-line RNA helicases (GLHs) [48], and the ubiquitin C-terminal hydrolase PGP9.5 [49]. However, the exact role of these interactions remains unclear. Several groups reported the occurrence of a free CSN5 subunit [50] or CSN5 as a component of a smaller complex [51], although the exact physiological function of the different CSN5 forms is so far unclear. It is also unknown whether the occurrence of the different CSN5 forms is regulated. Moreover, little is known about CSN5 interactions in vivo, how they are regulated and under what circumstances they take place. CSN6 like CSN8 exists in eukaryotes except in S. pombe [52]. There are only a few published interactions of CSN6 with other cellular proteins. It binds to the HIV-1 Vpr protein affecting cell-cycle-associated signaling [53] and to the RING-ﬁnger protein of the SCF-complex, Rbx1 [54, 55]. In addition, CSN6 is another binding site for Int-6/eIF3e [33]. Interestingly, in mammalian cells two homologs of CSN7, CSN7a and CSN7b, have been found [6]. S. pombe contains only one form of CSN7 whereas Arabidopsis contains two alternative splicing variants, CSN7i and CSN7ii [52, 56]. CSN7 interacts with the polyamine-modulated factor PMF-1 [57]. Interestingly, CSN7 also binds the protein kinase CK2, one of the CSN-associated kinases, which phosphorylates CSN7 [31]. Whether the phosphorylated form of CSN7 is necessary for CSN complex assembly or for other regulatory events is unclear. Little is known of CSN8 interactions. CSN8 like CSN3 and CSN4 binds to COP10 [34]. 3.3 PCI and MPN

Six of the CSN subunits contain PCI (proteasome, COP9 signalosome, initiation factor 3) domains and two contain MPN (Mpr-Pad1-N-terminal) domains [58]. These two characteristic domains have been found in three protein complexes: the CSN, the 26S proteasome lid complex (lid) and the eIF3 complex. The two domains are composed of about 150 to 200 amino acids at the N- or C-terminus of the CSN subunits. Apparently, the PCI domain has been shown to be important for interactions between CSN subunits. Thus, it might have a scaffolding function [22, 59].

6 The COP9 Signalosome and Its Role in the Ubiquitin System

The CSN subunit CSN5 has been shown to contain a metalloprotease motif localized on its MPN domain, which is essential for the cleavage of the ubiquitinlike modiﬁer NEDD8 from cullins [60] (see below). Apart from the catalytic activity of the MPN domain of CSN5 it appears to be the receptor for different cellular proteins associated with the CSN complex (see above and Figure 1). Interestingly, an MPN domain similar to that of CSN5 is located in the N-terminal region of CSN6. However, this MPN domain has no deneddylation catalytic center like CSN5. The function of the CSN6 MPN domain remains obscure.

4 Biochemical Activities Associated With the CSN 4.1 Deneddylation Activity

Studies in ﬁssion yeast and Arabidopsis have revealed that the CSN has a role in the cleavage of NEDD8 from cullins [54, 55, 61]. The MPN domain of CSN5, like its paralog subunit Rpn11 of the 26S proteasome lid complex, possesses a highly conserved pattern of four charged amino acid residues: one glutamate, two histidines and one aspartate. This pattern represents a new type of metalloprotease motif called the JAMM (Jab1/MPN domain metalloenzyme) or MPN+ motif [62, 63]. In CSN5 the catalytic region is important for the cleavage of the ubiquitinlike modiﬁer NEDD8 from its targets. Mutations in the conserved histidine and asparxstate residues of CSN5 led to suppression of its deneddylation activity [60]. Crystal-structure analysis obtained with bacterial CSN5/MPN+ domain-containing AF2198 protein conﬁrmed the metal-ion-dependent hydrolytic activity of CSN5, although it was inhibited by the alkylating agent NEM, an inhibitor of cysteine proteases [64]. NEDD8 is activated by a heterodimeric complex of APP-B1 and Ubα3 and is conjugated to target proteins by the conjugating enzyme Ubc12. So far, the only known targets are cullin-family proteins (CUL1–5), which are components of the cullin-based E3 ligase complexes. The covalent linkage of NEDD8 to cullins in vivo is thought to activate Ub-ligase complex activity by facilitating ubiquitin-conjugating enzyme E2 recruitment [65]. Deneddylation of cullins inactivates ubiquitination in vitro, but seems to stimulate the Ub E3 ligase complex activity in vivo [66, 67]. In cell lysates only a small fraction of CUL1 is neddylated, but in csn deletion cells 100% of CUL1 is modiﬁed by NEDD8. The puriﬁed CSN complex is able to deneddylate, although recombinant CSN5 protein cannot. Obviously CSN5 deneddylation activity is dependent on its assembly into the CSN complex [28, 54]. The fact that null mutants in most CSN subunits lack the deneddylation activity in the presence of excess CSN5 supports the fact that CSN5 alone is inactive in deneddylation [61]. So far, the exact role of deneddylation is questionable (see below).

4 Biochemical Activities Associated With the CSN 7

Fig. 2 Association of the CSN complex with enzymes. The Figure shows an electronmicroscopy image of puriﬁed CSN complex from human erythrocytes. As indicated by arrows the CSN is associated with the Ubspeciﬁc protease Ubp12, the proteasome, presumably with most of the cullin-based

Ub-ligase complexes, with a number of kinases, and with subunits of the translation initiation complex eIF3. In addition, subunit CSN5 has an intrinsic metalloprotease activity, which deneddylates cullins and also removes Ub conjugated to other proteins (for details see text).

4.2 Protein Kinases

The CSN is associated with enzymes such as kinases, proteases and Ub ligases, which perhaps, besides the intrinsic deneddylation activity, determine the speciﬁc function of the CSN in the Ub system. Here we summarize the associated (extrinsic) activities of the CSN shown in Figure 2. 4.2.1 Associated Protein Kinases Originally, a protein kinase was the ﬁrst enzyme identiﬁed with the CSN puriﬁed from human erythrocytes. The CSN-associated kinase activity phosphorylated several serine and threonine residues in the N-terminal region of c-Jun [5] resulting in stabilization of c-Jun and increased AP-1 transcriptional activity. The pathway responsible for this c-Jun stabilization/activation was called CSN-directed c-Jun signaling [68]. It was subsequently shown that the CSN-directed c-Jun signaling pathway controls most of the VEGF (vascular endothelial growth factor) production in tumor cells [13]. VEGF is essential for tumor angiogenesis (see below).

8 The COP9 Signalosome and Its Role in the Ubiquitin System

In contrast to c-Jun, phosphorylation of the tumor suppressor p53 by CSNassociated kinases targets the protein for degradation by the Ub system [35]. For p53 stability, modiﬁcation on Thr155 is most important as shown by mutational analysis [35] and by using different p53 peptides [31]. Mutation of Thr155 to Val led to stabilization of the transiently expressed p53 mutant in HeLa as well as in HL60 cells [35]. Inhibitors of CSN-associated kinases such as curcumin [18] caused stabilization of cellular p53 followed by massive cell death [35]. In addition to p53 and c-Jun, p27, ICSBP (interferon consensus sequence binding protein) and IκBα were identiﬁed as substrates of the CSN-associated kinases (for a review see Ref. [40]). Similar to p53, the phosphorylation of p27 results in accelerated degradation of the cyclin-dependent kinase inhibitor p27 by the Ub system (our unpublished data). In the case of ICSBP and IκBα, it is still unclear whether CSNmediated phosphorylation inﬂuences their stability. Interestingly, two of the CSN subunits, CSN2 and CSN7, are phosphorylated by the associated kinases [19, 69]. The physiological relevance of these modiﬁcations is currently obscure. 4.2.1.1 Identiﬁcation of associated protein kinases Based on phosphopeptide analyses it became clear that associated kinases modify principally serine and threonine residues. Moreover, the analysis of putative phosphorylation-speciﬁc consensus sequences of p53, c-Jun, p27, ICSBP and IκBα revealed that the protein kinase CK2 and a member of the protein kinase C family might be associated with the CSN. It has been shown by immunoblotting that CK2 and the protein kinase Cµ (also called protein kinase D, PKD) co-purify with the CSN from human erythrocytes [31]. In addition, the two kinases co-immunoprecipitated together with the CSN from HeLa cells. Interaction of CK2 as well as PKD with the CSN is mediated by CSN3, as is the interaction between CK2 and the CSN7 subunit [31]. Interestingly, CSN7 itself is phosphorylated. Majerus and co-workers have published work on the co-puriﬁcation of inositol 1,3,4-trisphosphate 5/6-kinase (5/6-kinase) with the CSN from bovine brain [24, 70]. Although the 5/6-kinase was not detected in the ﬁnal preparation of the CSN from human erythrocytes [31], it cannot be excluded that the enzyme is associated with another pool of CSN particles. The enzyme phosphorylates c-Jun, IκBα as well as p53 and is sensitive to curcumin. These characteristics are very similar to those described for CK2 and PKD. It has been shown that the 5/6-kinase interacts with CSN1 and that over-expression of CSN1 inhibits its activity [24]. It might be that it interacts with the N-terminal part of CSN1, which has been shown to suppress activation of an AP-1 promoter [22]. Future studies will show whether additional kinases besides 5/6-kinase, CK2 and PKD can interact with the CSN under certain circumstances. For example, an interaction of CSN3 with IKKγ , a component of the IKK kinase complex, has been published [32]. 4.2.1.2 Functions of associated protein kinases Phosphorylation of a number of Ub-dependent substrates by CSN-associated kinases regulates the stability of the proteins towards the Ub system [40], presumably by promoting substrate ubiquitination. Most of the proteins bind to the CSN via CSN5, are phosphorylated and

4 Biochemical Activities Associated With the CSN 9

subsequently channeled to the associated Ub ligase for ubiquitination (see below). Modiﬁcation of p53 induces a conformational change of the tumor suppressor, which leads to tighter binding to the Ub ligase [35]. In addition, there is evidence that phosphorylation might directly affect Ub-ligase activity. The transcriptional regulator Id3 interacts with the CSN, but is not phosphorylated. Nevertheless, inhibitors of CSN-associated kinases induce ubiquitination and degradation of the Id3 protein [41]. Because of associated kinases and their function in ubiquitination the CSN has been described as a complex “at the interface between signal transduction and ubiquitin-dependent proteolysis” [40]. This becomes even more signiﬁcant if upstream regulation of the associated kinases is taken into account. Unfortunately at the moment little is known about the receptors or signal-transduction pathways leading to modiﬁcation of the CSN and its associated kinase activities. It is also unclear whether there are interactions between the kinases and other associated activities of the CSN. 4.3 Deubiquitinating Enzymes

To date two deubiquitinating activities associated with the CSN have been identiﬁed. By mutational analysis one deubiquitinating activity has been mapped to the metalloprotease motif His–X–His–X10–Asp of the JAMM or MPN+ domain of CSN5 [11]. The conserved Asp residue of that motif was mutated and the mutant Flag-CSN5 was integrated into the CSN. The mutated CSN lost its ability to remove ubiquitin from the isopeptide bond of a mono-ubiquitinated conjugate [11]. Obviously the same intrinsic metalloprotease activity is responsible for deneddylation of mono-neddylated cullins [60], which is not surprising because of the homologies between Ub and NEDD8. It would be interesting to test whether the lid subunit Rpn11, which exhibits a deubiquitinating MPN+ domain [62], is able to deneddylate mono-neddylated proteins. In addition, another deubiquitinating activity associated with the CSN disassembles poly-Ub chains [11, 71]. This activity is catalyzed in ﬁssion yeast by the Ub-speciﬁc protease Ubp12, which is a CSN-associated enzyme [71]. The interaction of Ubp12 with the CSN is required for Ubp12 transport to the nucleus. Presumably in the nucleus, S. pombe Pcu1- and Pcu3-based Ub-ligase activities are inhibited by Ubp12 enzyme, since the deubiquitinating enzyme protects a speciﬁc adapter protein, Pop1p, from autocatalytic destruction [71]. Thus it seems that the CSN has dual activity in suppressing cullin-based Ub-ligase reactions: one is the intrinsic deneddylation and the other is deubiquitination via associated Ubp12; both reactions serve to inhibit cullin-based Ub ligases in vitro. Data have accumulated showing that CSN-associated deneddylation and deubiquitination are required for Ub-ligase activity in vivo. It has been hypothesized, therefore, that CSN-mediated inhibition of cullin-based ubiquitination might be necessary for the assembly of new cullin-based Ub-ligase complexes. After release from the CSN the new cullin-based complex would be active. It has to return to the

10 The COP9 Signalosome and Its Role in the Ubiquitin System

CSN for re-assembly or is degraded after auto-ubiquitination [71, 72]. For example, p27 has to be degraded at the transition from G1 to S phase. In a ﬁrst step p27 may bind to the CSN which signals, perhaps by phosphorylation, the assembly of the required SCF complex containing the speciﬁc F-box protein Skp2. After formation of the p27-speciﬁc SCF complex both p27 and the Ub ligase might be released from the CSN perhaps again by phosphorylation which then results in ubiquitination and complete degradation of p27. Finally the Skp2-containing SCF complex is auto-ubiquitinated and degraded unless additional substrate appears. 4.4 Ubiquitin Ligases

Data have been accumulated demonstrating interactions of the CSN with Ub ligases, in particular with the cullin-based Ub ligases. Cullins 1 to 7 (CUL1–CUL7) form a protein family detected in all eukaryotic cells, which is involved in protein ubiquitination. It is known that CUL1 to CUL5 interact with the RING-domain protein Rbx1, the Ub ligase of the cullin-based complexes. So far it has been shown that the CSN interacts with CUL1 to CUL4 [11, 12, 54, 55, 61, 73]. Binding studies with CUL1 and with CUL2 revealed that the two cullin proteins bind via CSN2 to the complex [28, 54, 55]. In addition, Rbx1 seems to interact with CSN6 [54, 55]. Moreover, CUL1 interacts with Skp1, which makes the connection to a substrate-speciﬁc F-box protein. Therefore, CUL1-based Ub ligases are called SCF complexes (Skp1CDC53/CUL1-F-box protein) (for a review see Ref. [74]). CUL2 can be linked to the substrate-adapter protein the von Hippel–Lindau tumor suppressor via elongin C and elongin B forming the so called VCB (von Hippel–Lindau–elongin C–elongin B) complex (for a review see Ref. [74]). BTB/POZ-domain proteins have been identiﬁed as possible substrate-speciﬁc adaptors of CUL3-based Ub ligases [73, 75, 76]. There are more than 200 putative BTB/POZ-domain proteins expressed in mammalian cells and together with the large number of possible F-box proteins one can estimate that several hundreds of different cullin-based Ub-ligase complexes with different substrate speciﬁcities can be formed. The CUL4–Rbx1 complex has been characterized, and seems to be important for checkpoint control [12], DNA repair [11] and ubiquitination of c-Jun [77]. Most likely all cullin-based complexes interact with the CSN. In other words, the CSN is associated with ubiquitinating activity (see Figure 2). There are just a few data on interactions of the CSN with other Ub ligases besides the cullin-based complexes. For example, Mdm2, the RING domain Ub ligase of the tumor suppressor p53, binds to the CSN and is modiﬁed by CSN-associated kinases (our unpublished data). Whether Mdm2 is also modiﬁed by other CSN-associated activities has to be tested in the future. In addition, COP1, a putative RING-domain Ub ligase, which probably cooperates with the COP1-interacting protein 8 (CIP8) also binds to the CSN (for a review see Ref. [4]). However, some data indicate that COP1 is associated with a CUL4A complex in which it acts together with DET1 as a heterodimeric substrate adapter [77]. In this complex the CSN interacts with both the CUL4A and the COP1.

5 Association of the CSN With Other Protein Complexes 11

5 Association of the CSN With Other Protein Complexes 5.1 The eIF3 Complex

MPN and PCI domains have been also found in subunits of the eIF3 complex. Because MPN and PCI domains are most likely involved in protein–protein interactions (see above), it is not surprising that there are also cross-interactions between subunits of the CSN, the eIF3 and the lid. It has been reported that eIF3e/INT6 possessing a PCI domain interacts with CSN7 [33, 78]. Another eIF3 subunit eIF3c/p105 co-immunoprecipitated with eIF3e/INT6, eIF3b, CSN1 and CSN8 [78]. eIF3e/INT6 was used as bait in a two-hybrid screen that revealed possible interactions with the 26S proteasome ATPase Rpt4, CSN3 and CSN6 but also with CSN7 [33]. Interestingly, the subunit of the CSN-like complex in Saccharomyces cerevisiae Pci8/CSN11 [79] seems also to be a subunit of the budding yeast eIF3 complex and perhaps plays a similar role to eIF3e/INT6 in eukaryotic cells [80]. It has been speculated that these interactions allow the CSN to control translation. Interactions between eIF3e/INT6 or eIF3i with the 26S proteasome have also been described [33, 81]. It has been shown that eIF3e/INT6 interacts with Rpn5 of the lid complex. This has an impact on 26S proteasome activity/localization, presumably affecting cell division and mitotic ﬁdelity [82]. Perhaps there exists a network of “PCI complexes” as suggested [83], which shares polypeptides and communicates via proteins such as eIF3e/INT6. 5.2 The Proteasome

In 1998 it was reported that the CSN co-fractionates with the 26S proteasome from human cells [5]. A yeast two-hybrid screen revealed that the C-terminal domain of the Arabidopsis atCSN1 subunit interacts with atRpn6 of the 26S proteasome lid [26]. Recently gel-ﬁltration size-fractionation of material from Arabidopsis in the presence of ATP and phosphatase inhibitors indicated that the CSN1 and CSN6 subunits co-elute in the same fractions as subunits of the 26S regulatory complex [84]. Based on these data it has been speculated that the CSN might be an alternative lid of the 26S proteasome [85]. The “alternative lid hypothesis”, however, makes little sense if the CSN interacts with the 26S proteasome via the lid component Rpn6 [26]. CSN pull-down experiments and subsequent mass-spectrometry analysis of co-precipitated proteins also revealed the presence of proteasome subunits in the precipitate [73]. However, since proteasome subunits are very abundant in cells, one has to be cautious with this type of data. So far there is no systematic binding study showing physical interaction of the CSN with sub-complexes of the 26S proteasome. Moreover, up to now there is no functional evidence for such a CSN/26S proteasome interaction.

12 The COP9 Signalosome and Its Role in the Ubiquitin System

6 Biological Functions of the CSN 6.1 Regulation of Ubiquitin Conjugate Formation

In general, and including all its activities, intrinsic as well as associated, the CSN seems to be a regulator of ubiquitination. Deneddylation, deubiquitination as well as CSN-mediated phosphorylation (at least with c-Jun and Id3 as substrates) cause inhibition of ubiquitination. It is likely that suppression of ligase activity is an essential step in the dynamic process of speciﬁc E3 complex assembly/reassembly. According to the model of Wolf et al. [72] cullin-based Ub-ligase complexes might assemble/reassemble in a protected environment produced by the CSN. In the CSN-associated-state, binding of any E2 to the Ub ligase is prevented, perhaps by deneddylation [65], self-ubiquitination is blocked by continuous deubiquitination [71] and substrate binding could be inhibited by phosphorylation [31]. Only under these conditions can the Ub ligase reassemble without itself being destroyed. For example, an SCF complex might associate with another F-box protein, or a CUL3Ub ligase with another BTB/POZ-domain protein, as an adaptation to the next phase of cell cycle or signal transduction upon the appearance of a new substrate, which has to be degraded. Following this argument a major question arises. How does the substrate signal the assembly of the required Ub ligase performing its ubiquitination? Is it by binding to the CSN and subsequent signaling via speciﬁc kinases? In the case of the SCF complexes, another protein called CAND1/Tip120A seems to be involved in the dynamic assembly/reassembly process of the E3 [86]. CAND1 binds to the deneddylated CUL1 and inhibits Ub-ligase activity by competing for the Skp1–F-box-protein unit of the SCF complex [87]. After the release of CAND1, a new Skp1–F-box-protein unit can dock to the CUL1-Rbx1 unit to form an SCF complex possessing the necessary substrate speciﬁcity. Now the freshly formed Ub ligase has to be released from the CSN to become active. At the moment it is unclear how the Ub ligase might be released from the CSN. The attractive model of CSN-assisted Ub-ligase-complex assembly has to be tested in the future. In this model the CSN would function as a platform for Ub-ligase assembly. Interestingly, there are no reports of interactions between the 26S proteasome lid complex and Ub ligases. Known E3s directly interacting with the 26S proteasome seem to bind via base ATPases [88, 89]. This is an interesting functional difference between the CSN and the lid developed during evolution. In an alternative model the CSN might be the platform for complete proteolysis. It forms supercomplexes consisting of both the ubiquitinating and the proteolytic machineries. According to this model, the substrate ﬁrst binds to the CSN, is then ubiquitinated by the associated Ub ligase and ﬁnally directly channeled into the 26S proteasome. Deneddylation, deubiquitination and phosphorylation are necessary to maintain the supercomplex, to protect the intermediates and to stimulate proteolysis.

6 Biological Functions of the CSN 13

6.1.1 Cell-cycle and Checkpoint Control Initial insight of the role of CSN in cell-cycle control came from the ﬁnding that csn1 and csn2 deletion S. pombe strains have an S-phase delay [52]. Interestingly, this effect did not occur in strains missing other CSN subunits. The S-phase delay was caused by the accumulation of the cell-cycle inhibitor Spd1 (S-phase delayed 1), which is involved in the misregulation of the ribonucleotide reductase (RNR). RNR catalyzes the production of deoxyribonucleotides for DNA synthesis and is composed of four subunits including Suc22. Activation of RNR is regulated by nuclear export of Suc22, which is suppressed by Spd1 [12]. Upon DNA damage or during S phase Spd1 is rapidly degraded, presumably leading to the RNRdependent production of dNTPs. However, in csn1 and csn2 deletion mutants, Spd1 accumulates, causing Suc22-dependent suppression of RNR connected with the S-phase delay and DNA-damage sensitivity [12, 15]. In mammalian cells, binding of HIV-1 Vpr-protein to the CSN6 results in cellcycle arrest at the G2/M phase [53]. Additionally, CSN is involved in the cell cycle via the nuclear export of cell-cycle kinase inhibitor p27kip1 (p27). CSN5 binds to p27 and promotes its nuclear export followed by its proteasome-dependent degradation. The over-expression of CSN5 in mouse ﬁbroblasts counteracts cell-cycle arrest induced by serum depletion [36, 51]. Microinjection of the puriﬁed CSN complex into synchronized G1 cells blocks the S-phase entry in a deneddylation-dependent manner [28]. Furthermore, the reduction of CSN subunit expression by RNAi in Caenorhabditis elegans causes the failure of Mei-1 degradation by regulation of its speciﬁc Ub-ligase CUL3-based complex, which leads to severe effects during mitotic cell division [76]. Moreover, the CSN is involved in checkpoint control. The double deletions of csn1 and csn2 mutants crossed with checkpoint pathway mutants such as rad3, chk, and cds1 are synthetically lethal in S. pombe [52]. Cds1 kinase is constitutively activated in csn1 mutants. Similarly, loss of csn5 in Drosophila results in activation of Mei-41, one of the ATM/ATR family kinases involved in meiotic checkpoint upon DNA damage [90]. 6.1.2 DNA Repair Two papers have assigned the CSN a function in DNA repair. One study reports on the existence of two different complexes containing human CSN and either one of the two nucleotide-excision-repair proteins, DDB2 or CSA. DDB2 is involved in the global genome-repair pathway (GGR) and CSA functions in the transcriptioncoupled repair pathway (TCR). Additionally, these complexes possess Ub-ligase activity and contain cullin-based Ub-ligase components such as CUL4 and Rbx1/Roc1, and DDB1, a UV-damage DNA-binding protein [11]. However, so far their targets remain unclear. CSN differentially regulates the ubiquitin-ligase activity of the DDB2- and CSA-containing complexes in response to UV irradiation. In support of direct involvement of the CSN is the ﬁnding that knockdown of CSN5 with RNAi causes a failure in NER mechanisms [11]. Similarly, CSN in combination with the CUL4–Rbx1 complex is involved in Ub-dependent degradation of CDT1, a licensing factor of the pre-replication complex (preRC), after UV- or γ -irradiation.

14 The COP9 Signalosome and Its Role in the Ubiquitin System

Knockdown of CSN completely suppresses CDT1 degradation, causing a defect G1 checkpoint in response to DNA damage [91]. 6.1.3 Developmental Processes Although CSN is not essential in yeast, the csn1 and csn2 S. pombe deletion mutants display slow growth and sensitivity to UV- and γ -irradiation [52]. Other csn mutants did not show signiﬁcant phenotypes apart from the loss of cullin’s deneddylation activity [61]. In mutants of CSN-like complexes in the budding yeast S. cerevisiae the sensitivity to the DNA-damage reagents is not affected [92]. In some S. cerevisiae mutants such as csn5, csn9 and csn12 deletions, increased mating efﬁciency and enhanced pheromone response has been observed [63]. In Drosophila, mutations of CSN causes lethality in early larval stages and defects during oogenesis or photoreceptor R cell differentiation [9, 90, 93, 94]. More specifically, lack of CSN5 leads to the activation of a DNA double-strand-break-dependent checkpoint mediated by Mei-41. This effect is caused by CSN5-dependent inhibition of gurken (Grk) protein translation [90]. In C. elegans, knockdown of CSN5 by RNAi resulted in a sterile phenotype, which could be explained by CSN interaction with germ-line RNA helicases [48]. The best studied physiological role of the CSN in developmental processes is derived from studies on Arabidopsis. Csn mutants can survive embryogenesis, but they die soon after germination. The csn mutants exhibit a defect in photomorphogenesis, a light-dependent developmental process of germinating seedlings. Even in total darkness the mutants display a light-dependent morphology and signalindependent expression of light-induced genes [3, 14, 95]. One key mechanism is the CSN-dependent regulation of the stability of the transcription factor HY5, a positive regulator of light-induced genes. In the dark it is degraded by the Ub system [8]. It has been suggested that in darkness the RING-ﬁnger protein COP1 ubiquitinates HY5 and triggers its degradation by the 26S proteasome (for a review see Ref. [4]). In the light, COP1 is relocated to the cytoplasm allowing expression of genes through HY5. Although the exact mechanism remains unclear, the CSN may be required for relocation of COP1 from cytoplasm to the nucleus in darkness. Identical phenotypes caused by different csn mutants in Arabidopsis could be explained by a role of the CSN as a whole complex (for a review see Ref. [20]). There is accumulating evidence for cooperation of the CSN and cullin-based complexes in speciﬁc developmental processes [96]. First insight has been provided by studies on auxin response where the CSN interacts with SCFTIR1 , modulating its activity [55]. Similarly, binding of the CSN to other cullin-based complexes regulates their activity in mediating various developmental processes such as ﬂower development, and plant defense responses [97, 98]. 6.2 Tumor Angiogenesis

Tumor angiogenesis is the vascularization of solid tumors, an essential requirement for tumor growth and metastasis. After a solid tumor has reached a size

6 Biological Functions of the CSN 15

of approximately 2 mm3 , it needs nutrient supply from blood vessels, otherwise it dies from necrosis. Many tumor cells are able to induce angiogenesis. In an initiation phase the tumor cells produce large amounts of pro-angiogenic factors such as vascular endothelial growth factor, VEGF. During proliferation and invasion VEGF stimulates migration of endothelial cells. Finally, after a maturation phase, vascularization of solid tumors is completed. Now the tumor can grow, and some tumor cells penetrate through vessel membranes and spread via the circulation. Therefore, inhibition of tumor angiogenesis has become an important strategy in tumor therapy. There is functional cooperation between the CSN and the Ub system in tumor angiogenesis [13]. It has been known for some time that curcumin is an inhibitor of angiogenesis [99]. However, only in 2001 did it become clear that it acts via inhibition of CSN-associated kinases [13]. It has been demonstrated that over-expression of CSN2 subunit leads to elevated amounts of de novo assembled CSN complex connected with increased c-Jun levels and enhanced AP-1 transactivation activity [68]. This c-Jun activation/stabilization is independent of the JNK and the MAP kinase pathway and is called CSN-directed c-Jun signaling [68]. This process can be inhibited by curcumin or other inhibitors of CSN-associated kinases (Figure

Fig. 3 The CSN-directed c-Jun signaling pathway. (A) The active CSN-directed cJun signaling pathway is shown. In case of active CSN-associated kinases c-Jun is phosphorylated, which stabilizes the transcription factor towards the Ub system. In addition, phosphorylation of the responsible E3 might inactivate the enzyme. In this situation Id1 and Id3 are also stabilized. Stable/active c-Jun causes enhanced AP-1 transactivation connected with an increase of VEGF production by tumor cells (see

text). VEGF is a major pro-angiogenic factor produced by many tumor cells. Id1 and Id3 are transcriptional regulators essential for tumor angiogenesis. (B) In the presence of curcumin or other kinase inhibitors the responsible Ub ligase is most likely active and ubiquitinates both c-Jun and Id3. In addition, unphosphorylated c-Jun might have higher afﬁnity to its Ub ligase. This leads to quick degradation of the proteins by the Ub system.

16 The COP9 Signalosome and Its Role in the Ubiquitin System

3). The CSN-directed c-Jun signaling controls up to 75% of VEGF production in tumor cells [13]. In addition, Id1 and Id3 are also essential factors of tumor angiogenesis [100] and are degraded in the presence of CSN-associated kinase inhibitors in an Ub-dependent manner just like c-Jun [41]. Therefore, speciﬁc inhibition of CSN-associated kinases might become important for tumor therapy. The application of CSN-associated kinase inhibitors in tumor therapy could be beneﬁcial owing to another effect of curcumin-like compounds, namely they stabilize cellular p53 and, at least in tumors with wild-type p53 protein, massive cell death can be observed [35].

7 Conclusions

The CSN is a regulatory complex of the Ub system. Physically it interacts with the proteasome and with Ub ligases. Although the exact mechanism remains obscure, the CSN regulates ubiquitination of important cell-cycle factors and transcriptional regulators. Its intrinsic deneddylating as well as the associated kinase and deubiquitinating activities seem to be required for determining protein stability towards the Ub system. As a major regulator of the Ub system the CSN is involved in processes such as DNA repair, cell-cycle progression and development. Its role in tumor angiogenesis makes the complex attractive for future tumor therapies.

Acknowledgment

We thank Rasmus Hartmann-Petersen for critical reading of the manuscript. The work was supported by research grants from the Deutsche Forschungsgemeinschaft and from the German-Israeli Foundation to W. D.

References 1 DENG, X. W., DUBIEL, W., WEI, N., HOFMANN, K., MUNDT, K., COLICELLI, J., KATO, J., NAUMANN, M., SEGAL, D., SEEGER, M., CARR, A., GLICKMAN, M., CHAMOVITZ, D. A. Uniﬁed nomenclature for the COP9 signalosome and its subunits: an essential regulator of development. Trends Genet. 2000, 16, 202–203. 2 WEI, N., CHAMOVITZ, D. A., DENG, X. W. Arabidopsis COP9 is a component of a novel signaling complex mediating light control of development. Cell 1994, 78, 117–124.

3 WEI, N., DENG, X. W. Making sense of the COP9 signalosome. A regulatory protein complex conserved from Arabidopsis to human. Trends Genet. 1999, 15, 98–103. 4 SERINO, G., DENG, X. W. The COP9 signalosome: regulating plant development through the control of proteolysis. Annu. Rev. Plant Biol. 2003, 54, 165–182. 5 SEEGER, M., KRAFT, R., FERRELL, K., BECH-OTSCHIR, D., DUMDEY, R., SCHADE, R., GORDON, C., NAUMANN, M., DUBIEL, W. A novel protein complex involved in

References

6

7

8

9

10

11

12

13

signal transduction possessing similarities to 26S proteasome subunits. FASEB J. 1998, 12, 469–478. WEI, N., TSUGE, T., SERINO, G., DOHMAE, N., TAKIO, K., MATSUI, M., DENG, X. W. The COP9 complex is conserved between plants and mammals and is related to the 26S proteasome regulatory complex. Curr. Biol. 1998, 8, 919–922. GLICKMAN, M. H., RUBIN, D. M., COUX, O., WEFES, I., PFEIFER, G., CJEKA, Z., BAUMEISTER, W., FRIED, V. A., FINLEY, D. A subcomplex of the proteasome regulatory particle required for ubiquitin-conjugate degradation and related to the COP9-signalosome and eIF3. Cell 1998, 94, 615–623. SCHWECHHEIMER, C., DENG, X. W. COP9 signalosome revisited: a novel mediator of protein degradation. Trends Cell Biol. 2001, 11, 420–426. FREILICH, S., ORON, E., KAPP, Y., NEVO-CASPI, Y., ORGAD, S., SEGAL, D., CHAMOVITZ, D. A. The COP9 signalosome is essential for development of Drosophila melanogaster. Curr. Biol. 1999, 9, 1187–1190. LYKKE-ANDERSEN, K., SCHAEFER, L., MENON, S., DENG, X. W., MILLER, J. B., WEI, N. Disruption of the COP9 signalosome Csn2 subunit in mice causes deﬁcient cell proliferation, accumulation of p53 and cyclin E, and early embryonic death. Mol. Cell. Biol. 2003, 23, 6790–6797. GROISMAN, R., POLANOWSKA, J., KURAOKA, I., SAWADA, J., SAIJO, M., DRAPKIN, R., KISSELEV, A. F., TANAKA, K., NAKATANI, Y. The ubiquitin ligase activity in the DDB2 and CSA complexes is differentially regulated by the COP9 signalosome in response to DNA damage. Cell 2003, 113, 357–367. LIU, C., POWELL, K. A., MUNDT, K., WU, L., CARR, A. M., CASPARI, T. Cop9/signalosome subunits and Pcu4 regulate ribonucleotide reductase by both checkpoint-dependent and -independent mechanisms. Genes Dev. 2003, 17, 1130–1140. POLLMANN, C., HUANG, X., MALL, J., BECH-OTSCHIR, D., NAUMANN, M.,

14

15

16

17

18

19

20

21

DUBIEL, W. The constitutive photomorphogenesis 9 signalosome directs vascular endothelial growth factor production in tumor cells. Cancer Res. 2001, 61, 8416–8421. OSTERLUND, M. T., WEI, N., DENG, X. W. The roles of photoreceptor systems and the COP1-targeted destabilization of HY5 in light control of Arabidopsis seedling development. Plant Physiol. 2000, 124, 1520–1524. CLARET, F. X., HIBI, M., DHUT, S., TODA, T., KARIN, M. A new group of conserved coactivators that increase the speciﬁcity of AP-1 transcription factors. Nature 1996, 383, 453–457. LEE, J. W., CHOI, H. S., GYURIS, J., BRENT, R., MOORE, D. D. Two classes of proteins dependent on either the presence or absence of thyroid hormone for interaction with the thyroid hormone receptor. Mol. Endocrinol. 1995, 9, 243–254. WEI, N., DENG, X. W. Characterization and puriﬁcation of the mammalian COP9 complex, a conserved nuclear regulator initially identiﬁed as a repressor of photomorphogenesis in higher plants. Photochem. Photobiol. 1998, 68, 237–241. HENKE, W., FERRELL, K., BECH-OTSCHIR, D., SEEGER, M., SCHADE, R., JUNGBLUT, P., NAUMANN, M., DUBIEL, W. Comparison of human COP9 signalsome and 26S proteasome lid’. Mol. Biol. Rep. 1999, 26, 29–34. KAPELARI, B., BECH-OTSCHIR, D., HEGERL, R., SCHADE, R., DUMDEY, R., DUBIEL, W. Electron microscopy and subunit–subunit interaction studies reveal a ﬁrst architecture of COP9 signalosome. J. Mol. Biol. 2000, 300, 1169–1178. WEI, N., DENG, X. W. The COP9 signalosome. Annu. Rev. Cell Dev. Biol. 2003, 19, 261–286. SPAIN, B. H., BOWDISH, K. S., PACAL, A. R., STAUB, S. F., KOO, D., CHANG, C. Y., XIE, W., COLICELLI, J. Two human cDNAs, including a homolog of Arabidopsis FUS6 (COP11), suppress G-protein- and mitogen-activated protein kinase-mediated signal transduction in

17

18 The COP9 Signalosome and Its Role in the Ubiquitin System

22

23

24

25

26

27

28

29

yeast and mammalian cells. Mol. Cell. Biol. 1996, 16, 6698–6706. TSUGE, T., MATSUI, M., WEI, N. The subunit 1 of the COP9 signalosome suppresses gene expression through its N-terminal domain and incorporates into the complex through the PCI domain. J. Mol. Biol. 2001, 305, 1–9. WANG, X., KANG, D., FENG, S., SERINO, G., SCHWECHHEIMER, C., WEI, N. CSN1 N-terminal-dependent activity is required for Arabidopsis development but not for Rub1/Nedd8 deconjugation of cullins: a structure-function study of CSN1 subunit of COP9 signalosome. Mol. Biol. Cell 2002, 13, 646–655. SUN, Y., WILSON, M. P., MAJERUS, P. W. Inositol 1,3,4-trisphosphate 5/6-kinase associates with the COP9 signalosome by binding to CSN1. J. Biol. Chem. 2002, 277, 45759–45764. KARNIOL, B., YAHALOM, A., KWOK, S., TSUGE, T., MATSUI, M., DENG, X. W., CHAMOVITZ, D. A. The Arabidopsis homologue of an eIF3 complex subunit associates with the COP9 complex. FEBS Lett. 1998, 439, 173–179. KWOK, S. F., STAUB, J. M., DENG, X. W. Characterization of two subunits of Arabidopsis 19S proteasome regulatory complex and its possible interaction with the COP9 complex. J. Mol. Biol. 1999, 285, 85–95. DRESSEL, U., THORMEYER, D., ALTINCICEK, B., PAULULAT, A., EGGERT, M., SCHNEIDER, S., TENBAUM, S. P., RENKAWITZ, R., BANIAHMAD, A. Alien, a highly conserved protein with characteristics of a corepressor for members of the nuclear hormone receptor superfamily. Mol. Cell. Biol. 1999, 19, 3383–3394. YANG, X., MENON, S., LYKKE-ANDERSEN, K., TSUGE, T., DI, X., WANG, X., RODRIGUEZ-SUAREZ, R. J., ZHANG, H., WEI, N. The COP9 signalosome inhibits p27(kip1) degradation and impedes G1-S phase progression via deneddylation of SCF Cul1. Curr. Biol. 2002, 12, 667–672. COHEN, H., AZRIEL, A., COHEN, T., MERARO, D., HASHMUELI, S., BECH-OTSCHIR, D., KRAFT, R., DUBIEL, W., LEVI, B. Z. Interaction between interferon consensus sequence-binding

30

31

32

33

34

35

36

37

protein and COP9/signalosome subunit CSN2 (Trip15). A possible link between interferon regulatory factor signaling and the COP9/signalosome. J. Biol. Chem. 2000, 275, 39081–39089. ALTINCICEK, B., TENBAUM, S. P., DRESSEL, U., THORMEYER, D., RENKAWITZ, R., BANIAHMAD, A. Interaction of the corepressor Alien with DAX-1 is abrogated by mutations of DAX-1 involved in adrenal hypoplasia congenita. J. Biol. Chem. 2000, 275, 7662–7667. UHLE, S., MEDALIA, O., WALDRON, R., DUMDEY, R., HENKLEIN, P., BECH-OTSCHIR, D., HUANG, X., BERSE, M., SPERLING, J., SCHADE, R., DUBIEL, W. Protein kinase CK2 and protein kinase D are associated with the COP9 signalosome. EMBO J. 2003, 22, 1302–1312. HONG, X., XU, L., LI, X., ZHAI, Z., SHU, H. CSN3 interacts with IKKgamma and inhibits TNF- but not IL-1-induced NF-kappaB activation. FEBS Lett. 2001, 499, 133–136. HOAREAU ALVES, K., BOCHARD, V., RETY, S., JALINOT, P. Association of the mammalian proto-oncoprotein Int-6 with the three protein complexes eIF3, COP9 signalosome and 26S proteasome. FEBS Lett. 2002, 527, 15–21. SUZUKI, G., YANAGAWA, Y., KWOK, S. F., MATSUI, M., DENG, X. W. Arabidopsis COP10 is a ubiquitin-conjugating enzyme variant that acts together with COP1 and the COP9 signalosome in repressing photomorphogenesis. Genes Dev. 2002, 16, 554–559. BECH-OTSCHIR, D., KRAFT, R., HUANG, X., HENKLEIN, P., KAPELARI, B., POLLMANN, C., DUBIEL, W. COP9 signalosome-speciﬁc phosphorylation targets p53 to degradation by the ubiquitin system. EMBO J. 2001, 20, 1630–1639. TOMODA, K., KUBOTA, Y., KATO, J. Degradation of the cyclin-dependent-kinase inhibitor p27Kip1 is instigated by Jab1. Nature 1999, 398, 160–165. LI, S., LIU, X., ASCOLI, M. p38JAB1 binds to the intracellular precursor of the

References

38

39

40

41

42

43

44

45

lutropin/choriogonadotropin receptor and promotes its degradation. J. Biol. Chem. 2000, 275, 13386–13393. WAN, M., CAO, X., WU, Y., BAI, S., WU, L., SHI, X., WANG, N. Jab1 antagonizes TGF-beta signaling by inducing Smad4 degradation. EMBO Rep. 2002, 3, 171–176. BAE, M. K., AHN, M. Y., JEONG, J. W., BAE, M. H., LEE, Y. M., BAE, S. K., PARK, J. W., KIM, K. R., KIM, K. W. Jab1 interacts directly with HIF-1alpha and regulates its stability. J. Biol. Chem. 2002, 277, 9–12. BECH-OTSCHIR, D., SEEGER, M., DUBIEL, W. The COP9 signalosome: at the interface between signal transduction and ubiquitin-dependent proteolysis. J. Cell Sci. 2002, 115, 467–473. BERSE, M., BOUNPHENG, M. A., HUANG, X., CHRISTY, B. A., POLLMANN, C., DUBIEL, W. Ubiquitin-dependent degradation of Id1 and Id3 is mediated by the COP9 signalosome. J. Mol. Biol. 2004, 343, 361–370. DECHEND, R., HIRANO, F., LEHMANN, K., HEISSMEYER, V., ANSIEAU, S., WULCZYN, F. G., SCHEIDEREIT, C., LEUTZ, A. The Bcl-3 oncoprotein acts as a bridging factor between NF-kappaB/Rel and nuclear coregulators. Oncogene 1999, 18, 3316–3323. CHAUCHEREAU, A., GEORGIAKAKI, M., PERRIN-WOLFF, M., MILGROM, E., LOOSFELT, H. JAB1 interacts with both the progesterone receptor and SRC-1. J. Biol. Chem. 2000, 275, 8540–8548. BIANCHI, E., DENTI, S., GRANATA, A., BOSSI, G., GEGINAT, J., VILLA, A., ROGGE, L., PARDI, R. Integrin LFA-1 interacts with the transcriptional co-activator JAB1 to modulate AP-1 activity. Nature 2000, 404, 617–621. KLEEMANN, R., HAUSSER, A., GEIGER, G., MISCHKE, R., BURGER-KENTISCHER, A., FLIEGER, O., JOHANNES, F. J., ROGER, T., CALANDRA, T., KAPURNIOTU, A., GRELL, M., FINKELMEIER, D., BRUNNER, H., BERNHAGEN, J. Intracellular action of the cytokine MIF to modulate AP-1 activity and the cell cycle through Jab1. Nature 2000, 408, 211–216.

46 GEMMILL, R. M., BEMIS, L. T., LEE, J. P., SOZEN, M. A., BARON, A., ZENG, C., ERICKSON, P. F., HOOPER, J. E., DRABKIN, H. A. The TRC8 hereditary kidney cancer gene suppresses growth and functions with VHL in a common pathway. Oncogene 2002, 21, 3507–3516. 47 LU, C., LI, Y., ZHAO, Y., XING, G., TANG, F., WANG, Q., SUN, Y., WEI, H., YANG, X., WU, C., CHEN, J., GUAN, K. L., ZHANG, C., CHEN, H., HE, F. Intracrine hepatopoietin potentiates AP-1 activity through JAB1 independent of MAPK pathway. FASEB J. 2002, 16, 90–92. 48 SMITH, P., LEUNG-CHIU, W. M., MONTGOMERY, R., ORSBORN, A., KUZNICKI, K., GRESSMAN-COBERLY, E., MUTAPCIC, L., BENNETT, K. The GLH proteins, Caenorhabditis elegans P granule components, associate with CSN-5 and KGB-1, proteins necessary for fertility, and with ZYX-1, a predicted cytoskeletal protein. Dev. Biol. 2002, 251, 333–347. 49 CABALLERO, O. L., RESTO, V., PATTURAJAN, M., MEERZAMAN, D., GUO, M. Z., ENGLES, J., YOCHEM, R., RATOVITSKI, E., SIDRANSKY, D., JEN, J. Interaction and colocalization of PGP9.5 with JAB1 and p27(Kip1). Oncogene 2002, 21, 3003–3010. 50 CHAMOVITZ, D. A., SEGAL, D. JAB1/CSN5 and the COP9 signalosome. A complex situation. EMBO Rep. 2001, 2, 96–101. 51 TOMODA, K., KUBOTA, Y., ARATA, Y., MORI, S., MAEDA, M., TANAKA, T., YOSHIDA, M., YONEDA-KATO, N., KATO, J. Y. The cytoplasmic shuttling and subsequent degradation of p27Kip1 mediated by Jab1/CSN5 and the COP9 signalosome complex. J. Biol. Chem. 2002, 277, 2302–2310. 52 MUNDT, K. E., PORTE, J., MURRAY, J. M., BRIKOS, C., CHRISTENSEN, P. U., CASPARI, T., HAGAN, I. M., MILLAR, J. B., SIMANIS, V., HOFMANN, K., CARR, A. M. The COP9/signalosome complex is conserved in ﬁssion yeast and has a role in S phase. Curr. Biol. 1999, 9, 1427–1430. 53 MAHALINGAM, S., AYYAVOO, V., PATEL, M., KIEBER-EMMONS, T., KAO, G. D., MUSCHEL, R. J., WEINER, D. B. HIV-1 Vpr

19

20 The COP9 Signalosome and Its Role in the Ubiquitin System

54

55

56

57

58

59

60

61

62

interacts with a human 34-kDa mov34 homologue, a cellular factor linked to the G2/M phase transition of the mammalian cell cycle. Proc. Natl. Acad. Sci. USA 1998, 95, 3419–3424. LYAPINA, S., COPE, G., SHEVCHENKO, A., SERINO, G., TSUGE, T., ZHOU, C., WOLF, D. A., WEI, N., DESHAIES, R. J. Promotion of NEDD-CUL1 conjugate cleavage by COP9 signalosome. Science 2001, 292, 1382–1385. SCHWECHHEIMER, C., SERINO, G., CALLIS, J., CROSBY, W. L., LYAPINA, S., DESHAIES, R. J., GRAY, W. M., ESTELLE, M., DENG, X. W. Interactions of the COP9 signalosome with the E3 ubiquitin ligase SCFTIRI in mediating auxin response. Science 2001, 292, 1379–1382. FU, H., REIS, N., LEE, Y., GLICKMAN, M. H., VIERSTRA, R. D. Subunit interaction maps for the regulatory particle of the 26S proteasome and the COP9 signalosome. EMBO J. 2001, 20, 7096–7107. WANG, Y., DEVEREUX, W., STEWART, T. M., CASERO, R. A., Jr. Polyamine-modulated factor 1 binds to the human homologue of the 7a subunit of the Arabidopsis COP9 signalosome: implications in gene expression. Biochem. J. 2002, 366, 79–86. HOFMANN, K., BUCHER, P. The PCI domain: a common theme in three multiprotein complexes. Trends Biochem. Sci. 1998, 23, 204–205. KIM, T., HOFMANN, K., VON ARNIM, A. G., CHAMOVITZ, D. A. PCI complexes: pretty complex interactions in diverse signaling pathways. Trends Plant Sci. 2001, 6, 379–386. COPE, G. A., SUH, G. S., ARAVIND, L., SCHWARZ, S. E., ZIPURSKY, S. L., KOONIN, E. V., DESHAIES, R. J. Role of predicted metalloprotease motif of Jab1/Csn5 in cleavage of Nedd8 from Cul1. Science 2002, 298, 608–611. ZHOU, C., SEIBERT, V., GEYER, R., RHEE, E., LYAPINA, S., COPE, G., DESHAIES, R. J., WOLF, D. A. The ﬁssion yeast COP9/signalosome is involved in cullin modiﬁcation by ubiquitin-related Ned8p. BMC BioChem. 2001, 2, 7. VERMA, R., ARAVIND, L., OANIA, R., MCDONALD, W. H., YATES, J. R., 3rd,

63

64

65

66

67

68

69

70

KOONIN, E. V., DESHAIES, R. J. Role of Rpn11 metalloprotease in deubiquitination and degradation by the 26S proteasome. Science 2002, 298, 611–615. MAYTAL-KIVITY, V., REIS, N., HOFMANN, K., GLICKMAN, M. H. MPN+, a putative catalytic motif found in a subset of MPN domain proteins from eukaryotes and prokaryotes, is critical for Rpn11 function. BMC BioChem. 2002, 3, 28. TRAN, H. J., ALLEN, M. D., LOWE, J., BYCROFT, M. Structure of the Jab1/MPN domain and its implications for proteasome function. Biochemistry 2003, 42, 11460–11465. KAWAKAMI, T., CHIBA, T., SUZUKI, T., IWAI, K., YAMANAKA, K., MINATO, N., SUZUKI, H., SHIMBARA, N., HIDAKA, Y., OSAKA, F., OMATA, M., TANAKA, K. NEDD8 recruits E2-ubiquitin to SCF E3 ligase. EMBO J. 2001, 20, 4003–4012. PODUST, V. N., BROWNELL, J. E., GLADYSHEVA, T. B., LUO, R. S., WANG, C., COGGINS, M. B., PIERCE, J. W., LIGHTCAP, E. S., CHAU, V. A Nedd8 conjugation pathway is essential for proteolytic targeting of p27Kip1 by ubiquitination. Proc. Natl. Acad. Sci. USA 2000, 97, 4579–4584. READ, M. A., BROWNELL, J. E., GLADYSHEVA, T. B., HOTTELET, M., PARENT, L. A., COGGINS, M. B., PIERCE, J. W., PODUST, V. N., LUO, R. S., CHAU, V., PALOMBELLA, V. J. Nedd8 modiﬁcation of cul-1 activates SCF(beta(TrCP))-dependent ubiquitination of IkappaBalpha. Mol. Cell. Biol. 2000, 20, 2326–2333. NAUMANN, M., BECH-OTSCHIR, D., HUANG, X., FERRELL, K., DUBIEL, W. COP9 signalosome-directed c-Jun activation/stabilization is independent of JNK. J. Biol. Chem. 1999, 274, 35297–35300. KARNIOL, B., MALEC, P., CHAMOVITZ, D. A. Arabidopsis FUSCA5 encodes a novel phosphoprotein that is a component of the COP9 complex. Plant Cell 1999, 11, 839–848. WILSON, M. P., SUN, Y., CAO, L., MAJERUS, P. W. Inositol 1,3,4-trisphosphate 5/6-kinase is a

References

71

72

73

74

75

76

77

78

79

protein kinase that phosphorylates the transcription factors c-Jun and ATF-2. J. Biol. Chem. 2001, 276, 40998–41004. ZHOU, C., WEE, S., RHEE, E., NAUMANN, M., DUBIEL, W., WOLF, D. A. Fission yeast COP9/signalosome suppresses cullin activity through recruitment of the deubiquitylating enzyme Ubp12p. Mol. Cell 2003, 11, 927–938. WOLF, D. A., ZHOU, C., WEE, S. The COP9 signalosome: an assembly and maintenance platform for cullin ubiquitin ligases? Nat. Cell Biol. 2003, 5, 1029–1033. GEYER, R., WEE, S., ANDERSON, S., YATES, J., WOLF, D. A. BTB/POZ domain proteins are putative substrate adaptors for cullin 3 ubiquitin ligases. Mol. Cell 2003, 12, 783–790. TYERS, M., JORGENSEN, P. Proteolysis and the cell cycle: with this RING I do thee destroy. Curr. Opin. Genet. Dev. 2000, 10, 54–64. XU, L., WEI, Y., REBOUL, J., VAGLIO, P., SHIN, T. H., VIDAL, M., ELLEDGE, S. J., HARPER, J. W. BTB proteins are substrate-speciﬁc adaptors in an SCF-like modular ubiquitin ligase containing CUL-3. Nature 2003, 425, 316–321. PINTARD, L., WILLIS, J. H., WILLEMS, A., JOHNSON, J. L., SRAYKO, M., KURZ, T., GLASER, S., MAINS, P. E., TYERS, M., BOWERMAN, B., PETER, M. The BTB protein MEL-26 is a substrate-speciﬁc adaptor of the CUL-3 ubiquitin-ligase. Nature 2003, 425, 311–316. WERTZ, I. E., O’ROURKE, K. M., ZHANG, Z., DORNAN, D., ARNOTT, D., DESHAIES, R. J., DIXIT, V. M. Human De-etiolated-1 regulates c-Jun by assembling a CUL4A ubiquitin ligase. Science 2004, 303, 1371–1374. YAHALOM, A., KIM, T. H., WINTER, E., KARNIOL, B., VON ARNIM, A. G., CHAMOVITZ, D. A. Arabidopsis eIF3e (INT-6) associates with both eIF3c and the COP9 signalosome subunit CSN7. J. Biol. Chem. 2001, 276, 334–340. MAYTAL-KIVITY, V., PICK, E., PIRAN, R., HOFMANN, K., GLICKMAN, M. H. The COP9 signalosome-like complex in S. cerevisiae and links to other PCI

80

81

82

83

84

85

86

87

88

complexes. Int. J. Biochem. Cell Biol. 2003, 35, 706–715. SHALEV, A., VALASEK, L., PISE-MASISON, C. A., RADONOVICH, M., PHAN, L., CLAYTON, J., HE, H., BRADY, J. N., HINNEBUSCH, A. G., ASANO, K. Saccharomyces cerevisiae protein Pci8p and human protein eIF3e/Int-6 interact with the eIF3 core complex by binding to cognate eIF3b subunits. J. Biol. Chem. 2001, 276, 34948–34957. DUNAND-SAUTHIER, I., WALKER, C., WILKINSON, C., GORDON, C., CRANE, R., NORBURY, C., HUMPHREY, T. Sum1, a component of the ﬁssion yeast eIF3 translation initiation complex, is rapidly relocalized during environmental stress and interacts with components of the 26S proteasome. Mol. Biol. Cell 2002, 13, 1626–1640. YEN, H. C., GORDON, C., CHANG, E. C. Schizosaccharomyces pombe Int6 and Ras homologs regulate cell division and mitotic ﬁdelity via the proteasome. Cell 2003, 112, 207–217. VON ARNIM, A. G., CHAMOVITZ, D. A. Protein homeostasis: a degrading role for Int6/eIF3e. Curr. Biol. 2003, 13, R323–325. PENG, Z., SHEN, Y., FENG, S., WANG, X., CHITTETI, B. N., VIERSTRA, R. D., DENG, X. W. Evidence for a physical association of the COP9 signalosome, the proteasome, and speciﬁc SCF E3 ligases in vivo. Curr. Biol. 2003, 13, R504–505. LI, L., DENG, X. W. The COP9 signalosome: an alternative lid for the 26S proteasome? Trends Cell Biol. 2003, 13, 507–509. COPE, G. A., DESHAIES, R. J. COP9 signalosome: a multifunctional regulator of SCF and other cullin-based ubiquitin ligases. Cell 2003, 114, 663–671. ZHENG, J., YANG, X., HARRELL, J. M., RYZHIKOV, S., SHIM, E. H., LYKKE-ANDERSEN, K., WEI, N., SUN, H., KOBAYASHI, R., ZHANG, H. CAND1 binds to unneddylated CUL1 and regulates the formation of SCF ubiquitin E3 ligase complex. Mol. Cell 2002, 10, 1519–1526. XIE, Y., VARSHAVSKY, A. UFD4 lacking the proteasome-binding region catalyses ubiquitination but is impaired in

21

22 The COP9 Signalosome and Its Role in the Ubiquitin System

89

90

91

92

93

94

proteolysis. Nat Cell Biol. 2002, 4, 1003–1007. CORN, P. G., MCDONALD, E. R., 3rd, HERMAN, J. G., EL-DEIRY, W. S. Tat-binding protein-1, a component of the 26S proteasome, contributes to the E3 ubiquitin ligase function of the von Hippel-Lindau protein. Nat. Genet. 2003, 35, 229–237. DORONKIN, S., DJAGAEVA, I., BECKENDORF, S. K. CSN5/Jab1 mutations affect axis formation in the Drosophila oocyte by activating a meiotic checkpoint. Development 2002, 129, 5053–5064. HIGA, L. A., MIHAYLOV, I. S., BANKS, D. P., ZHENG, J., ZHANG, H. Radiation-mediated proteolysis of CDT1 by CUL4-ROC1 and CSN complexes constitutes a new checkpoint. Nat Cell Biol. 2003, 5, 1008–1015. WEE, S., HETFELD, B., DUBIEL, W., WOLF, D. A. Conservation of the COP9/signalosome in budding yeast. BMC Genet. 2002, 3, 15. ORON, E., MANNERVIK, M., RENCUS, S., HARARI-STEINBERG, O., NEUMAN-SILBERBERG, S., SEGAL, D., CHAMOVITZ, D. A. COP9 signalosome subunits 4 and 5 regulate multiple pleiotropic pathways in Drosophila melanogaster. Development 2002, 129, 4399–4409. SUH, G. S., POECK, B., CHOUARD, T., ORON, E., SEGAL, D., CHAMOVITZ, D. A., ZIPURSKY, S. L. Drosophila JAB1/CSN5

95

96

97

98

99

100

acts in photoreceptor cells to induce glial cells. Neuron 2002, 33, 35–46. MA, L., ZHAO, H., DENG, X. W. Analysis of the mutational effects of the COP/DET/FUS loci on genome expression proﬁles reveals their overlapping yet not identical roles in regulating Arabidopsis seedling development. Development 2003, 130, 969–981. SCHWECHHEIMER, C., SERINO, G., DENG, X. W. Multiple ubiquitin ligase-mediated processes require COP9 signalosome and AXR1 function. Plant Cell 2002, 14, 2553–2563. FENG, S., MA, L., WANG, X., XIE, D., DINESH-KUMAR, S. P., WEI, N., DENG, X. W. The COP9 signalosome interacts physically with SCF COI1 and modulates jasmonate responses. Plant Cell 2003, 15, 1083–1094. WANG, X., FENG, S., NAKAYAMA, N., CROSBY, W. L., IRISH, V., DENG, X. W., WEI, N. The COP9 signalosome interacts with SCF UFO and participates in Arabidopsis ﬂower development. Plant Cell 2003, 15, 1071–1082. ARBISER, J. L., KLAUBER, N., ROHAN, R., van LEEUWEN, R., HUANG, M. T., FISHER, C., FLYNN, E., BYERS, H. R. Curcumin is an in vivo inhibitor of angiogenesis. Mol. Med. 1998, 4, 376–383. BENEZRA, R., RAFII, S., LYDEN, D. The Id proteins and angiogenesis. Oncogene 2001, 20, 8334–8341.

1

Molecular Chaperones and the Ubiquitin–Proteasome System Cam Patterson

University of North Carolina, Chapel Hill, USA

J¨org H¨ohfeld Rheinische Friedrich-Wilhelms-Universit¨at Bonn, Bonn, Germany

Originally published in: Protein Degradation, Volume 2. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31130-0

1 Introduction

The biological activity of a protein is deﬁned by its unique three-dimensional structure. Attaining this structure, however, is a delicate process. A recent study suggests that up to 30% of all newly synthesized proteins never reach their native state [1]. As protein misfolding poses a major threat to cell function and viability, molecular mechanisms must have evolved to prevent the accumulation of misfolded proteins and thus aggregate formation. Two protective strategies appear to be followed. Molecular chaperones are employed to stabilize nonnative protein conformations and to promote folding to the native state whenever possible. Alternatively, misfolded proteins are removed by degradation, involving, for example, the ubiquitin–proteasome system. For a long time molecular chaperones and cellular degradation systems were therefore viewed as opposing forces. However, recent evidence suggests that certain chaperones (in particular members of the 70- and 90-kDa heat shock protein families) are able to cooperate with the ubiquitin–proteasome system. Protein fate thus appears to be determined by a tight interplay of cellular protein-folding and protein-degradation systems.

2 A Biomedical Perspective

The aggregation and accumulation of misfolded proteins is now recognized as a common characteristic of a number of degenerative disorders, many Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Molecular Chaperones and the Ubiquitin–Proteasome System

of which have neurological manifestations [2, 3]. These diseases include prionopathies, Alzheimer’s and Parkinson’s diseases, and polyglutamine expansion diseases such as Huntington’s disease and spinocerebellar ataxia. At the cellular level, these diseases are characterized by the accumulation of aberrant proteins either intracellularly or extracellularly in speciﬁc groups of cells that subsequently undergo death. The precise association between protein accumulation and cell death remains incompletely understood and may vary from disease to disease. In some cases, misfolded protein accumulations may themselves be toxic or exert spatial constraints on cells that affect their ability to function normally. In other cases, the sequestering of proteins in aggregates may itself be a protective mechanism, and it is the overwhelming of pathways that consolidate aberrant proteins that is the toxic event. In either case, lessons learned from genetically determined neurodegenerative diseases have helped us to understand the inciting events of protein aggregation that ultimately lead to degenerative diseases. Mutations resulting in neurodegenerative diseases fall into two broad classes. The ﬁrst class comprises mutations that affect proteins, irrespective of their native function, and cause them to misfold. The classic example of this is Huntington’s disease [4, 5]. The protein encoded by the huntingtin gene contains a stretch of glutamine residues (or polyglutamine repeat), and the genomic DNA sequence that codes for this polyglutamine repeat is subject to misreading and expansion. When the length of the polyglutamine repeat in huntingtin reaches a critical threshold of approximately 35 residues, the protein becomes prone to misfolding and aggregation [6]. This appears to be the proximate cause of neurotoxicity in this invariably fatal disease [7, 8]. A number of other neurodegenerative diseases are caused by polyglutamine expansions [9, 10]. For example, spinocerebellar ataxia is caused by polyglutamine expansions in the protein ataxin-1 [11]. In other diseases, protein misfolding occurs due to other mutations that induce misfolding and aggregation; for example, mutations in superoxide dismutase-1 lead to aggregation and neurotoxicity in amyotrophic lateral sclerosis [12, 13]. Other mutations that result in neurodegenerative diseases are instructive in that they directly implicate the ubiquitin–proteasome system in the pathogenesis of these diseases [14]. For example, mutations in the gene encoding the protein parkin are associated with juvenile-onset Parkinson’s disease [15, 16]. Parkin is a RING ﬁnger–containing ubiquitin ligase, and mutations in this ubiquitin ligase cause accumulation of target proteins that ultimately result in the neurotoxicity and motor dysfunction associated with Parkinson’s disease [17–20]. Repressor screens of neurodegeneration phenotypes in animal models have also linked the molecular chaperone machinery to neurodegeneration [21–24]. Taken together, the pathophysiology of neurodegenerative diseases provides a compelling demonstration of the importance of the regulated metabolism of misfolded proteins and provides direct evidence of the role of both molecular chaperones and the ubiquitin–proteasome system in guarding against protein misfolding and its consequent toxicity.

3 Molecular Chaperones: Mode of Action and Cellular Functions 3

3 Molecular Chaperones: Mode of Action and Cellular Functions

Molecular chaperones are deﬁned by their ability to bind and stabilize nonnative conformations of other proteins [25, 26]. Although they are an amazingly diverse group of conserved and ubiquitous proteins, they are also among the most abundant intracellular proteins. The classical function of chaperones is to facilitate protein folding, inhibit misfolding, and prevent aggregation. These folding events are regulated by interactions between chaperones and ancillary proteins, the co-chaperones, which in general assist in cycling unfolded substrate proteins on and off the active chaperone complex [25, 27, 28]. In agreement with their essential function under normal growth conditions, chaperones are ubiquitously expressed and are found in all cellular compartments of the eukaryotic cell (except for peroxisomes). In addition, cells greatly increase chaperone concentration as a response to diverse stresses, when proteins become unfolded and require protection and stabilization [29]. Accordingly, many chaperones are heat shock proteins (Hsps). Four main families of cytoplasmic chaperones can be distinguished: the Hsp70 family, the Hsp90 family, the small heat shock proteins, and the chaperonins. 3.1 The Hsp70 Family

The Hsp70 proteins bind to misfolded proteins promiscuously during translation or after stress-mediated protein damage [26, 30]. Members of this family are highly conserved throughout evolution and are found throughout the prokaryotic and eukaryotic phylogeny. It is common for a single cell to contain multiple homologues, even within a single cellular compartment; for example, mammalian cells express two inducible homologues (Hsp70.1 and Hsp70.3) and a constitutive homologue (Hsc70) in the cytoplasm. These homologues have overlapping but not totally redundant cellular functions. Members of this family are typically in the range of 70 kDa in size and contain three functional domains: an amino-terminal ATPase domain, a central peptide-binding cleft, and a carboxyl terminus that seems to form a lid over the peptide-binding cleft [28] (Figure 1). The chaperones recognize short segments of the protein substrate, which are composed of clusters of hydrophobic amino acids ﬂanked by basic residues [31]. Such binding motifs occur frequently within protein sequences and are found exposed on nonnative proteins. In fact, mammalian Hsp70 binds to a wide range of nascent and newly synthesized proteins, comprising about 15–20% of total protein [32]. This percentage is most likely further increased under stress conditions. Hsp70 proteins apparently prevent protein aggregation and promote proper folding by shielding hydrophobic segments of the protein substrate. The hydrophobic segments are recognized by the central peptide-binding domain of Hsp70 proteins (Figure 1). The domain is composed of two sheets of β strands that together with connecting loops form a cleft to accommodate extended peptides of about seven amino acids in length, as revealed

4 Molecular Chaperones and the Ubiquitin–Proteasome System

Fig. 1 Schematic presentation of the domain architecture and chaperone cycle of Hsp70. Hsp70 proteins display a characteristic domain structure comprising an amino-terminal ATPase domain (ATP), a peptide-binding domain (P), and a carboxylterminal domain (C) that is supposed to form a lid over the peptide-binding domain. In the ATP-bound conformation, the binding

pocket is open, resulting in a low afﬁnity for the binding of a chaperone substrate. ATP hydrolysis induces stable substrate binding through a closure of the peptide-binding pocket. Substrate release is induced upon nucleotide exchange. ATP hydrolysis and nucleotide exchange are regulated by diverse co-chaperones.

in crystallographic studies of bacterial Hsp70 [33]. In the obtained crystal structure, the adjacent carboxyl-terminal domain of Hsp70 folds back over the β sandwich, suggesting that the domain may function as a lid in permitting entry and release of protein substrates (Figure 1). According to this model, ATP binding and hydrolysis by the amino-terminal ATPase domain of Hsp70 induce conformational changes of the carboxyl terminus, which lead to lid opening and closure [28]. In the ATP-bound conformation of Hsp70, the peptide-binding pocket is open, resulting in rapid binding and release of the substrate and consequently in a low binding afﬁnity (Figure 1). Stable holding of the protein substrate requires closing of the binding pocket, which is induced upon ATP hydrolysis and conversion of Hsp70 to the ADP-bound conformation. The dynamic association of Hsp70 with nonnative polypeptide substrates thus depends on ongoing cycles of ATP binding, hydrolysis, and nucleotide exchange. Importantly, ancillary co-chaperones are employed to regulate the ATPase cycle [27, 30]. Co-chaperones of the Hsp40 family (also termed J proteins due to their founding member bacterial DnaJ) stimulate the ATP hydrolysis step within the Hsp70 reaction cycle and in this way promote substrate binding [34] (Figure 1). In contrast, the carboxyl terminus of Hsp70-interacting protein CHIP attenuates ATP hydrolysis [35]. Similarly, nucleotide exchange on Hsp70 is under the control of stimulating and inhibiting co-chaperones. The Hsp70-interacting protein Hip slows down nucleotide exchange by stabilizing the ADP-bound conformation of the chaperone [36], whereas nucleotide exchange is stimulated by the co-chaperone BAG-1 (Bcl-2-associated athanogene 1), which assists substrate unloading from Hsp70 [37–39]. By altering the AT-Pase cycle, the co-chaperones directly modulate the folding activity of Hsp70. In addition to chaperone-recognition motifs, co-chaperones often possess other functional

3 Molecular Chaperones: Mode of Action and Cellular Functions 5

Fig. 2 Domain architecture of diverse cochaperones of Hsp70. DnaJ: domain related to the bacterial co-chaperone DnaJ; TPR: tetratricopeptide repeat; Sti1: domain related to the yeast co-chaperone Sti1; CC: coiled-coil domain; U box: E2-interacting domain present in certain ubiquitin ligases;

PG: polyglycine region; ARM: armadillo repeat; TRSEEX: repeat motif found at the amino terminus of BAG-1 isoforms; ubl: ubiquitin-like domain; BAG: Hsp70-binding domain present in BAG proteins; WW: protein interaction domain.

domains and therefore link chaperone activity to distinct cellular processes [27, 40] (Figure 2). Indeed, as discussed below, the co-chaperones BAG-1 and CHIP apparently modulate Hsp70 function during protein degradation. 3.2 The Hsp90 Family

The 90-kDa cytoplasmic chaperones are members of the Hsp90 family, and in mammals two isoforms exist: Hsp90α and Hsp90β. The Hsp70 and Hsp90 families exhibit several common features: both possess ATPase activity and are regulated by ATP binding and hydrolysis, and both are further regulated by ancillary co-chaperones [41–48]. Unlike Hsp70, however, cytoplasmic Hsp90 is not generally involved in the folding of newly synthesized polypeptide chains. Instead it plays a key role in the regulation of signal transduction networks, as most of the known substrates of Hsp90 are signaling proteins, the classical examples being steroid hormone receptors and signaling kinases. On a molecular level, Hsp90 binds

6 Molecular Chaperones and the Ubiquitin–Proteasome System

Fig. 3 Cooperation of Hsp70 and Hsp90 during the regulation of signal transduction pathways. The inactive signaling protein, e.g., a steroid hormone receptor, is initially recognized by Hsp40 and delivered to Hsp70. Subsequently, a multi-chaperone complex assembles that contains the Hsp70 co-chaperone Hip and the Hsp70/Hsp90organizing protein Hop. Hop stimulates recruitment of an Hsp90 dimer that accepts the substrate from Hsp70. At the ﬁnal stage

of the chaperone pathway, Hsp90 associates with p23 and diverse cyclophilins (cycloph.) to mediate conformational changes of the signaling protein necessary to reach an activatable state. Upon activation, i.e., hormone binding in the case of the steroid receptor, the signaling protein is released from Hsp90. In the absence of an activating stimulus, the signaling protein folds back to the inactive state when released and enters a new cycle of chaperone binding.

to substrates at a late stage of the folding pathway, when the substrate is poised for activation by ligand binding or associations with other factors. Consequently, Hsp90 accepts partially folded conformations from Hsp70 for further processing. In the case of the chaperone-assisted activation of the glucocorticoid hormone receptor and also of the progesterone receptor, the sequence of events leading to attaining an active conformation is fairly well understood [49–53]. It appears that the receptors are initially recognized by Hsp40 and are then delivered to Hsp70 [54] (Figure 3). Subsequent transfer onto Hsp90 requires the Hsp70/Hsp90-organizing protein Hop, which possesses non-overlapping binding sites for Hsp70 and Hsp90 and therefore acts as a coupling factor between the two chaperones [55]. In conjunction with p23 and different cyclophilins, Hsp90 eventually mediates conformational changes that enable the receptor to reach a high-afﬁnity state for ligand binding. On other signaling pathways Hsp90 serves as a scaffolding factor to permit interactions between kinases and their substrates, as is the case for Akt kinase and endothelial nitric oxide synthase [56]. Since many of the Hsp90 substrate proteins are involved in regulating cell proliferation and cell death, it is not surprising that the chaperone recently emerged as a drug target in tumor therapy [57–59]. The antibiotics

3 Molecular Chaperones: Mode of Action and Cellular Functions 7

Fig. 4 Alteration of chaperone action during signal transduction induced by Hsp90 inhibitors such as geldanamycin and radicicol. In the presence of the inhibitors the activation pathway is blocked, and signaling

proteins are targeted to the proteasome for degradation in a process that involves the co-chaperone CHIP and other E3 ubiquitin ligases that remain to be identiﬁed.

geldanamycin and radicicol speciﬁcally bind to Hsp90 in mammalian cells and inhibit the function of the chaperone by occupying its ATP-binding pocket [60–63]. Drugs based on these compounds are now being developed as anticancer agents, as they potentially inactivate multiple signaling pathways that drive carcinogenesis. Remarkably, drug-induced inhibition of Hsp90 blocks the chaperone-assisted activation of signaling proteins and leads to their rapid degradation via the ubiquitin–proteasome pathway [64–69] (Figure 4). Hsp90 inhibitors therefore have emerged as helpful tools to study chaperone- proteasome cooperation. 3.3 The Small Heat Shock Proteins

The precise functions of small heat shock proteins (sHsps) including Hsp27 and the eye-lens protein αB-crystallin are incompletely understood. However, they seem to play a major role in preventing protein aggregation under conditions of cellular stress [70–73]. All members investigated so far form large oligomeric complexes of spherical or cylindrical appearance [74, 75]. Complex formation is independent of ATP binding and hydrolysis, but appears to be regulated by temperature and phosphorylation. The structural analysis of wheat Hsp16.9 suggested that the oligomeric complex acts as a storage form rather than an enclosure for substrates, as the active chaperone appears to be a dimer [75]. In agreement with this notion, dissociation of

8 Molecular Chaperones and the Ubiquitin–Proteasome System

the oligomeric complex formed by yeast Hsp26 was found to be a prerequisite for efﬁcient chaperone activity [76]. Subsequent refolding may occur spontaneously or may involve cooperation with other chaperones such as Hsp70 [77]. 3.4 Chaperonins

The chaperone proteins best understood with regard to their mode of action are certainly the so-called chaperonins, which are deﬁned by a barrel-shaped, doublering structure [25, 28]. Members include bacterial GroEL, Hsp60 of mitochondria and chloroplasts, and the TriC–CCT complex localized in the eukaryotic cytoplasm. Based on their characteristic ring structure, a central cavity is formed, which accommodates nonnative proteins via hydrophobic interactions. Conformational changes of the chaperonin subunits induced through ATP hydrolysis change the inner lining of the cavity from a hydrophobic to a hydrophilic character [78–80]. As a consequence the unfolded polypeptide is released into the central chamber and can proceed on its folding pathway in a protected environment [81]. The chaperonins are therefore capable of folding proteins such as actin that cannot be properly folded via other mechanisms [82].

4 Chaperones: Central Players During Protein Quality Control

Due to their ability to recognize nonnative conformations of other proteins, molecular chaperones are of central importance during protein quality control. This was elegantly revealed in studies on the inﬂuence of the Hsp70 chaperone system on polyglutamine diseases using the fruit ﬂy Drosophila melanogaster as a model organism (reviewed in Refs. [23] and [83]). Hallmarks of the polyglutamine disease spinocerebellar ataxia type 3 (SCA3), for example, were recapitulated in transgenic ﬂies that expressed a pathological polyQ tract of the ataxin-3 protein in the eye disc [84]. Transgene expression caused formation of abnormal protein inclusions and progressive neuronal degeneration. Intriguingly, co-expression of human cytoplasmic Hsp70 suppressed polyQ-induced neurotoxicity. In a similar experimental approach, Hsp40 family members protected neuronal cells against toxic polyQ expression [22]. Enhancing the activity of the Hsp70/Hsp40 chaperone system apparently mitigates cytotoxicity caused by the accumulation of aggregationprone proteins. These ﬁndings obtained in Drosophila were conﬁrmed in a mouse model of spinocerebellar ataxia type 1 (SCA1) [85, 86]. Unexpectedly, however, the Hsp70 chaperone system was unable to prevent the formation of protein aggregates in these models of polyglutamine diseases and upon polyQ expression in yeast and mammalian cells [84, 85, 87–89]. Elevating the cellular levels of Hsp70 and of some Hsp40 family members affected the number of protein aggregates and their biochemical properties, but did not inhibit the formation of polyQ aggregates. Notably, Hsp70 and Hsp40 profoundly modulated the aggregation process of

5 Chaperones and Protein Degradation

polyQ tracts in biochemical experiments; this led to the formation of amorphous, SDS-soluble aggregates, instead of the ordered, SDS-insoluble amyloid ﬁbrils that form in the absence of the chaperone system [88]. These biochemical data were conﬁrmed in yeast and mammalian cells [88, 90]. Although unable to prevent the formation of protein aggregates, the Hsp70 chaperone system apparently prevents the ordered oligomerization and ﬁbril growth that is characteristic of the disease process. In an alternate but not mutually exclusive model to explain their protective role, the chaperones may cover potentially dangerous surfaces exposed by polyQcontaining proteins during the oligomerization process or by the ﬁnal oligomers. Intriguingly, elevated expression of Hsp70 also suppresses the toxicity of the nonpolyQ-containing protein α-synuclein in a Drosophila model of Parkinson’s disease without inhibiting aggregate formation [24]. Hsp70 may thus exert a rather general function in protecting cells against toxic protein aggregation. This raises the exciting possibility that treatment of diverse forms of human neurodegenerative diseases may be achieved through upregulation of Hsp70 activity. The mentioned examples illustrate that one does not have to evoke the refolding of an aberrant protein to the native state in order to explain the protective activity of Hsp70 observed in models of amyloid diseases. In some cases it might be sufﬁcient for Hsp70 to modulate the aggregation process or to shield interaction surfaces of the misfolded protein to decrease cytotoxic effects. Another option may involve presentation of the misfolded protein to the ubiquitin–proteasome system for degradation.

5 Chaperones and Protein Degradation

Hsp70 and Hsp90 family members as well as small heat shock proteins have all been implicated to participate in protein degradation. For example, the small heat shock protein Hsp27 was recently shown to stimulate the degradation of phosphorylated IκBα via the ubiquitin–proteasome pathway, which may account for the antiapoptotic function of Hsp27 [91]. Similarly, Hsp27 facilitates the proteasomal degradation of phosphorylated tau, a microtubule-binding protein and component of protein deposits in Alzheimer’s disease [92]. Hsp70 participates in the degradation of apolipoprotein B100 (apoB), which is essential for the assembly and secretion of very low-density lipoproteins from the liver [93]. Under conditions of limited availability of core lipids, apoB translocation across the ER membrane is attenuated, resulting in the exposure of some domains of the protein into the cytoplasm and their recognition by Hsp70. This is followed by the degradation of apoB via the ubiquitin–proteasome pathway. Elevating cellular Hsp70 levels stimulated the degradation of the membrane protein, suggesting that the chaperone facilitates sorting to the proteasome. Genetic studies in yeast indicate that cytoplasmic Hsp70 may fulﬁll a rather general role in the degradation of ER-membrane proteins that display large domains into the cytoplasm [94]. In agreement with this notion, Hsp70 also takes part in the degradation of immaturely glycosylated and aberrantly

9

10 Molecular Chaperones and the Ubiquitin–Proteasome System

folded forms of the cystic ﬁbrosis transmembrane conductance regulator (CFTR) [95–98]. CFTR is an ion channel localized at the apical surface of epithelial cells. Its functional absence causes cystic ﬁbrosis, the most common fatal genetic disease in Caucasians [99, 100]. The disease-causing allele, F508, which is expressed in more than 70% of all patients, drastically interferes with the protein’s ability to fold, essentially barring it from functional expression in the plasma membrane. However, wild-type CFTR also folds very inefﬁciently, and less than 30% of the protein reaches the plasma membrane [99]. While trafﬁcking from the endoplasmic reticulum (ER) to the Golgi apparatus, immature forms of CFTR are recognized by quality-control systems and are eventually directed to the proteasome for degradation [101–104]. A critical step during CFTR biogenesis is the inefﬁcient folding of the ﬁrst of two cytoplasmically exposed nucleotide-binding domains (NBD1) of the membrane protein [105, 106]. The disease-causing F508 mutation localizes to NBD1 and further decreases the folding propensity of this domain. During the co-translational insertion of CFTR into the ER membrane, cytoplasmic Hsp70 and its co-chaperone Hdj-2 bind to NBD1 and facilitate intramolecular interactions between the domain and another cytoplasmic region of CFTR, the regulatory R-domain [96, 107]. However, Hsp70 is also able to present CFTR to the ubiquitin–proteasome system [97], and heterologous expression of CFTR in yeast revealed an essential role of cytoplasmic Hsp70 in CFTR turnover [98]. Hsp70 is thus a key player in the cellular surveillance system that monitors the folded state of CFTR at the ER membrane. Interestingly, CFTR and the disease form F508 are deposited in distinct pericentriolar structures, termed aggresomes, upon overexpression or proteasome inhibition [108]. Subsequent studies established that aggresomes are induced upon ectopic expression of many different aggregation-prone proteins (reviewed in Refs. [109] and [110]). Aggresomes form near the microtubule-organizing center in a manner dependent on the microtubule-associated motor protein dynein, and are surrounded by a “cage” of ﬁlamentous vimentin [108, 111]. Aggresome formation is apparently a speciﬁc and active cellular response when production of misfolded proteins exceeds the capacity of the ubiquitin–proteasome system to tag and remove these proteins. They likely serve to protect the cell from toxic “gain-of-function” activities acquired by misfolded proteins. Aggresomes are also of clinical relevance as they share remarkable biochemical and structural features, for example, with Lewy bodies, the cytoplasmic inclusion bodies found in neurons affected by Parkinson’s disease [112]. The pathways that regulate aggresome assembly are only now being explicated. Histone deacetylase 6 (HDAC6) appears to be a key regulator of aggresome assembly [113]. HDAC6 is a microtubule-associated deacetylase that has the capacity to bind both multi-ubiquitinated proteins and dynein motors and is believed to recruit misfolded proteins to the pericentriolar region for aggresome assembly. Deletion of HDAC6 prevents aggresome formation and sensitizes cells to the toxic effects of misfolded proteins, which supports the hypothesis that aggresomes sequester misfolded proteins to protect against their toxic activities. Components of the ubiquitin–proteasome system and chaperones such as Hsp70 are abundantly present in and are actively recruited to aggresomes [114–116]. Furthermore, elevating cellular Hsp70 levels can reduce aggresome formation by

5 Chaperones and Protein Degradation

stimulating proteasomal degradation [117]. It appears that these subcellular structures are major sites of chaperone – proteasome cooperation to mediate the metabolism of misfolded proteins. The formation of aggresome-like structures is also observed in dendritic cells that present foreign antigens to other immune cells [118]. Immature dendritic cells are located in tissues throughout the body, including skin and gut. When they encounter invading microbes, the pathogens are endocytosed and processed in a manner that involves the generation of antigenic peptides by the ubiquitin–proteasome system. Upon induction of dendritic cell maturation, ubiquitinated proteins transiently accumulate in large cytosolic structures that resemble aggresomes and were therefore termed DALIS (dendritic cell aggresome-like induced structures). It was speculated that DALIS formation may enable dendritic cells to regulate antigen processing and presentation. DALIS contain components of the ubiquitin–proteasome machinery as well as Hsp70 and the co-chaperone CHIP [118, 119]. Again, an interplay of molecular chaperones and the ubiquitin–proteasome system during regulated protein turnover is suggested. The cellular function of molecular chaperones is apparently not restricted to mediating protein folding; instead, chaperones emerge also as vital components on protein-degradation pathways. Remarkably, the balance between folding and degradation activities of chaperones can be manipulated. In cells treated with Hsp90 inhibitors, for example, with geldanamycin (see above), the chaperone-assisted activation of signaling proteins is abrogated and chaperone substrates such as the protein kinases Raf-1 and ErbB2 are rapidly degraded by the ubiquitin–proteasome system [64–69, 120]. This appears to be due, in part, to transfer of the substrates back to Hsp70 and progression toward the ubiquitin-dependent degradation pathway. Substrate interactions with chaperones – and consequently their commitment either toward the folding pathway or to their degradation via the ubiquitin– proteasome machinery – apparently serve as an essential post-translational protein quality-control mechanism within eukaryotic cells. The partitioning of proteins to either one of these mutually exclusive pathways is referred to as “protein triage” [121]. Although some misfolded proteins may be directly recognized by the proteasome [122], speciﬁc pathways within the ubiquitin–proteasome system are probably relied on for the degradation of most misfolded and damaged proteins. For example, E2 enzymes of the Ubc4/5 family selectively mediate the ubiquitylation of abnormal proteins as revealed in genetic studies in Saccharomyces cerevisiae [123]. It is well accepted that chaperones play a central role in the triage decision; however, less well understood are the events that lead to the cessation of efforts to fold a substrate, and the diversion of the substrate to the terminal degradative pathway. It is possible that chaperones and components of the ubiquitin–proteasome pathway exist in a state of competition for these substrates and that repeated cycling of a substrate on and off a chaperone maintains the substrate in a soluble state and increases, in a stochastic fashion, its likelihood of interactions with the ubiquitin machinery (Figure 5A). However, some data argue for a more direct role of the chaperones in the degradation process. Hsp70 plays an active and necessary role in the ubiquitylation of some substrates [124]; this activity of Hsp70

11

12 Molecular Chaperones and the Ubiquitin–Proteasome System

Fig. 5 Interplay of molecular chaperones with the ubiquitin–proteasome system. (A) Chaperones and the degradation machinery (i.e., ubiquitylation systems) compete with each other in the recognition of folding intermediates. Interaction with the chaperones directs the substrate towards folding. However, when the protein substrate is unable to attain a folded conformation, the chaperones maintain the folding intermediate in a soluble state that can be recog-

nized by the degradation machinery. (B) The chaperones are actively involved in protein degradation. Through an association with certain components of the ubiquitin conjugation machinery (degrading partner), the chaperones participate in the targeting of protein substrates to the proteasome. A competition between degrading partners and folding partners determines chaperone action and the fate of the protein substrate.

requires its chaperone function, indicating that conformational changes within substrates may facilitate recognition by the ubiquitylation machinery. Plausible hypotheses to explain these observations include direct associations between the chaperone and ubiquitin–proteasome machinery to facilitate transfer of a substrate from one pathway to the other, or conversion of the chaperone itself to a ubiquitylation complex (Figure 5B). It is also entirely possible that several quality-control pathways may exist and that the endogenous triage decision may involve aspects of each of these hypotheses.

6 The CHIP Ubiquitin Ligase: A Link Between Folding and Degradation Systems

Major insights into molecular mechanisms that underlie the cooperation of molecular chaperones with the ubiquitin–proteasome system were obtained through the functional characterization of the co-chaperone CHIP (reviewed in Ref. [40]). CHIP was initially identiﬁed in a screen for proteins containing tetratricopeptide repeat (TPR) domains, which are found in several co-chaperones – including Hip, Hop, and the cyclophilins – as chaperone-binding domains [27, 55] (Figure 2). CHIP

6 The CHIP Ubiquitin Ligase: A Link Between Folding and Degradation Systems 13

contains three TPR domains at its amino terminus, which are used for binding to Hsp70 and Hsp90 [35, 125]. Besides the TPR domains, CHIP possesses a Ubox domain at its carboxyl terminus [35] (Figure 2). U-box domains are similar to RING ﬁnger domains, but they lack the metal-chelating residues and instead are structured by intramolecular interactions [126]. The predicted structural similarity suggests that U boxes, like RING ﬁngers, may also play a role in targeting proteins for ubiquitylation and subsequent proteasome-dependent degradation, and this possibility is borne out in functional analyses of U box–containing proteins [127, 128]. The TPR and U-box domains in CHIP are separated by a central domain rich in charged residues. The charged domain of CHIP is necessary for TPRdependent interactions with Hsp70 [35] and is also required for homodimerization of CHIP [129]. The tissue distribution of CHIP supports the notion that it participates in protein folding and degradation decisions, as it is most highly expressed in tissues with high metabolic activity and protein turnover: skeletal muscle, heart, and brain. Although it is also present in all other organs, including pancreas, lung, liver, placenta, and kidney, the expression levels are much lower. CHIP is also detectable in most cultured cells, and is particularly abundant in muscle and neuronal cells and in tumor-derived cell lines [35]. Intracellularly, CHIP is primarily localized to the cytoplasm under quiescent conditions [35], although a fraction of CHIP is present in the nucleus [97]. In addition, cytoplasmic CHIP trafﬁcs into the nucleus in response to environmental challenge in cultured cells, which may serve as a protective mechanism or to regulate transcriptional responses in the setting of stress [130]. CHIP is distinguished among co-chaperones in that it is a bona ﬁde interaction partner with both of the major cytoplasmic chaperones Hsp90 and Hsp70, based on their interactions with CHIP in the yeast two-hybrid system and in vivo binding assays [35, 125]. CHIP interacts with the terminal-terminal EEVD motifs of Hsp70 and Hsp90, similar to other TPR domain–containing co-chaperones such as Hop [55, 131, 132]. When bound to Hsp70, CHIP inhibits ATP hydrolysis and therefore attenuates substrate binding and refolding, resulting in inhibition of the “forward” Hsp70 substrate folding/refolding pathway, at least in in vitro assays [35]. The cellular consequences of this “anti-chaperone” function are not yet clear, and in fact CHIP may actually facilitate protein folding under conditions of stress, perhaps by slowing the Hsc70 reaction cycle [130, 133]. CHIP interacts with Hsp90 with approximately equivalent afﬁnity to its interaction with Hsp70 [125]. This interaction results in remodeling of Hsp90 chaperone complexes, such that the co-chaperone p23, which is required for the appropriate activation of many, if not all, Hsp90 client proteins, is excluded. The mechanism for this activity is unclear – p23 and CHIP bind Hsp90 through different sites – yet the consequence of this action is predictable: CHIP should inhibit the function of proteins that require Hsp90 for conformational activation. The glucocorticoid receptor is an Hsp90 client that undergoes activation through a well-described sequence of events that depend on interactions of the glucocorticoid receptor with Hsp90 and various Hsp90 co-chaperones, including p23, making it an excellent model to test this prediction. Indeed, CHIP inhibits glucocorticoid receptor substrate binding and

14 Molecular Chaperones and the Ubiquitin–Proteasome System

steroid-dependent transactivation ability [125]. Surprisingly, this effect of CHIP is accompanied by decreased steady-state levels of glucocorticoid receptor, and CHIP induces ubiquitylation of the glucocorticoid receptor in vivo and in vitro, as well as subsequent proteasome-dependent degradation. This effect is both U-box- and TPR-domain-dependent, suggesting that CHIP’s effects on GR require direct interaction with Hsp90 and direct ubiquitylation of GR and delivery to the proteasome. These observations are not limited to the glucocorticoid receptor. ErbB2, another Hsp90 client, is also degraded by CHIP in a proteasome-dependent fashion [120]. Nor are they limited to Hsp90 clients. For example, CHIP cooperates with Hsp70 during the degradation of immature forms of the CFTR protein at the ER membrane and during the ubiquitylation of phosphorylated forms of the microtubule-binding protein tau, which is of clinical importance due to its role in the pathology of Alzheimer’s disease [97, 134]. The effects of CHIP are dependent on both the TPR domain, indicating a necessity for interactions with molecular chaperones, and the U box, which suggests that the U box is most likely the “business end” with respect to ubiquitylation. The means by which CHIP-dependent ubiquitylation occurs is not clear. In the case of ErbB2, ubiquitylation depends on a transfer of the client protein from Hsp90 to Hsp70 [120], indicating that the ﬁnal ubiquitylation complex consists of CHIP, Hsp70 (but not Hsp90), and the client protein. In any event, the studies are consistent in supporting a role for CHIP as a key component of the chaperone-dependent quality-control mechanism. CHIP efﬁciently targets client proteins, particularly when they are partially unfolded (as is the case for most Hsp90 clients when bound to the chaperone) or frankly misfolded (as is the case for most proteins binding to Hsp70 through exposed hydrophobic residues). Once the ubiquitylation activity of CHIP was recognized, it was logical to speculate that its U box might function in a manner analogous to that of RING ﬁngers, which have recently been appreciated as key components of the largest family of ubiquitin ligases. If CHIP is a ubiquitin ligase, then its ability to ubiquitylate a substrate should be reconstituted in vitro when a substrate is added in the presence of CHIP, E1, an E2, and ubiquitin. Indeed, this is the case [135–137] (Figure 6). CHIP is thus the ﬁrst described chaperone-associated E3 ligase. The ubiquitin ligase activity of CHIP depends on functional and physical interactions with a speciﬁc family of E2 enzymes, the Ubc4/Ubc5 family, which in humans comprises the E2s UbcH5a, UbcH5b, and UbcH5c. Of interest is the fact that the Ubc4/Ubc5 E2s are stress-activated, ubiquitin-conjugating enzymes [123]. CHIP can therefore be seen as a co-chaperone that, in addition to inhibiting traditional chaperone activity, converts chaperone complexes into chaperone-dependent ubiquitin ligases. Indeed, the chaperones themselves seem to act as the main substrate-recognition components of these ubiquitin ligase complexes, as efﬁcient ubiquitylation of chaperone substrates by CHIP depends on the presence of Hsp70 or Hsp90 in reconstituted systems [136, 137] (Figure 6). The chaperones apparently function in a manner analogous to F-box proteins, which are required as substrate recognition modules in many RING ﬁnger–containing ubiquitin ligase complexes [138–140]. Recently, another surprising function for CHIP has been identiﬁed, that of activation of the stress-responsive transcription factor heat shock factor-1 (HSF1)

7 Other Proteins That May Inﬂuence the Balance

Fig. 6 Characterization of CHIP as a chaperone-associated ubiquitin ligase. Puriﬁed CHIP, UbcH5b, the ubiquitinactivating enzyme E1, ubiquitin, and the Hsp70–Hsp40 chaperone system were incubated with the bacterially expressed protein kinase Raf-1 (for details, see Ref. [137]). Raf-1 and ubiquitylated forms of the kinase

(ub(n) -Raf-1) were detected by immunoblotting using a speciﬁc anti-Raf-1 antibody. Efﬁcient ubiquitylation of Raf-1 through the CHIP conjugation machinery depends on the recognition of the chaperone substrate by Hsp70, which presents the kinase to the conjugation machinery.

[130]. Through this association, CHIP regulates the expression of chaperones such as Hsp70 independently of its ability to modify their function through direct interactions. The mechanisms through which CHIP activates HSF1 are not entirely clear, but they are dependent in part on the induction of HSF1 trimerization, which is required for nuclear import and DNA binding. In addition, activation of HSF1 by CHIP seems to be independent of CHIP’s ubiquitin ligase activity. The consequences of this activation are important for the response to stress, in that cells lacking CHIP are prone to stress-dependent apoptosis and mice deﬁcient in CHIP (through homologous recombination) succumb rapidly to thermal challenge. These data indicate that CHIP plays a heretofore unsuspected role in coordinating the response to stress, not only by serving as a rate-limiting step in the degradation of damaged proteins but also by increasing the buffering capacity of the chaperone system to guard against stress-dependent proteotoxicity.

7 Other Proteins That May Inﬂuence the Balance Between Chaperone-assisted Folding and Degradation

CHIP is ideally suited to mediate chaperone-proteasome cooperation, as it combines a chaperone-binding motif and a domain that functions in ubiquitindependent degradation within its protein structure (Figure 2). Some other cochaperones display a similar structural arrangement [40]. For example, BAG-1 contacts Hsp70 through a BAG-domain located at its carboxyl terminus and, in

15

16 Molecular Chaperones and the Ubiquitin–Proteasome System

Fig. 7 Schematic presentation of the BAG1–Hsp70–CHIP complex. BAG-1 associates with the ATPase domain of Hsp70, while CHIP is bound to the carboxyl terminus. BAG-1 mediates an association of Hsp70 with the proteasome via its ubiquitin-like

domain (ubl), whereas CHIP acts in conjunction with Ubc4/5 as a chaperoneassociated ubiquitin ligase to mediate the attachment of a polyubiquitin chain to the chaperone substrate.

addition, possesses a central ubiquitin-like domain that is used for binding to the proteasome [141] (Figure 2). The co-chaperone thus belongs to a family of ubiquitin domain proteins (UDPs), many of which were shown to be associated with the proteasome [142]. This domain architecture enables BAG-1 to provide a physical link between Hsp70 and the proteasome, and elevating the cellular levels of BAG-1 results in a recruitment of the chaperone to the proteolytic complex. Notably, BAG-1 and CHIP occupy distinct domains on Hsp70 (Figure 7). Whereas BAG-1 associates with the amino-terminal ATPase domain, CHIP binds to the carboxylterminal EEVD motif of Hsp70 [35, 37]. Ternary complexes that comprise both co-chaperones associated with Hsp70 can be isolated from mammalian cells, suggesting a cooperation of BAG-1 and CHIP in the regulation of Hsp70 activity on certain degradation pathways. In fact, BAG-1 is able to stimulate the CHIPinduced degradation of the glucocorticoid hormone receptor [137]. A cooperation of diverse co-chaperones apparently provides additional levels of regulation to alter chaperone-assisted folding and degradation pathways. Interestingly, BAG-1 and also Hsp70 and Hsp90 are themselves substrates of the CHIP ubiquitin ligase [135, 143] (J.H. unpublished). Yet, CHIP-mediated ubiquitylation of the chaperones and the co-chaperone does not induce their proteasomal degradation. Instead, it seems to provide additional means to regulate the association of the chaperone systems with the proteasome. In the case of BAG-1, ubiquitylation mediated by CHIP indeed stimulates the binding of the co-chaperone to the proteasome [143]. It remains to be elucidated, however, why Hsp70 and BAG-1 are not degraded when sorted to the proteasome through CHIP-induced ubiquitylation, in contrast to chaperone substrates such as the glucocorticoid hormone receptor. Possibly, the folded state of the proteins may serve to distinguish targeting factors and substrates doomed for degradation. Efﬁcient ubiquitylation of BAG-1 mediated by CHIP is dependent on the formation of the ternary BAG-1–Hsp70–CHIP complex [143]. The formed chaperone complex would thus expose multiple signals for sorting to the proteasome, e.g., the integrated ubiquitin-like domain of BAG-1 and polyubiquitin chains attached to BAG-1, Hsp70, and the bound protein substrate. Such a redundancy of sorting information might be considered unnecessary. Intriguingly, however, several

7 Other Proteins That May Inﬂuence the Balance

subunits of the regulatory 19S particle of the proteasome are currently thought to act as receptors for polyubiquitin chains and integrated ubiquitin-like domains, including Rpn1, Rpn2, Rpt5, and Rpn10. The Rpn10 subunit was initially identiﬁed as a polyubiquitin chain receptor and was later shown to also bind integrated ubiquitin-like domains presented by UDPs [144–146]. Rpn10 possesses two distinct ubiquitin-binding domains, of which only one is used for UDP recognition [145–147]. However, conﬂicting data exist as to whether the subunit acts as a ubiquitin receptor in the context of the assembled 19S complex [148, 149]. More recently, Rpn1 was identiﬁed as a receptor for integrated ubiquitin-like domains [149], and a similar function may be fulﬁlled by the Rpn1-related subunit Rpn2 [150]. Polyubiquitin chains seem to be recognized by the Rpt5 subunit, one of the AAA ATPases present in the ring-like base of the regulatory 19S complex [151]. Its receptor function was revealed when tetraubiquitin was cross-linked to intact proteasomes [148]. Multiple docking sites for ubiquitin-like domains and polyubiquitin chains are apparently displayed by the regulatory particle of the proteasome. This may provide a structural basis for the recognition of multiple sorting signals exposed by the CHIP–chaperone complex (Figure 8). A similar mechanism involving multiple-site binding at the proteasome was recently proposed based on the

Fig. 8 The co-chaperone network that determines folding and degradation activities of Hsp70. BAG-1 and CHIP associate with Hsp70 to induce the proteasomal degradation of a Hsp70-bound protein substrate. When BAG-1 is displaced by binding of HspBP1 to the ATPase domain of Hsp70, the ubiquitin ligase activity of CHIP is attenuated in the formed complex, enabling

CHIP to modulate Hsp70 activity without inducing degradation. The ATPase domain can also be occupied by Hip, which stimulates the chaperone activity of Hsp70 and participates in the Hsp70/Hsp90-mediated regulation of signal transduction pathways. At the same time, Hop displaces CHIP from the carboxyl terminus of Hsp70 and recruits Hsp90 to the chaperone complex.

17

18 Molecular Chaperones and the Ubiquitin–Proteasome System

observation that two unrelated yeast ubiquitin ligases associate with speciﬁc subunits of the 19S regulatory complex [152]. In these cases substrate delivery involves interactions of proteasomal subunits with the substrate-bound ubiquitin ligase, with the polyubiquitin chain attached to the substrate, and with the substrate itself. Multiple-site binding may function to slow down dissociation of the substrate from the proteasome and to facilitate transfer into the central proteolytic chamber through ATP-dependent movements of the subunits of the 19S particle. Human cells contain several BAG-1-related proteins: BAG-2, BAG-3 (CAIR-1; Bis), BAG-4 (SODD), BAG-5, and BAG-6 (Scythe, BAT3) [153] (Figure 2). It appears that BAG proteins act as nucleotide-exchange factors to induce substrate unloading from Hsp70 on diverse protein folding, assembly, and degradation pathways. Notably, BAG-6 is another likely candidate for a co-chaperone that regulates protein degradation via the ubiquitin–proteasome pathway. Similar to BAG-1, BAG-6 contains a ubiquitin-like domain that is possibly used for proteasome binding [154]. However, experimental data verifying a role of BAG-6 in protein degradation remain elusive so far. The cooperation of diverse co-chaperones not only may allow promotion of chaperone-associated degradation but also may provide the means to conﬁne the destructive activity of CHIP. The Hsp70-binding protein 1 (HspBP1) seems to fulﬁll such a regulatory function [155]. HspBP1 was initially identiﬁed in a screen for proteins that associate with the ATPase domain of Hsp70 and was shown to stimulate nucleotide release from the chaperone [156, 157]. Notably, association of HspBP1 with the ATPase domain blocks binding of BAG-1 to Hsp70 and at the same time promotes an interaction of CHIP with Hsp70’s carboxyl terminus. In the formed ternary HspBP1–Hsp70–CHIP complex, the ubiquitin ligase activity of CHIP is attenuated and Hsp70 as well as a chaperone substrate are no longer efﬁciently ubiquitylated [155]. By interfering with CHIP-mediated ubiquitylation, HspBP1 stimulates the maturation of CFTR and promotes the sorting of the membrane protein to the cell surface. HspBP1 apparently functions as an antagonist of the CHIP ubiquitin ligase to regulate Hsp70-assisted folding and degradation pathways (Figure 8). The HspBP1-mediated inhibition of the ubiquitin ligase activity may enable CHIP to modulate the Hsp70 ATPase cycle without inducing degradation. In fact, degradation-independent functions of CHIP have recently emerged [130, 133, 158, 159]. CHIP was shown to regulate the chaperone-assisted folding and sorting of the androgen receptor and of endothelial nitric oxide synthase without inducing degradation [158, 159]. Moreover, CHIP fulﬁlls an essential role in the chaperone-mediated regulation of the heat shock transcription factor, independent of its degradation-inducing activity [130]. It remains to be seen, however, whether HspBP1 cooperates with CHIP in these instances, as HspBP1 displayed a certain speciﬁcity with regard to chaperone substrates. The co-chaperone interfered with the degradation of CFTR, but did not inﬂuence the CHIP-mediated turnover of the glucocorticoid hormone receptor. Such a client speciﬁcity may arise in part from the fact that HspBP1 inhibits the ubiquitin ligase activity of CHIP in a complex with Hsc70, but leaves Hsp90-associated

8 Further Considerations

ubiquitylation unaffected [155]. In addition, direct interactions between HspBP1 and a subset of chaperone substrates may contribute to substrate selection. In any case, the cooperation of CHIP with other co-chaperones apparently provides a means to regulate chaperone-assisted protein degradation. It is likely that there are multiple degradation pathways for misfolded proteins in the eukaryotic cytoplasm. Although CHIP participates in the degradation of chaperone substrates induced by applying Hsp90 inhibitors to cell cultures (see above), drug-induced degradation is not abrogated in cells that lack the CHIP ubiquitin ligase [120]. Furthermore, CHIP cooperates with Hsp70 in the ER-associated degradation of CFTR, but the Hsp70-assisted degradation of apoB at the cytoplasmic face of the ER membrane does not involve CHIP [97]. Taken together, these data strongly argue for the existence of other, yet to be identiﬁed, ubiquitin ligases that are able to target chaperone substrates to the proteasome. A likely candidate in this regard is Parkin, a RING ﬁnger ubiquitin ligase, whose activity is impaired in juvenile forms of Parkinson’s disease [17]. Hsp70 and CHIP were found to be associated with Parkin in neuronal cells, suggesting an involvement of Parkin in the proteasomal degradation of chaperone substrates [160]. Interestingly, α-synuclein, the main component of protein deposits observed in dopaminergic neurons of Parkinson patients, and synphilin, a protein that binds α-synuclein and induces deposit formation, both associate with yet other ubiquitin ligases: Siah-1 and Siah-2 [161, 162]. In the case of Siah-1, a link to cytoplasmic chaperone systems is suggested by the ﬁnding that the Hsp70 co-chaperone BAG-1 is a binding partner of the ubiquitin ligase and suppresses some of the cellular activities of Siah-1 [163]. Taken together, it is tempting to speculate about a role of Parkin and Siah on chaperone-assisted degradation pathways; yet, this remains to be explored in detail.

8 Further Considerations

Although the appreciation of interplay between molecular chaperones and ubiquitin-dependent proteolysis has greatly expanded over the past decade, a number of critical issues remain to be resolved. It is not entirely clear what determines whether a misfolded protein will undergo repeated attempts at misfolding versus diversion to the ubiquitin–proteasome pathway. Recruitment of CHIP into chaperone complexes appears to be a critical component of this reaction, which therefore begs the question as to what regulates this step. Since this step in protein quality control must be both rapidly activated and easily reversible, it is likely that regulation occurs at the post-translational level rather than through changes in steady-state protein levels. The precise sorting mechanisms for ubiquitinated proteins are also unclear. BAG-1 is a player, and it is also likely that overlap exists to some extent for sorting of the cytoplasmic and endoplasmic reticulum quality-control pathways. Nevertheless, much remains to be learned about these steps. From a broader perspective, it is now also imperative to understand the pathophysiological roles of cytoplasmic quality-control mechanisms regulated by

19

20 Molecular Chaperones and the Ubiquitin–Proteasome System

chaperone-proteasome interactions. As mentioned previously, there is a strong association between chaperone dysfunction and accumulations of misfolded proteins that characterizes genetic neurodegenerative diseases. An imbalance between protein folding and degradation may also contribute to some features of senescence and organismal aging. The link between chaperone systems and aging is based on increasing appreciation that modiﬁed, misfolded, and aggregated proteins accumulate with age [164]. Dysregulation of chaperone expression has been observed with aging and is therefore implicated in aging-related changes [165]; in general, it is accepted that induction of the major chaperones is impaired with aging, a fact conﬁrmed by recent gene-proﬁling experiments in vivo [166], although given the diversity of chaperones it is probably not surprising that age-related changes in expression are fairly complicated [167]. The mechanism underlying this dysregulation is not entirely clear, but seems to be due in part to impaired activation of the stress-responsive transcription factor HSF1. Overexpression of heat shock proteins in yeast, C. elegans, and Drosophila leads to increased longevity [168–170]. More recently, conclusive genetic evidence from C. elegans indicates that mutation of HSF1 causes a dramatic and signiﬁcant reduction in lifespan [170, 171], further implicating the accumulation of misfolded proteins in age-related phenotypes.

9 Conclusions

The associations between molecular chaperones and the ubiquitin–proteasome system represent a critical step in the response to proteotoxic damage. Whether attempts should be made to refold damaged proteins (thus conserving cellular resources) or degrade them instead (to prevent the possibility of protein aggregation and concomitant toxicity) requires a consideration of cellular economy. Defects in the quality-control mechanisms may have enormous consequences even if only slight imbalances occur between protein folding and degradation, as these imbalances can cause accumulated toxicity over time. The relationship between chaperone–proteasome interactions and pathophysiological events is only now being unraveled. Modulation of this system may provide a unique therapeutic target for degenerative diseases and pathologies associated with aging. References 1 U. SCHUBERT, L. C. ANTON, J. GIBBS, C. C. NORBURY, J. W. YEWDELL, J. R. BENNINK, Rapid degradation of a large fraction of newly synthesized proteins by proteasomes, Nature 404 (2000) 770–774. 2 J. P. TAYLOR, J. HARDY, K. H. FISCHBECK, Toxic proteins in neurodegenerative disease, Science 296 (2002) 1991–1995.

3 A. HORWICH, Protein aggregation in disease: a role for folding intermediates forming speciﬁc multimeric interactions, J. Clin. Invest. 110 (2002) 1221–1232. 4 E. SCHERZINGER, R. LURZ, M. TURMAINE, L. MANGIARINI, B. HOLLENBACH, R. HASENBANK, G. P. BATES, S. W. DAVIES, H. LEHRACH, E. E. WANKER,

References

5

6

7

8

9

10

11

12

Huntingtin-encoded polyglutamine expansions form amyloid-like protein aggregates in vitro and in vivo, Cell 90 (1997) 549–558. E. SCHERZINGER, A. SITTLER, K. SCHWEIGER, V. HEISER, R. LURZ, R. HASENBANK, G. P. BATES, H. LEHRACH, E. E. WANKER, Self-assembly of polyglutamine-containing huntingtin fragments into amyloid-like ﬁbrils: implications for Huntington’s disease pathology, Proc. Natl. Acad. Sci. USA 96 (1999) 4604–4609. A. KAZANTSEV, E. PREISINGER, A. DRANOVSKY, D. GOLDGABER, D. HOUSMAN, Insoluble detergent-resistant aggregates form between pathological and nonpathological lengths of polyglutamine in mammalian cells, Proc. Natl. Acad. Sci. USA 96 (1999) 11404–11409. D. MARTINDALE, A. HACKAM, A. WIECZOREK, L. ELLERBY, C. WELLINGTON, K. MCCUTCHEON, R. SINGARAJA, P. KAZEMI-ESFARJANI, R. DEVON, S. U. KIM, D. E. BREDESEN, F. TUFARO, M. R. HAYDEN, Length of huntingtin and its polyglutamine tract inﬂuences localization and frequency of intracellular aggregates. Nat. Genet. 18 (1998) 150–154. I. SANCHEZ, C. MAHLKE, J. YUAN, Pivotal role of oligomerization in expanded polyglutamine neurodegenerative disorders, Nature 421 (2003) 373–379. M. F. PERUTZ, Glutamine repeats and neurodegenerative diseases: molecular aspects, Trends Biochem. Sci. 24 (1999) 58–63. H. Y. ZOGHBI, H. T. ORR, Glutamine repeats and neurodegeneration, Annu. Rev. Neurosci. 23 (2000) 217–247. H. T. ORR, M. Y. CHUNG, S. BANFI, T. J. Jr. KWIATKOWSKI, A. SERVADIO, A. L. BEAUDET, A. E. MCCALL, L. A. DUVICK, L. P. RANUM, H. Y. ZOGHBI, Expansion of an unstable trinucleotide CAG repeat in spinocerebellar ataxia type 1, Nat. Genet. 4 (1993) 221–226. D. R. ROSEN, T. SIDDIQUE, D. PATTERSON, D. A. FIGLEWICZ, P. SAPP, A. HENTATI, D. DONALDSON, J. GOTO, J. P. O’REGAN, H. X. DENG, et al. Mutations in Cu/Zn

13

14

15

16

17

18

19

20

superoxide dismutase gene are associated with familial amyotrophic lateral sclerosis. Nature 362 (1993) 59–62. J. A. JOHNSTON, M. J. DALTON, M. E. GURNEY, R. R. KOPITO, Formation of high molecular weight complexes of mutant Cu, Zn-superoxide dismutase in a mouse model for familial amyotrophic lateral sclerosis, Proc. Natl. Acad. Sci. USA 97 (2000) 12571–12576. B. I. GIASSON, V. M. LEE, Are ubiquitination pathways central to Parkinson’s disease? Cell 114 (2003) 1–8. T. KITADA, S. ASAKAWA, N. HATTORI, H. MATSUMINE, Y. YAMAMURA, S. MINOSHIMA, M. YOKOCHI, Y. MIZUNO, N. SHIMIZU, Mutations in the parkin gene cause autosomal recessive juvenile parkinsonism, Nature 392 (1998) 605–608. C. B. L¨UCKING, A. DURR, V. BONIFATI, J. VAUGHAN, G. De MICHELE, T. GASSER, B. S. HARHANGI, G. MECO, P. DENEFLE, N. W. WOOD, Y. AGID, A. BRICE, Association between early-onset Parkinson’s disease and mutations in the parkin gene, N. Engl. J. Med. 342 (2000) 1560–1567. H. SHIMURA, N. HATTORI, S. KUBO, Y. MIZUNO, S. ASAKAWA, S. MINOSHIMA, N. SHIMIZU, K. IWAI, T. CHIBA, K. TANAKA, T. SUZUKI, Familial Parkinson disease gene product, parkin, is a ubiquitin-protein ligase, Nat. Genet. 25 (2000) 302–305. K. K. CHUNG, Y. ZHANG, K. L. LIM, Y. TANAKA, H. HUANG, J. GAO, C. A. ROSS, V. L. DAWSON, T. M. DAWSON, Parkin ubiquitinates the α-synuclein-interacting protein, synphilin-1: implications for Lewy-body formation in Parkinson disease, Nat. Med. 7 (2001) 1144–1150. H. SHIMURA, M. G. SCHLOSSMACHER, N. HATTORI, M. P. FROSCH, A. TROCKENBACHER, R. SCHNEIDER, Y. MIZUNO, K. S. KOSIK, D. J. SELKOE, Ubiquitination of a new form of α-synuclein by parkin from human brain: implications for Parkinson’s disease, Science 293 (2001) 263–269. Y. IMAI, M. SODA, H. INOUE, N. HATTORI, Y. MIZUNO, R. TAKAHASHI, An unfolded putative transmembrane polypeptide, which can lead to endoplasmic

21

22 Molecular Chaperones and the Ubiquitin–Proteasome System

21

22

23

24

25

26

27

28

29

30

31

reticulum stress, is a substrate of Parkin, Cell 105 (2001) 891–902. P. FERNANDEZ-FUNEZ, M. L. NINO-ROSALES, B. de GOUYON, W. C. SHE, J. M. LUCHAK, P. MARTINEZ, E. TURIEGANO, J. BENITO, M. CAPOVILLA, P. J. SKINNER, A. MCCALL, I. CANAL, H. T. ORR, H. Y. ZOGHBI, J. BOTAS, Identiﬁcation of genes that modify ataxin-1-induced neurodegeneration, Nature 408 (2000) 101–106. P. KAZEMI-ESFARJANI, S. BENZER, Genetic suppression of polyglutamine toxicity in Drosophila, Science 287 (2000) 1837–1840. N. M. BONINI, Chaperoning brain degeneration, Proc. Natl. Acad. Sci. USA 99 Suppl. 4 (2002) 16407–16411. P. K. AULUCK, H. Y. CHAN, J. Q. TROJANOWSKI, V. M. LEE, N. M. BONINI, Chaperone suppression of α-synuclein toxicity in a Drosophila model for Parkinson’s disease, Science 295 (2002) 865–868. F. U. HARTL, M. HAYER-HARTL, Molecular chaperones in the cytosol: from nascent chain to folded protein, Science 295 (2002) 1852–1858. J. FRYDMAN, Folding of newly translated proteins in vivo: the role of molecular chaperones, Annu. Rev. Biochem. 70 (2001) 603–647. J. FRYDMAN, J. H¨ohfeld, Chaperones get in touch: the Hip-Hop connection, Trends Biochem. Sci. 22 (1997) 87–92. B. BUKAU, A. L. HORWICH, The Hsp70 and Hsp60 chaperone machines, Cell 92 (1998) 351–366. R. I. MORIMOTO, Regulation of the heat shock transcriptional response: cross talk between a family of heat shock factors, molecular chaperones, and negative regulators. Genes Dev. 12 (1998) 3788–3796. M. P. MAYER, D. BREHMER, C. S. G¨ASSLER, B. BUKAU, Hsp70 chaperone machines, Adv. Protein Chem. 59 (2001) 1–44. S. R¨UDIGER, L. GERMEROTH, J. SCHNEIDER-MERGENER, B. BUKAU, Substrate speciﬁcity of the DnaK chaperone determined by screening cellulose-bound peptide libraries, EMBO J. 16 (1997) 1501–1507.

32 V. THULASIRAMAN, C. F. YANG, J. FRYDMAN, In vivo newly translated polypeptides are sequestered in a protected folding environment, EMBO J. 18 (1999) 85–95. 33 X. ZHU, X. ZHAO, W. F. BURKHOLDER, A. GRAGEROV, C. M. OGATA, M. E. GOTTESMAN, W. A. HENDRICKSON, Structural analysis of substrate binding by the molecular chaperone DnaK, Science 272 (1996) 1606–1614. 34 Y. MINAMI, J. H¨OHFELD, K. OHTSUKA, F. U. HARTL, Regulation of the heat-shock protein 70 reaction cycle by the mammalian DnaJ homolog, Hsp40, J. Biol. Chem. 271 (1996) 19617–19624. 35 C. A. BALLINGER, P. CONNELL, Y. WU, Z. HU, L. J. THOMPSON, L. Y. YIN, C. PATTERSON, Identiﬁcation of CHIP, a novel tetratricopeptide repeat-containing protein that interacts with heat shock proteins and negatively regulates chaperone functions, Mol. Cell. Biol. 19 (1999) 4535–4545. 36 J. H¨OHFELD, Y. MINAMI, F. U. HARTL, Hip, a novel cochaperone involved in the eukaryotic Hsc70/Hsp40 reaction cycle, Cell 83 (1995) 589–598. 37 J. H¨OHFELD, S. JENTSCH, GrpE-like regulation of the hsc70 chaperone by the anti-apoptotic protein BAG-1, Embo J. 16 (1997) 6209–6216. 38 H. SONDERMANN, C. SCHEUFLER, C. SCHNEIDER, J. H¨OHFELD, F. U. HARTL, I. MOAREFI, Structure of a Bag/Hsc70 complex: convergent functional evolution of Hsp70 nucleotide exchange factors, Science 291 (2001) 1553– 1557. 39 C. S. G¨ASSLER, T. WIEDERKEHR, D. BREHMER, B. BUKAU, M. P. MAYER, Bag-1M accelerates nucleotide release for human Hsc70 and Hsp70 and can act concentration-dependent as positive and negative cofactor, J. Biol. Chem. 276 (2001) 32538–32544. 40 J. H¨OHFELD, D. M. CYR, C. PATTERSON, From the cradle to the grave: molecular chaperones that may choose between folding and degradation, EMBO Rep. 2 (2001) 885–890. 41 W. M. OBERMANN, H. SONDERMANN, A. A. RUSSO, N. P. PAVLETICH, F. U. HARTL, In

References

42

43

44

45

46

47

48

49

50

vivo function of Hsp90 is dependent on ATP binding and ATP hydrolysis, J. Cell Biol. 143 (1998) 901–910. J. BUCHNER, Hsp90 & Co. – a holding for folding, Trends Biochem. Sci. 24 (1999) 136–141. J. C. YOUNG, I. MOAREFI, F. U. HARTL, Hsp90: a specialized but essential protein-folding tool, J. Cell Biol. 154 (2001) 267–273. L. H. PEARL, C. PRODROMOU, Structur, function and mechanism of the Hsp90 molecular chaperone, Adv. Protein Chem. 59 (2001) 157–186. B. PANARETOU, G. SILIGARDI, P. MEYER, A. MALONEY, J. K. SULLIVAN, S. SINGH, S. H. MILLSON, P. A. CLARKE, S. NAABY-HANSEN, R. STEIN, R. CRAMER, M. MOOAPOUR, P. WORKMAN, P. W. PIPER, L. H. PEAL, C. PRODROMOU, Activation of the ATPase activity of hsp90 by the stress-regulated cochaperone aha1, Mol. Cell 10 (2002) 1307–1318. J. C. YOUNG, N. J. HOOGENRAAD, F. U. HARTL, Molecular chaperones Hsp90 and Hsp70 deliver preproteins to the mitochondrial import receptor Tom70, Cell 112 (2003) 41–50. A. BRYCHZY, T. REIN, K. F. WINKLHOFER, F. U. HARTL, J. C. YOUNG, W. M. OBERMANN, Cofactor Tpr2 combines two TPR domains and a J domain to regulate the Hsp70/Hsp90 chaperone system, EMBO J. 22 (2003) 3613–3623. S. M. ROE, M. M. ALI, P. MEYER, C. K. VAUGHAN, B. PANARETOU, P. W. PIPER, C. PRODROMOU, L. H. PEARL, The mechanism of Hsp90 regulation by the protein kinase-speciﬁc cochaperone p50 (cdc37), Cell 116 (2004) 87–98. K. D. DITTMAR, W. B. PRATT, Folding of the glucocorticoid receptor be the reconstituted Hsp90-based chaperone machinery. The initial hsp90.p60.hsp70 dependent step is sufﬁcient for creating the steroid binding conformation, J. Biol. Chem. 272 (1997) 13047–13054. K. D. DITTMAR, D. R. DEMADY, L. F. STANCATO, P. KRISHNA, W. B. PRATT, Folding of the glucocorticoid receptor by the heat shock protein (hsp) 90-based chaperone machinery. The role of p23 is to stabilize receptor.hsp90

51

52

53

54

55

56

57

58

59

heterocomplexes formed by hsp90.p60.hsp70, J. Biol. Chem. 272 (1997) 21213–21220. S. CHEN, V. PRAPAPANICH, R. A. RIMERMAN, B. HONORE, D. F. SMITH, Interactions of p60, a mediator of progesterone receptor assembly, with heat shock proteins hsp90 and hsp70. Mol. Endocrinol. 10 (1996) 682–693. H. KOSANO, B. STENSGARD, M. C. CHARLESWORTH, N. MCMAHON, D. TOFT, The assembly of progesterone receptor-hsp90 complexes using puriﬁed proteins, J. Biol. Chem. 273 (1998) 32973–32979. W. B. PRATT, D. O. TOFT, Regulation of signaling protein function and trafﬁcking by the hsp90/hsp70-based chaperone machinery, Exp. Biol. Med. 228 (2003) 111–133. M. P. HERNANDEZ, A. CHADLI, D. O. TOFT, Hsp40 binding is the ﬁrst step in the Hsp90 chaperoning pathway for the progesterone receptor, J. Biol. Chem. 277 (2002) 11873–11881. C. SCHEUFLER, A. BRINKER, G. BOURENKOV, S. PEGORARO, L. MORODER, H. BARTUNIK, F. U. HARTL, I. MOAREFI, Structure of TPR domain-peptide complexes: critical elements in the assembly of the Hsp70-Hsp90 multichaperone machine, Cell 101 (2000) 199–210. J. FONTANA, D. FULTON, Y. CHEN, T. A. FAIRCHILD, T. J. MCCABE, N. FUJITA, T. TSURUO, W. C. SESSA, Domain mapping studies reveal that the M domain of hsp90 serves as a molecular scaffold to regulate Akt-dependent phosphorylation of endothelioal nitric oxide synthase and NO release, Circ. Res. 90 (2002) 866–873. D. B. SOLIT, H. I. SCHER, N. ROSEN, Hsp90 as a therapeutic target in prostate cancer, Semin. Oncol. 30 (2003) 709–716. L. NECKERS, S. P. IVY, Heat shock protein 90, Curr. Opin. Oncol. 15 (2003) 419– 424. A. KAMAL, L. THAO, J. SENSINTAFFAR, L. ZHANG, M. F. BOEHM, L. C. FRITZ, F. J. BURROWS, A high-afﬁnity conformation of Hsp90 confers tumour selectivity on Hsp90 inhibitors, Nature 425 (2003) 407–410.

23

24 Molecular Chaperones and the Ubiquitin–Proteasome System

60 C. PRODROMOU, S. M. ROE, R. O’BRIEN, J. E. LADBURY, P. W. PIPER, L. H. PEARL, Identiﬁcation and structural characterization of the ATP/ADP-binding site in the Hsp90 molecular chaperone, Cell 90 (1997) 65–75. 61 C. E. STEBBINS, A. A. RUSSO, C. SCHNEIDER, N. ROSEN, F. U. HARTL, N. P. PAVLETICH, Crystal structure of an Hsp90-geldanamycin complex: targeting of a protein chaperone by an antitumor agent, Cell 89 (1997) 239–250. 62 S. V. SHARMA, T. AGATSUMA, H. NAKANO, Targeting of the protein chaperone, Hsp90, by the transformation suppressing agent, radicicol, Oncogene 16 (1998) 2639–2645. 63 T. W. SCHULTE, S. AKINAGA, T. MURAKATA, T. AGATSUMA, S. SUGIMOTO, H. NAKANO, Y. S. LEE, B. B. SIMEN, Y. ARGON, S. FELTS, D. O. TOFT, L. M. NECKERS, S. V. SHARMA, Interaction of radicicol with members of the heat shock protein 90 family of molecular chaperones, Mol. Endocrinol. 13 (1999) 1435–1448. 64 C. SCHNEIDER, L. SEPP-LORENZINO, E. NIMMESGERN, O. OUERFELLI, S. DANISHEFSKY, N. ROSEN, F. U. HARTL, Pharmacologic shifting of a balance between protein refolding and degradation mediated by Hsp90, Proc. Natl. Acad. Sci. USA 93 (1996) 14536–14541. 65 E. G. MIMNAUGH, C. CHAVANY, L. NECKERS, Polyubiquitination and proteasomal degradation of the p185c-erbB-2 receptor protein-tyrosine kinase induced by geldanamycin, J. Biol. Chem. 271 (1996) 22796–22801. 66 M. G. MARCU, T. W. SCHULTE, L. NECKERS, Novobiocin and related coumarins and depletion of heat shock protein 90-dependent signaling proteins, J. Natl. Cancer Inst. 92 (2000) 242–248. 67 W. XU, E. MIMNAUGH, M. F. ROSSER, C. NICCHITTA, M. MARCU, Y. YARDEN, L. NECKERS, Sensitivity of mature Erbb2 to geldanamycin is conferred by its kinase domain and is mediated by the chaperone protein Hsp90, J. Biol. Chem. 276 (2001) 3702–3708.

68 D. B. SOLIT, F. F. ZHENG, M. DROBNJAK, P. N. MUNSTER, B. HIGGINS, D. VERBEL, G. HELLER, W. TONG, C. CORDON-CARDO, D. B. AGUS, H. I. SCHER, N. ROSEN, 17Allylamino-17-demethoxygeldanamycin induces the degradation of androgen receptor and HER-2/neu and inhibits the growth of prostate cancer xenografts, Clin. Cancer Res. 8 (2002) 986–993. 69 A. CITRI, I. ALROY, S. LAVI, C. RUBIN, W. XU, N. GRAMMATIKAKIS, C. PATTERSON, L. NECKERS, D. W. FRY, Y. YARDEN, Drug-induced ubiquitylation and degradation of ErbB receptor tyrosine kinases: implications for cancer therapy, Embo J. 21 (2002) 2407–2417. 70 R. van MONTFORT, C. SLINGSBY, E. VIERLING, Structure and function of the small heat shock protein/alpha-crystallin family of molecular chaperones, Adv. Protein Chem. 59 (2001) 105–156. 71 M. HASLBECK, N. BRAUN, T. STROMER, B. RICHTER, N. MODEL, S. WEINKAUF, J. BUCHNER, Hsp42 is the general small heat shock protein in the cytosol of Saccharomyces cerevisiae, EMBO J. 23 (2004) 638–649. 72 T. STROMER, E. FISCHER, K. RICHTER, M. HASLBECK, J. BUCHNER, Analysis of the regulation of the molecular chaperone Hsp26 by temperature-induced dissociation: the N-terminal domain is important for oligomer assembly and the binding of unfolding proteins. J. Biol. Chem. 279 (2004) 11222–11228. 73 E. BASHA, G. J. LEE, L. A. BRECI, A. C. HAUSRATH, N. R. BUAN, K. C. GIESE, E. VIERLING, The identity of proteins associated with a small heat shock protein during heat stress in vivo indicates that these chaperones protect a wide range of cellular functions, J. Biol. Chem. 279 (2004) 7566–7575. 74 K. K. KIM, R. KIM, S. H. KIM, Crystal structure of a small heat-shock protein, Nature 394 (1998) 595–599. 75 R. L. van MONTFORT, E. BASHA, K. L. FRIEDRICH, C. SLINGSBY, E. VIERLING, Crystal structure and assembly of a eukaryotic small heat shock protein. Nat. Struct. Biol. 8 (2001) 1025–1030. 76 M. HASLBECK, S. WALKE, T. STROMER, M. EHRNSPERGER, H. E. WHITE, S. CHEN, H.

References

77

78

79

80

81

82

83

84

85

86

R. SAIBIL, J. BUCHNER, Hsp26: a temperature-regulated chaperone, EMBO J. 18 (1999) 6744–6751. G. J. LEE, A. M. ROSEMAN, H. R. SAIBIL, E. VIERLING, A small heat shock protein stably binds heat-denatured model substrates and can maintain a substrate in a folding-competent state, EMBO J. 16 (1997) 659–671. Z. XU, A. L. HORWICH, P. B. SIGLER, The crystal structure of the asymmetric GroEL-GroES-(ADP)7 chapero-nin complex, Nature 388 (1997) 741–750. H. R. SAIBIL, N. A. RANSON, The chaperonin folding machine, Trends Biochem. Sci. 27 (2002) 627–632. A. S. MEYER, J. R. GILLESPIE, D. WALTHER, I. S. MILLET, S. DONIACH, J. FRYDMAN, Closing the folding chamber of the eukaryotic chaperonin requires the transition state of ATP hydrolysis, Cell 113 (2003) 369–381. V. THULASIRAMAN, C. F. YANG, J. FRYDMAN, In vivo newly translated polypeptides are sequestered in a protected folding environment, EMBO J. 18 (1999) 85–95. S. A. LEWIS, G. TIAN, I. E. VAINBERG, N. J. COWAN, Chaperonin-mediated folding of actin and tubulin, J. Cell Biol. 132 (1996) 1–4. H. SAKAHIRA, P. BREUER, M. K. HAYER-HARTL, F. U. HARTL, Molecular chaperones as modulators of polyglutamine protein aggregation and toxicity, Proc. Natl. Acad. Sci. USA 99 Suppl. 4 (2002) 16412–16418. J. M. WARRICK, H. Y. CHAN, G. L. GRAY-BOARD, Y. CHAI, H. L. PAULSON, N. M. BONINI, Suppression of polyglutamine-mediated neuro-degeneration in Drosophila by the molecular chaperone Hsp70. Nat. Genet. 23 (1999) 425–428. C. J. CUMMINGS, M. A. MANCINI, B. ANTALFFY, D. B. DEFRANCO, H. T. ORR, H. Y. ZOGHBI, Chaperone suppression of aggregation and altered subcellular proteasome localization imply protein misfolding in SCA1, Nat. Genet. 19 (1998) 148–154. C. J. CUMMINGS, Y. SUN, P. OPAL, B. ANTALFFY, R. MESTRIL, H. T. ORR, W. H.

87

88

89

90

91

92

93

DILLMANN, H. Y. ZOGHBI, Over-expression of inducible HSP70 chaperone suppresses neuropathology and improves motor function in SCA1 mice, Hum. Mol. Genet. 10 (2001) 1511–1518. S. KROBITSCH, S. LINDQUIST, Aggregation of huntingtin in yeast varies with the length of the polyglutamine expansion and the expression of chaperone proteins, Proc. Natl. Acad. Sci. USA 97 (2000) 1589–1594. P. J. MUCHOWSKI, G. SCHAFFAR, A. SITTLER, E. E. WANKER, M. K. HAYER-HARTL, F. U. HARTL, Hsp70 and hsp40 chaperones can inhibit self-assembly of polyglutamine proteins into amyloid-like ﬁbrils, Proc. Natl. Acad. Sci. USA 97 (2000) 7841–7846. J. Z. CHUANG, H. ZHOU, M. ZHU, S. H. LI, X. J. LI, C. H. SUNG, Characterization of a brain-enriched chaperone, MRJ, that inhibits Huntingtin aggregation and toxicity independently, J. Biol. Chem. 277 (2002) 19831–19838. A. SITTLER, R. LURZ, G. LUEDER, J. PRILLER, H. LEHRACH, M. K. HAYER-HARTL, F. U. HARTL, E. E. WANKER, Geldanamycin activates a heat shock response and inhibits huntingtin aggregation in a cell culture model of Huntington’s disease, Hum. Mol. Genet. 10 (2001) 1307–1315. A. PARCELLIER, E. SCHMITT, S. GURBUXANI, D. SEIGNEURIN-BERNY, A. PANCE, A. CHANTOME, S. PLENCHETTE, S. KHOCHBIN, E. SOLARY, C. GARRIDO, Hsp27 is a ubiquitin-binding protein involved in IκBα proteasomal degradation, Mol. Cell Biol. 23 (2003) 5790–5802. H. SHIMURA, Y. MIURA-SHIMURA, K. S. KOSICK, Binding of tau to heat shock protein 27 leads to decreased concentration of hyperphosphorylated tau and enhanced cell survival, J. Biol. Chem. 279 (2004) 17957–17962. V. GUSAROVA, A. J. CAPLAN, J. L. BRODSKY, E. A. FISHER, Apoprotein B degradation is promoted by the molecular chaperones hsp90 and hsp70, J. Biol. Chem. 276 (2001) 24891– 24900.

25

26 Molecular Chaperones and the Ubiquitin–Proteasome System

94 C. TAXIS, R. HITT, S. H. PARK, P. M. DEAK, Z. KOSTOVA, D. H. WOLF, Use of modular substrates demonstrates mechanistic diversity and reveals differences in chaperone requirement of ERAD, J. Biol. Chem. 278 (2003) 35903–35913. 95 Y. YANG, S. JANICH, J. A. COHN, J. M. WILSON, The common variant of cystic ﬁbrosis transmembrane conductance regulator is recognized by hsp70 and degraded in a pre-Golgi nonlysosomal compartment, Proc. Natl. Acad. Sci. USA 90 (1993) 9480–9484. 96 G. C. MEACHAM, Z. LU, S. KING, E. SORSCHER, A. TOUSSON, D. M. CYR, The Hdj-2/Hsc70 chaperone pair facilitates early steps in CFTR biogenesis, Embo J. 18 (1999) 1492–1505. 97 G. C. MEACHAM, C. PATTERSON, W. ZHANG, J. M. YOUNGER, D. M. CYR, The Hsc70 co-chaperone CHIP targets immature CFTR for proteasomal degradation, Nat. Cell Biol. 3 (2001) 100–105. 98 Y. ZHANG, G. NIJBROEK, M. L. SULLIVAN, A. A. MCCRACKEN, S. C. WATKINS, S. MICHAELIS, J. L. BRODSKY, Hsp70 molecular chaperone facilitates endoplasmic reticulum-associated protein degradation of cystic ﬁbrosis transmembrane conductance regulator in yeast, Mol. Biol. Cell 12 (2001) 1303–1314. 99 R. R. KOPITO, Biosynthesis and degradation of CFTR, Physiol. Rev. 79 (1999) S167–173. 100 J. R. RIORDAN, Cystic ﬁbrosis as a disease of misprocessing of the cystic ﬁbrosis transmembrane conductance regulator glycoprotein, Am. J. Hum. Genet. 64 (1999) 1499–1504. 101 C. L. WARD, S. OMURA, R. R. KOPITO, Degradation of CFTR by the ubiquitin proteasome pathway, Cell 83 (1995) 121–127. 102 T. J. JENSEN, M. A. LOO, S. PIND, D. B. WILLIAMS, A. L. GOLDBERG, J. R. RIORDAN, Multiple proteolytic systems, including the proteasome, contribute to CFTR processing, Cell 83 (1995) 129–135. 103 M. S. GELMAN, E. S. KANNEGAARD, R. R. KOPITO, A principal role for the

104

105

106

107

108

109

110

111

112

113

proteasome in endoplasmic reticulum-associated degradation of misfolded intracellular cystic ﬁbrosis transmembrane conductance regulator, J. Biol. Chem. 277 (2002) 11709–11714. Z. KOSTOVA, D. H. WOLF, For whom the bell tolls: protein quality control of the endoplasmic reticulum and the ubiquitin proteasome connection, EMBO J. 22 (2003) 2309–2317. S. SATO, C. L. WARD, M. E. KROUSE, J. J. WINE, R. R. KOPITO, Glycerol reverses the misfolding phenotype of the most common cystic ﬁbrosis mutation, J. Biol. Chem. 271 (1996) 635–638. B. H. QU, P. J. THOMAS, Alteration of the cystic ﬁbrosis transmembrane conductance regulator folding pathway, J. Biol. Chem. 271 (1996) 7261–7264. E. STRICKLAND, B. H. QU, L. MILLEN, P. J. THOMAS, The molecular chapeorne Hsc70 assists the in vitro folding of the N-terminal nucleotide-binding domain of the cystic ﬁbrosis transmembrane conductance regulator, J. Biol. Chem. 272 (1997) 25421–25424. J. A. JOHNSTON, C. L. WARD, R. R. KOPITO, Aggresomes: a cellular response to misfolded proteins, J. Cell Biol. 143 (1998) 1883–1898. R. R. KOPITO, Aggresomes, inclusion bodies and protein aggregation, Trends Cell Biol. 10 (2000) 524–530. S. WAELTER, A. BOEDDRICH, R. LURZ, E. SCHERZINGER, G. LUEDER, H. LEHRACH, E. E. WANKER, Accumulation of mutant huntingtin fragments in aggresome-like inclusion bodies as a result of insufﬁcient protein degradation, Mol. Biol. Cell 12 (2001) 1393–1407. R. GARCIA-MATA, Z. BEBOK, E. J. SORSCHER, E. S. SZTUL, Characterization and dynamics of aggresome formation by a cytosolic GFP-chimera, J. Cell Biol. 146 (1999) 1239–1254. K. S. MCNAUGHT, P. SHASHIDHARAN, D. P. PERL, P. JENNER, C. W. OLANOW, Aggresome-related biogenesis of Lewy bodies, Eur. J. Neurosci. 16 (2002) 2136–2148. Y. KAWAGUCHI, J. J. KOVACS, A. MCLAURIN, J. M. VANCE, A. ITO, T. P. YAO, The deacetylase HDAC6 regulates

References

114

115

116

117

118

119

120

121

122

aggresome formation and cell viability in response to misfolded protein stress, Cell 115 (2003) 727–738. R. GARCIA-MATA, Z. BEBOK, E. J. SORSCHER, E. S. SZTUL, Characterization and dynamics of aggresome formation by a cytosolic GFP-chimera, J. Cell Biol. 146 (1999) 1239–1254. W. C. WIGLEY, R. P. FABUNMI, M. G. LEE, C. R. MARINO, S. MUALLEM, G. N. DEMARTINO, P. J. THOMAS, Dynamic association of proteasomal machinery with the centrosome, J. Cell Biol. 145 (1999) 481–490. A. WYTTENBACH, J. CARMICHAEL, J. SWARTZ, R. A. FURLONG, Y. NARAIN, J. RANKIN, D. C. RUBINSZTEIN, Effects of heat shock, heat shock protein 40 (HDJ-2), and proteasome inhibition on protein aggregation in cellular models of Huntington’s disease, Proc. Natl. Acad. Sci. USA 97 (2000) 2898–2903. J. L. DUL, D. P. DAVIS, E. K. WILLIAMSON, F. J. STEVENS, Y. ARGON, Hsp70 and antiﬁbrillogenic peptides promote degradation and inhibit intracellular aggregation of amyloidogenic light chains, J. Cell Biol. 152 (2001) 705–716. H. LELOUARD, E. GATTI, F. CAPPELLO, O. GRESSER, V. CAMOSSETO, P. PIERRE, Transient aggregation of ubiquitinated proteins during dendritic cell maturation, Nature 417 (2002) 177–182. H. LELOUARD, V. FERRAND, D. MARGUET, J. BANIA, V. CAMOSSETO, A. DAVID, E. GATTI, P. PIERRE, Dendritic cell aggresome-like induced structures are dedicated areas for ubiquitination and storage of newly synthesized defective proteins, J. Cell Biol. 164 (2004) 667–675. W. XU, M. MARCU, X. YUAN, E. MIMNAUGH, C. PATTERSON, L. NECKERS, Chaperone-dependent E3 ubiquitin ligase CHIP mediates a degradative pathway for c-ErbB2/Neu, Proc. Natl. Acad. Sci. USA 99 (2002) 12847–12852. S. WICKNER, M. R. MAURIZI, S. GOTTESMAN, Posttranslational quality control: folding, refolding, and degrading proteins, Science 286 (1999) 1888–1893. C.-W. LIU, M. J. CORBOY, G. N. DEMARTINO, P. J. THOMAS,

123

124

125

126

127

128

129

130

131

Endoproteolytic activity of the proteasome, Science 299 (2003) 408–411. W. SEUFERT, S. JENTSCH, Ubiquitin-conjugating enzymes UBC4 and UBC5 mediate selective degradation of short-lived and abnormal proteins, EMBO J. 9 (1990) 543–550. B. BERCOVICH, I. STANCOVSKI, A. MAYER, N. BLUMENFELD, A. LASZLO, A. L. SCHWARTZ, A. CIECHANOVER, Ubiquitin-dependent degradation of certain protein substrates in vitro requires the molecular chaperone Hsc70, J. Biol. Chem. 272 (1997) 9002–9010. P. CONNELL, C. A. BALLINGER, J. JIANG, Y. WU, L. J. THOMPSON, J. H¨OHFELD, C. PATTERSON, The co-chaperone CHIP regulates protein triage decisions mediated by heat-shock proteins, Nat. Cell Biol. 3 (2001) 93–96. L. ARAVIND, E. V. KOONIN, The U box is a modiﬁed RING ﬁnger – a common domain in ubiquitination, Curr. Biol. 10 (2000) R132–134. S. HATAKEYAMA, M. YADA, M. MATSUMOTO, N. ISHIDA, K. I. NAKAYAMA, U box proteins as a new family of ubiquitin-protein ligases, J. Biol. Chem. 276 (2001) 33111–33120. E. PRINGA, G. MARTINEZ-NOEL, U. M¨ULLER, K. HARBERS, Interaction of the ring ﬁnger-related U-box motif of a nuclear dot protein with ubiquitin-conjugating enzymes, J. Biol. Chem. 276 (2001) 19617– 19623. R. NIKOLAY, T. WIEDERKEHR, W. RIST, G. KRAMER, M. P. MAYER, B. BUKAU, Dimerization of the human E3 ligase CHIP via a coiled-coil domain is essential for its activity, J. Biol. Chem. 279 (2004) 2673–2678. Q. DAI, C. ZHANG, Y. WU, H. MCDONOUGH, R. A. WHALEY, V. GODFREY, H. H. LI, N. MADAMANCHI, W. XU, L. NECKERS, D. CYR, C. PATTERSON, CHIP activates HSF1 and confers protection against apoptosis and cellular stress, EMBO J. 22 (2003) 5446–5458. J. DEMAND, J. L¨UDERS, J. H¨OHFELD, The carboxyl-terminal domain of Hsc70 provides binding sites for a distinct set

27

28 Molecular Chaperones and the Ubiquitin–Proteasome System

132

133

134

135

136

137

138

139

140

of chaperone cofactors, Mol. Cell. Biol. 18 (1998) 2023–2028. A. BRINKER, C. SCHEUFLER, F. VON DER MULBE, B. FLECKENSTEIN, C. HERRMANN, G. JUNG, I. MOAREFI, F. U. HARTL, Ligand discrimination by TPR domains. Relevance and selectivity of EEVD-recognition in Hsp70 xHop xHsp90 complexes, J. Biol. Chem. 277 (2002) 19265–19275. H. H. KAMPINGA, B. KANON, F. A. SALOMONS, A. E. KABAKOV, C. PATTERSON, Overexpression of the cochaperone CHIP enhances Hsp70-dependent folding activity in mammalian cells, Mol. Cell. Biol. 23 (2003) 4948–4958. H. SHIMURA, D. SCHWARTZ, S. P. GYGI, K. S. KOSIK, CHIP-Hsc70 complex ubiquitinates phosphorylated tau and enhances cell survival, J. Biol. Chem. 279 (2004) 4869–4876. J. JIANG, C. A. BALLINGER, Y. WU, Q. DAI, D. M. CYR, J. H¨OHFELD, C. PATTERSON, CHIP is a U-box-dependent E3 ubiquitin ligase: identiﬁcation of Hsc70 as a target for ubiquitylation, J. Biol. Chem 276 (2001) 42938–42944. S. MURATA, Y. MINAMI, M. MINAMI, T. CHIBA, K. TANAKA, CHIP is a chaperone-dependent E3 ligase that ubiquitylates unfolded protein, EMBO Rep. 2 (2001) 1133–1138. J. DEMAND, S. ALBERTI, C. PATTERSON, J. H¨ohfeld, Cooperation of a ubiquitin domain protein and an E3 ubiquitin ligase during chaperone/proteasome coupling, Curr. Biol. 11 (2001) 1569–1577. P. K. JACKSON, A. G. ELDRIDGE, E. FREED, L. FURSTENTHAL, J. Y. HSU, B. K. KAISER, J. D. REIMANN, The lore of the RINGs: substrate recognition and catalysis by ubiquitin ligases, Trends Cell. Biol. 10 (2000) 429–439. P. K. JACKSON, A. G. ELDRIDGE, The SCF ubiquitin ligase: an extended look, Mol. Cell 9 (2002) 923–925. N. ZHENG, B. A. SCHULMAN, L. SONG, J. J. MILLER, P. D. JEFFREY, P. WANG, C. CHU, D. M. KOEPP, S. J. ELLEDGE, M. PAGANO, R. C. CONAWAY, J. W. CONAWAY, J. W. HARPER, N. P. PAVLETICH, Structure of the Cul1-Rbx1-Skp1-F boxSkp2 SCF

141

142

143

144

145

146

147

148

149

ubiquitin ligase complex, Nature 416 (2002) 703–709. J. L¨UDERS, J. DEMAND, J. H¨ohfeld, The ubiquitin-related BAG-1 provides a link between the molecular chaperones Hsc70/Hsp70 and the proteasome, J. Biol. Chem. 275 (2000) 4613–4617. S. JENTSCH, G. PYROWOLAKIS, Ubiquitin and its kin: how close are the family ties? Trends Cell. Biol. 10 (2000) 335– 342. S. ALBERTI, J. DEMAND, C. ESSER, N. EMMERICH, H. SCHILD, J. H¨OHFELD, Ubiquitylation of BAG-1 suggests a novel regulatory mechanism during the sorting of chaperone substrates to the proteasome, J. Biol. Chem. 277 (2002) 45920–45927; Erratum in J. Biol. Chem. 278 (2003) 15702–15703. Q. DEVERAUX, V. USTRELL, C. PICKART, M. RECHSTEINER, A 26 S protease subunit that binds ubiquitin conjugates, J. Biol. Chem. 269 (1994) 7059–7061. P. YOUNG, Q. DEVERAUX, R. E. BEAL, C. M. PICKART, M. RECHSTEINER, Characterization of two polyubiquitin binding sites in the 26 S protease subunit 5a, J. Biol. Chem. 273 (1998) 5461–5467. H. HIYAMA, M. YOKOI, C. MASUTANI, K. SUGASAWA, T. MAEKAWA, K. TANAKA, J. H. HOEIJMAKERS, F. HANAOKA, Interaction of hHR23 with S5a. The ubiquitin-like domain of hHR23 mediates interaction with S5a subunit of 26 S proteasome, J. Biol. Chem. 274 (1999) 28019–28025. K. J. WALTERS, M. F. KLEIJNEN, A. M. GOH, G. WAGNER, P. M. HOWLEY, Structural studies of the interaction between ubiquitin family proteins and proteasome subunit S5a, Biochemistry 41 (2002) 1767–1777. Y. A. LAM, T. G. LAWSON, M. VELAYUTHAM, J. L. ZWEIER, C. M. PICKART, A proteasomal ATPase subunit recognizes the polyubiquitin degradation signal, Nature 416 (2002) 763–767. S. ELSASSER, R. R. GALI, M. SCHWICKART, C. N. LARSEN, D. S. LEGGETT, B. MULLER, M. T. FENG, F. TUBING, G. A. DITTMAR, D. FINLEY, Proteasome subunit Rpn1 binds ubiquitin-like protein domains, Nat. Cell Biol. 4 (2002) 725–730.

References 150 Y. SAEKI, T. SONE, A. TOHE, H. YOKOSAWA, Identiﬁcation of ubiquitin-like protein-binding subunits of the 26S proteasome, Biochem. Biophys. Res. Commun. 296 (2002) 813–819. 151 M. H. GLICKMAN, D. M. RUBIN, O. COUX, I. WEFES, G. PFEIFER, Z. CJEKA, W. BAUMEISTER, V. A. FRIED, D. FINLEY, A subcomplex of the proteasome regulatory particle required for ubiquitin-conjugate degradation and related to the COP9-signalosome and eIF3, Cell 94 (1998) 615–623. 152 Y. XIE, A. VARSHAVSKY, UFD4 lacking the proteasome-binding region catalyses ubiquitination but is impaired in proteolysis, Nat. Cell Biol. 4 (2002) 1003–1007. 153 S. TAKAYAMA, J. C. REED, Molecular chaperone targeting and regulation by BAG family proteins, Nat. Cell Biol. 3 (2001) E237–241. 154 K. THRESS, J. SONG, R. I. MORIMOTO, S. KORNBLUTH, Reversible inhibition of Hsp70 chaperone function by Scythe and Reaper, EMBO J. 20 (2001) 1033–1041. 155 S. ALBERTI, K. B¨OHSE, V. ARNDT, A. SCHMITZ, J. H¨ohfeld, The co-chaperone HspBP1 inhibits the CHIP ubiquitin ligase and stimulates the maturation of the cystic ﬁbrosis transmembrane conductance regulator, Mol. Biol. Cell (2004), in press. 156 D. A. RAYNES, V. GUERRIERO, Inhibition of Hsp70 ATPase activity and protein renaturation by a novel Hsp70-binding protein, J. Biol. Chem. 273 (1998) 32883–32888. 157 M. KABANI, C. MCLELLAN, D. A. RAYNES, V. GUERRIERO, J. L. BRODSKY, HspBP1, a homologue of the yeast Fes1 and Sls1 proteins, is an Hsc70 nucleotide exchange factor, FEBS Lett. 531 (2002) 339–342. 158 C. P. CARDOZO, C. MICHAUD, M. C. OST, A. E. FLISS, E. YANG, C. PATTERSON, S. J. HALL, A. J. CAPLAN, C-terminal Hsp-interacting protein slows androgen receptor synthesis and reduces its rate of degradation, Arch. Biochem. Biophys. 410 (2003) 134–140. 159 J. JIANG, D. CYR, R. W. BABBITT, W. C. SESSA, C. PATTERSON,

160

161

162

163

164

165

166

167

Chaperone-dependent regulation of endothelial nitric-oxide synthase intracellular trafﬁcking by the co-chaperone/ubiquitin ligase CHIP, J. Biol. Chem. 278 (2003) 49332– 49341. Y. IMAI, M. SODA, S. HATAKEYAMA, T. AKAGI, T. HASHIKAWA, K. I. NAKAYAMA, R. TAKAHASHI, CHIP is associated with Parkin, a gene responsible for familial Parkinson’s disease, and enhances its ubiquitin ligase activity, Mol. Cell 10 (2002) 55–67. Y. NAGANO, H. YAMASHITA, T. TAKAHASHI, S. KISHIDA, T. NAKAMURA, E. ISEKI, N. HATTORI, Y. MIZUNO, A. KIKUCHI, M. MATSUMOTO, Siah-1 facilitates ubiquitination and degradation of synphilin-1, J. Biol. Chem. 278 (2003) 51504–51514. E. LIANI, A. EYAL, E. AVRAHAM, R. SHERMER, R. SZARGEL, D. BERG, A. BORNEMANN, O. RIESS, C. A. ROSS, R. ROTT, S. ENGELENDER, Ubiquitylation of synphilin-1 and alpha-synuclein by SIAH and its presence in cellular inclusions and Lewy bodies imply a role in Parkinson’s disease, Proc. Natl. Acad. Sci. USA 101 (2004) 5500–5505. S. MATSUZAWA, S. TAKAYAMA, B. A. FROESCH, J. M. ZAPATA, J. C. REED, p53-inducible homologue of Drosophila seven in absentia (Siah) inhibits cell growth: suppression by BAG-1, EMBO J. 17 (1998) 2736–2747. C. SOTI, P. CSERMELY, Aging and molecular chaperones, Exp. Gerontol. 38 (2003) 1037–1040. J. FARGNOLI, T. KUNISADA, A. J. Jr. FORNACE, E. L. SCHNEIDER, N. J. HOLBROOK, Decreased expression of heat shock protein 70 mRNA and protein after heat tretament in cells of aged rats, Proc. Natl. Acad. Sci. USA 87 (1990) 846–850. C. K. LEE, R. G. KLOPP, R. WEINDRUCH, T. A. PROLLA, Gene expression proﬁle of aging and its retardation by caloric restriction, Science 285 (1999) 1390–1393. S. X. CAO, J. M. DHAHBI, P. L. MOTE, S. R. SPINDLER, Genomic proﬁling of shortand long-term caloric restriction effects

29

30 Molecular Chaperones and the Ubiquitin–Proteasome System

in the liver of aging mice, Proc. Natl. Acad. Sci. USA 98 (2001) 10630–10635. 168 M. TATAR, A. A. KHAZAELI, J. W. CURTSINGER, Chaperoning extended life, Nature 390 (1997) 30. 169 G. MORROW, R. M. TANGUAY, Heat shock proteins and aging in Drosophila melanogaster, Semin. Cell. Dev. Biol. 14 (2003) 291–299.

170 J. F. MORLEY, R. I. MORIMOTO, Regulation of longevity in Caenorhabditis elegans by heat shock factor and molecular chaperones, Mol. Biol. Cell 15 (2004) 657–664. 171 A. L. HSU, C. T. MURPHY, C. KENYON, Regulation of aging and age-related disease by DAF-16 and heat-shock factor, Science 300 (2003) 1142–1145.

1

Ubiquitin-conjugating Enzymes Michael J. Eddins Johns Hopkins School of Medicine, Baltimore, USA

Cecile M. Pickart Johns Hopkins Bloomberg School of Public Health, Baltimore, USA

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8

1 Introduction

In this chapter we review the biochemical, structural, and biological properties of ubiquitin-conjugating enzymes (also called E2 enzymes). Because length restrictions preclude a comprehensive treatment, we focus on key findings that have revealed important general insights and principles. Throughout the piece we try to point out important unanswered questions concerning the E2 enzyme family. A few words about nomenclature are necessary. The yeast E2 genes were numbered in the order of their discovery, but the situation is more complicated in mammals. There are currently three naming systems in use for human E2s: one based on protein molecular mass (e.g. E225K is a 25-kD E2), one based on temporal order of gene cloning (e.g. UbcH10 is specified by the tenth E2 gene cloned in humans), and one based on relationship to yeast E2s (e.g. HR6A is one of two human homologs of yeast Rad6/Ubc2). The second system is the least ambiguous, but also the least informative. In this chapter, we generally name mammalian E2s according to their relationship to yeast E2s. When this is not possible, we use one of the published names.

2 Historical Background

Ubiquitin’s best-understood function is that of a protein cofactor in an intracellular protein-degradation pathway that terminates with the destruction of Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Ubiquitin-conjugating Enzymes

Fig. 1 The ubiquitin-conjugation pathway. Steps in ubiquitin activation and substrate modification. E1, ubiquitin activating enzyme; E2, ubiquitin-conjugating enzyme; E3, ubiquitin-protein ligase. Atoms involved in the thiol ester and amide bonds are shown.

ubiquitin-tagged substrates by 26S proteasomes [1]. The discovery in 1980 that this 76-residue protein is conjugated to proteolytic substrates through the formation of a peptide-like bond, and in an ATP-dependent manner, suggested that ubiquitin activation would be part of the conjugation process [2, 3]. A ubiquitin activating enzyme (E1) was soon identified and shown to employ an aminoacyl-tRNA-synthetase-like mechanism [4]. E1 first catalyzes the addition of an adenylate moiety to the carboxyl group of ubiquitin’s C-terminal residue, G76. The AMP-bound ubiquitin is then transferred to a cysteine residue in the E1 active site, concomitant with the formation of a new molecule of ubiquitin adenylate. The thiol-linked ubiquitin is the proximal source of activated ubiquitin for downstream steps. From a chemical point of view, the E1/ubiquitin thiol ester should be competent to donate ubiquitin to a substrate amino group. In fact, aminoacyl-enzyme thiol esters are used in exactly this way in non-ribosomal polypeptide synthesis, a process that was discovered around the same time as ubiquitin–protein conjugation [5]. In spite of the attractive simplicity of this model, however, biochemical reconstitution studies showed that besides E1 two additional fractions were required to conjugate ubiquitin to a model substrate. They were called ubiquitin carrier protein (E2) and ubiquitin-protein ligase (E3), respectively, since the respective factors seemed to act sequentially [6]. Interestingly, the E2 factor apparently formed a thiol ester with ubiquitin. Based on these results, Hershko and co-workers proposed the “ubiquitin conjugation cascade” (Figure 1). Multiple thiol ester-forming proteins were present in the E2 fraction [6], but only the smallest of them reconstituted substrate ubiquitination catalyzed by the thenknown E3 [7]. This result suggested that there could be multiple E2s with distinct functional properties. Confirmation of this hypothesis came with the cloning of the first two E2 genes, RAD6/UBC2 and CDC34/UBC3, which indeed encoded homologous yeast proteins with a signature cysteine-containing active-site motif [8, 9]. The two E2s functioned in distinct biological processes – DNA damage tolerance [8, 10] and cell-cycle control [9] – providing the first hint that ubiquitination might regulate a broad range of biological processes. A family of E2 enzymes naturally suggested that there would also be a family of E3 enzymes. This prediction has since been strikingly confirmed. We now know that specific E2/E3 complexes function to modify specific substrates with ubiquitin.

3 What is an E2? 3

3 What is an E2?

A protein can be identified as an E2 enzyme according to several different criteria. Functionally, the E2 occupies an intermediate position in the conjugation cascade – that is, it acts between the E1 and the E3 (Figure 1). This property accounts for the original name of ubiquitin carrier protein, which drew an analogy to the acyl carrier proteins used in fatty acid biosynthesis and non-ribosomal peptide synthesis [6]. Subsequently, with the recognition that E2 enzymes often play an active role in conjugation, the conjugating enzyme name gained favor. Mechanistically, the E2 first participates in a thiol ester transfer reaction, in which the activated ubiquitin is moved from the active-site cysteine of E1 to that of the E2 (Figure 1). The E2/ubiquitin thiol ester intermediate is strictly required for downstream steps, as shown by ablation of substrate ubiquitination following mutation of the active-site cysteine residues of different E2s (see, for example, Refs. [11, 12]). The ubiquitin is then transferred from the E2 active site to the ε-amino group of the substrate’s lysine residue, forming an isopeptide bond. The conjugation site can also be a specific lysine on a previously conjugated ubiquitin, which leads to polyubiquitin chain elongation; chains linked through K48 are the principal signal for targeting substrates to proteasomes [1]. Transfer of ubiquitin to the substrate requires the assistance of the E3 [1, 6]. If this enzyme belongs to the HECT domain family (Homologous to E6AP C-Terminus), the ubiquitin is first transferred to an active-site cysteine residue of the E3; if the E3 belongs to the RING domain family (Really Interesting New Gene), ubiquitin is transferred directly to the substrate’s amino group (Section 6.3). Collectively, these properties constitute the biochemical definition of an E2 enzyme: it is a protein that accepts ubiquitin in thiol ester linkage from E1, and cooperates with an E3 enzyme to deliver this ubiquitin to the substrate. The functional specialization of individual E2s (Section 4) reflects the specificity of interaction of each E2 with its cognate E3(s), in conjunction with the E3’s substrate specificity. Therefore an E2 enzyme can also be defined according to the cognate E3(s) with which it interacts. The E3 partners of many of the eleven ubiquitin-dedicated E2s in Saccharomyces cerevisiae are conserved in higher organisms (Section 4). However, both the E2 and E3 families are much larger in higher organisms than in yeast. Present accounting suggests that there are 50–70 E2s, and hundreds of E3s, in mammals [13, 14]. The amino acids surrounding the thiol ester-forming cysteine residue are particularly highly conserved, but there is sequence similarity throughout the 150-residue E2 core domain (Figure 2). This bioinformatic definition makes it easy to identify E2 genes in sequenced genomes [13, 14]. In fact, many E2s consist of just this core domain (Figure 2). The fact that such E2s are often functionally distinct from one another indicates that modest sequence variation within the core domain can be highly significant. Structural biology has begun to shed light on this structure/function correlation (Section 6). Other E2s display N- and C-terminal extensions to the core

4 Ubiquitin-conjugating Enzymes

3 What is an E2? 5

Fig. 2 Ubc13 (1JBB). Canonical "/$ E2 fold with the active-site cysteine shown in ball-and-stick.

domain (Figure 2), which may play a role in E3 and/or substrate specificity (see Refs. [15–19]). Structural biology provides a final way to define an E2 enzyme. As expected from the strong sequence conservation, the E2 core domain adopts a conserved fold. At the time this article was being prepared, twelve different E2 structures had been deposited in the Protein Data Bank. The average root-mean-square deviation of the 150 Cα positions of these structures is less than 2Å. E2s are α/β proteins containing a central anti-parallel four-stranded β-sheet (S1–S4), four α-helices (H1, H3, H4, H5), and a small 310 helix (H2) (Figure 3) [20, 21]. The cysteine is located on an extended loop after β-strand 4 and immediately before the short 310 helix H2. The active-site cysteine sits in a shallow groove composed of residues from the H3–H4 and S4–H2 loops. The canonical α/β E2 fold is highly versatile, allowing E2 enzymes to associate with several different proteins in the ubiquitin conjugation cascade without any perturbation of the E2’s tertiary structure (Section 6). Residues occupying the face opposite the active site are less conserved than those surrounding the cysteine [21]. Sequence variation in this region contributes to the functional diversity of the E2 family by permitting specific interactions of individual E2s with cognate E3s and (perhaps) substrates.

6 Ubiquitin-conjugating Enzymes

4 Functional Diversity of Ubiquitin-conjugating Enzymes

The functional range of the ubiquitin-conjugating enzyme family is easily appreciated by considering the family members in a single organism. Table 1 summarizes key properties of the complete set of E2s in the yeast S. cerevisiae, including notable structural features, known cognate E3(s) and their key substrates, and biological functions (see also [22, 23]). We cannot give a comprehensive review of E2 functions in higher organisms, but we do comment on some notable instances of functional conservation, expansion, and divergence (see also [23]). 4.1 Functions Related to Proteasome Proteolysis

In many cases, the specific function(s) of a given E2 enzyme reflect its role in targeting one or more substrates for degradation by 26S proteasomes. The scope of this function varies considerably between E2 family members, however. At one extreme, the functionally redundant enzymes Ubc4 and Ubc5 are necessary for the turnover of many substrates, as shown by a marked reduction in the rate of turnover of endogenous short-lived and abnormal proteins in a ubc4ubc5 yeast strain [24]. The slow growth and stress sensitivity of this strain [24, 25] can also be ascribed to inhibition of proteasomal proteolysis since these phenotypes are characteristic of proteasome mutants [22, 26]. Despite the important role of Ubc4/5 in proteasome degradation, few E3 partners relevant to this function have been identified. One is Ufd4, a HECT-domain E3 that mediates the degradation of linear ubiquitin fusion proteins [27]. A ufd4 strain grows normally, however, indicating that Ubc4 has other cognate E3s. Rsp5, an essential HECT-domain E3, is one likely candidate since this E3 partners with Ubc4 in other pathways (see below). UBC1 is essential for viability in the ubc4ubc5 strain, suggesting that Ubc1 shares substantial functional overlap with Ubc4/5 in directing substrates to proteasomes for degradation [28]. The Ubc4/5 sub-family of E2s is much larger in mammals, where it includes both constitutively and selectively expressed enzymes. Notable human E2s in this group are UbcH5a/b/c, UbcH7, and UbcH8 (see Ref. [23]). The expansion is likely to reflect the much larger size of the E3 family in higher organisms. However, while the results of in vitro conjugation assays and protein-protein interaction studies suggest that certain E3s partner specifically with individual Ubc4/5 sub-family members, the degree of E3 (hence, functional) selectivity in the cellular setting remains quite uncertain (discussed in Refs. [23, 29]). RNA interference studies and mouse knockout models may be helpful in addressing this question in the future. Ubc3/Cdc34 supports the proteasome-mediated proteolysis of numerous substrates through its role as the specific E2 partner of a large family of multi-subunit RING E3s called SCF E3s (Skp-Cullin-F-box, Section 6.3). This role is preserved in higher organisms [30]. Yeast ubc3 mutants arrest in G1 phase of the cell cycle because they fail to degrade Sic1 [31], an inhibitor of the G1/S transition that is

4 Functional Diversity of Ubiquitin-conjugating Enzymes 7 Table 1 E2 enzymes of the yeast Saccharomyces cerevisiae

Gene

Amino acids

UBC1

215 [28]

Cognate E3

Unknown

Hrd1 [39] UBC2 (RAD6)

172 [8]

Ubr1 [156]

Ubr1 [52]

Rad18 [73], [157] Bre1 [67, 69]

UBC3 (CDC34)

295 [9]

SCF E3s

UBC4

148 [24]

Unknown Doa10 Rsp5

UBC5

148 [24]

UBC6

250 [46]

See UBC4

Unknown

Doa10 [42] Doa10 [42] UBC7

165 [25] Hrd1 [39]

Doa10 [42] UBC8

206 [160]

Unknown

UBC10

165 [75]

Unknown

Functions and substrates Short C-terminal tail harbors ubiquitin-associated (UBA) domain [155] Essential in ubc4ubc5 genetic background, suggesting a redundant role with Ubc4/5 in proteasomal turnover of short-lived and abnormal proteins [28] Role in ERAD that is not fulfilled by Ubc4/5 [39, 40] Proteasomal degradation of N-end rule [158] substrates, including cohesin fragment [51] Proteasomal degradation of Cup9 transcriptional repressor regulates peptide import DNA-damage tolerance [8] via monoubiquitination of PCNA [65] (non-proteolytic function) Ubiquitination of histone H2B [68] regulates gene transcription and silencing [71] (non-proteolytic function) Essential gene; long C-terminal tail; targets diverse substrates for proteasomal degradation [30, 34, 36]; regulation of cell-cycle progression Proteasomal degradation of diverse shortlived proteins [24] Proteasomal degradation of MATα2 transcriptional repressor [49] Endocytosis of membrane proteins [60, 61]; protein trafficking (see Ref. [63]) 92% identical to Ubc4; functionally redundant [24] C-terminal tail provides anchoring to ER membrane [46] Together with Ubc7, proteasomal degradation of some ERAD substrates [47, 48, 159] Proteasomal turnover of Ubc6 is Ubc6-, Ubc7-, and Doa10-dependent [42, 50] In conjunction with Ubc7, proteasomal turnover of MATα2 Localized to ER membrane via Cue1 [37] Role in proteasomal degradation via ERAD [38, 48] confers resistance to cadmium and other ER stresses [25] In conjunction with Ubc7, proteasomal turnover of MATα2 Glucose-induced proteasome degradation of fructose-1,6-bisphosphatase [57] Also called Pas2/Pex10. Peroxisome biogenesis [75]; Pex10 is a candidate E3 [78]

8 Ubiquitin-conjugating Enzymes Table 1 (continued)

Gene

Amino acids

Cognate E3

Functions and substrates

UBC11

156 [84]

Unknown

UBC13

153

Unknown; similar to clam E2-C (E2-C functions in mitotic cyclin turnover [79], but Ubc11 is dispensable for this process in yeast [84]) Heterodimerizes with Mms2 (UEV) [72] DNA-damage tolerance [72] via polyubiquitination of PCNA [65] (non-proteolytic function) Essential gene; E2 dedicated to Smt3 (SUMO) [163] Septin modification E2 dedicated to Rub1 (Nedd8) Modification of specific cullin lysine residue activates cullin-based E3s [165]

Rad5 [73]

UBC9

157 [161]

UBC12

188 [164]

Siz1/2 [162] SCF E3s

recognized and polyubiquitinated by a specific SCF E3 [32, 33]. Although this is the only essential function of yeast Ubc3 [31], this E2 partners with many other SCF E3s to target diverse substrates for degradation by proteasomes (see Refs. [30, 34–36]). Although studies in yeast suggest that Cdc34 is the main E2 partner of SCF E3s, some SCF E3s seem to partner with Ubc4/5-type E3s (see Refs. [23, 36]). Ubc3 has a long C-terminal tail (Table 1), making it the most distinctive yeast E2 in terms of primary structure. A chimeric E2 in which the Ubc3 tail is appended to the core domain of Ubc2 fulfills the essential function of Ubc3 in yeast, suggesting that the tail of Ubc3 is necessary for key interactions with the E3 or Sic1 [15, 16]. Ubc7 is localized to the endoplasmic reticulum (ER) through an interaction with a partner protein, Cue1 [37], and plays a major role in proteasome degradation. Ubc7 acts on misfolded proteins of the ER, which are ejected from this compartment as a prelude to ubiquitination at the cytosolic face of the ER membrane and degradation by cytosolic proteasomes [38]. Ubc7’s role in ERAD (ER Associated Degradation) explains why a ubc7 strain is conditionally sensitive to agents that induce protein misfolding in the ER [25, 39–41]. Ubc7 frequently partners with Hrd1, an ERlocalized RING E3 [39], but some ERAD substrates of Ubc7 seem to be recognized in cooperation with a different ER-localized RING E3, Doa10 [42]. Consistent with Ubc7’s prominent role in ERAD, the yeast UBC7 and CUE1 genes are induced as part of the Unfolded Protein Response (UPR) and there is a synthetic lethal relationship between certain ERAD and UPR genes [40, 41]. Yeast Ubc1 also plays a significant, but poorly-defined, role in ERAD [39, 40]. Mammalian Ubc7 also functions in ERAD [43–45]. Ubc6 localizes to the ER through its own C-terminal membrane anchor [46]. Although Ubc6 plays a role in ERAD, its function in this process is less conspicuous than that of Ubc7 [43, 47, 48]. Interestingly, Ubc6 and Ubc7 both contribute to the Doa10-dependent degradation of a soluble nuclear protein [42, 49], and Ubc6 is

4 Functional Diversity of Ubiquitin-conjugating Enzymes 9

itself rapidly degraded by proteasomes in a manner that depends on its own activesite cysteine, its C-terminal membrane anchor, functional Ubc7, and Doa10 [42, 50]. The purpose of this instability remains mysterious. Ubc2 functions rather selectively in proteasome proteolysis. In yeast, two specific E3 partners are known, leading to proteasome degradation events that regulate chromosome stability [51], peptide import [52], and homing endonuclease stability [53]. Mammals have two closely-related Ubc2 isoforms, each of which complements most of the functions of the yeast ubc2 strain [54]. But the mammalian Ubc2 isoforms also have specialized functions – one of them is required for spermatogenesis in the mouse [55] and at least one of them can be inferred to be necessary for cardiovascular development [56]. So far, Ubc8 has been implicated in the regulated turnover of just one substrate, and its E3 partner(s) remain unknown [57]. Interestingly, the closest mammalian relative of yeast Ubc8 is expressed with a restricted tissue specificity and (in some tissues) in a regulated manner [58, 59]. 4.2 Endocytosis and Trafficking

Just because an E2 functions in proteasome proteolysis does not mean that its functions are limited to this pathway. This is because the E2’s functional range is largely determined by the substrate specificity of its E3 partner(s). For example, yeast Ubc4 and Ubc5 play a prominent role in proteasome degradation, but they also cooperate with a HECT E3, Rsp5, to mono-ubiquitinate certain plasma membrane receptors [60–62]. This modification signals receptor endocytosis, leading to degradation in the vacuole (equivalent to the mammalian lysosome). Ubc1 is partially redundant with Ubc4/5 in endocytosis [61, 62], as seen in ubiquitination reactions leading to proteasome degradation (above). Thus, Ubc1, 4, and 5 may be able to substitute for one another in complexes involving many different E3s, probably reflecting the strong conservation of the core domain between Ubc1 and Ubc4/5. Ubc4 and Ubc5 may also act with Rsp5 to regulate protein trafficking downstream of endocytosis (reviewed in Ref. [63]). 4.3 Non-proteolytic Functions

As mentioned in Section 2, Ubc2 is the defining player in a conserved DNA damagetolerance pathway [8, 10, 64]. Here Ubc2 partners with a RING E3, Rad18, to modify a DNA polymerase processivity factor with a single ubiquitin [65]. This modification signals error-prone bypass of DNA lesions [66]. Ubc2 partners with a different RING E3 (Table 1) to mono-ubiquitinate histone H2B [67–69]. This modification promotes histone methylation, which in turn regulates transcription and silencing [70, 71]. Ubc13 participates in the same DNA damage-tolerance pathway as Ubc2 [72]. Ubc13 collaborates with two enzyme partners (Table 1) to modify the DNA polymerase cofactor (above) with a K63-linked polyubiquitin chain, which signals

10 Ubiquitin-conjugating Enzymes

error-free lesion bypass [65, 72, 73]. In higher organisms, Ubc13 also helps to synthesize K63-linked polyubiquitin chains in a second non-proteolytic signaling pathway (see Ref. [74]). The function and mechanism of Ubc13 are discussed in more detail below (Section 7). 4.4 E2s of Uncertain Function

Ubc10 is required for the biogenesis of the peroxisome, an oxidative organelle [75]. This E2 plays a role in peroxisomal protein import [76] and is recruited to the peroxisomal membrane through an interaction with a partner protein [77]. Membrane-localized Ubc10 also seems to be spatially proximal to Pex10, which has a RING-like domain [78]. Whether Pex10 is a cognate E3 of Ubc10 remains to be determined, as does the mechanistic role of ubiquitin conjugation in peroxisome biogenesis. The function of the remaining yeast E2, Ubc11, remains uncertain. Ubc11 is very similar to E2-C, a clam E2 that acts with an essential multi-subunit RING E3, the anaphase promoting complex (APC) or cyclosome, to ubiquitinate mitotic cyclins [79]. This reaction leads to cyclin degradation by proteasomes, which drives exit from mitosis [35, 80]. Mitotic cyclin ubiquitination can be reconstituted in vitro with apparent amphibian and fission yeast orthologs of either Ubc11 or Ubc4 [81, 82] and other data implicate both E2s in this process in higher cells [82, 83]. However, mitotic cyclins are efficiently degraded in budding yeast ubc4 and ubc11 strains, indicating that other E2 enzymes can support this essential function in S. cerevisiae [84]. 4.5 E2 Enzymes and Disease

There are now several striking examples of disease-related defects in ubiquitin conjugation, but most of them involve E3s rather than E2s. This is not surprising given the paramount role of E3s in substrate selection and the corresponding intensity of research effort that has been focused on E3s. Still, there are several hints that defects at the E2 level of the conjugation cascade can also contribute to disease. Many viruses subvert the ubiquitin system to evade the host cell’s defenses or modulate the cellular environment so as to promote viral replication (see Refs. [85, 86] and Section 7). The genome of African swine fever virus encodes an E2 enzyme that is somewhat similar to yeast Ubc3 [87, 88]. This enzyme might alter the activity or specificity of the host cell’s conjugation cascade so as to benefit the virus, or it could act on specific viral proteins. Herpes Simplex Virus-1 (HSV-1) encodes an E3 enzyme that specifically binds the host cell’s Ubc3/Cdc34 enzyme and targets this E2 for ubiquitination and (presumably) degradation – events that may help to stabilize specific cyclins and promote viral replication [89]. A different kind of relationship between an E2 enzyme and disease is exemplified by the finding that the Alzheimer’s amyloid-β peptide induces the expression of

6 The Biochemistry of E2 Enzymes

E225K , a mammalian relative of yeast Ubc1 [90]. E225K was found to play a major role in amyloid-β-dependent neuronal cell killing. This effect may be related to the E225K -dependent production of aberrant polyubiquitin chains, leading to the inhibition of proteasomes [90, 91]. Other studies showed that the human homolog of yeast Ubc11 is over-expressed in numerous cancer cell lines and primary tumors and that forced over-expression of this E2 in cultured cells can drive proliferation and transformation [92]. Similarly, transformation and chromosomal abnormalities were observed following over-expression of human Ubc2b [93]. Such disease-related over-expression effects could arise in two different ways. The higher E2 concentration could lead to a relaxation of specificity – that is, pairings with non-cognate E3s – leading to inappropriate ubiquitination events. Alternatively, specificity could be maintained, but an inappropriately high flux through the normal E2/E3 pathway could lead to the excessive ubiquitination of cognate substrates.

5 E2 Enzymes Dedicated to Ubiquitin-like Proteins (UbLs)

Ubiquitin is just one member of a family of protein modifiers that share a common fold and a common mechanism of isopeptide tagging [70, 94, 95]. Like ubiquitin, individual UbLs are activated at a C-terminal glycine residue by a specific E1 enzyme. Often, the next step is transfer to a specific E2 enzyme. Certain UbL-specific E2 enzymes are so similar to ubiquitin-conjugating enzymes that they were initially thought to be members of the ubiquitin-conjugating enzyme family. This was true of yeast Ubc9 and Ubc12, which are dedicated to Smt3/SUMO and Rub1/Nedd8, respectively (Table 1). SUMO modifies numerous cellular proteins and has a broad functional range [94], but the only known target of Nedd8 is a specific lysine residue in one subunit (the cullin) of SCF E3s. Nedd8 modification activates these E3s (see Ref. [70]). The reader should consult earlier reviews [70, 94, 95] for a detailed discussion of UbL biology and biochemistry. There are two important points for the current discussion. First, the conjugation cascades of UbLs differ from that of ubiquitin chiefly in terms of complexity – there is one conjugating enzyme per UbL, and many fewer E3s. Second, because modifier proteins (including ubiquitin) do not interact strongly with their dedicated E2s (Section 6.1), it is believed that E1 enzymes play the major role in matching E2s with the correct modifier protein (see Ref. [96]).

6 The Biochemistry of E2 Enzymes 6.1 E1 Interaction

An E2 needs to associate with several different proteins in the course of the ubiquitin conjugation cascade, with the first being the E1. Mutational studies conducted with

11

12 Ubiquitin-conjugating Enzymes

the SUMO-specific E2 Ubc9 suggest that free Ubc9 associates with its free cognate E1 through a surface of Ubc9 that includes the C-terminal residues of α-helix H1 and residues in a loop between β-strands S1 and S2 (Figure 3) [97]. This surface of Ubc9 is also important for thiol ester bond formation [97]. Several residues in α-helix H1, particularly the C-terminal residues, are poorly conserved among E2s, and Ubc9 contains a five-residue insertion in the loop between β-strands S1 and S2 (Figure 2). Thus, this region of E2s may contribute to specificity for their cognate E1s. Consistent with this idea, the N-terminal helix (H1) of a ubiquitin E2 was found to be important for E2/ubiquitin thiol ester formation [98]. One cautionary note is that Ubc9 displays a substantial affinity for its free E1 [97], whereas ubiquitin E2s bind tightly to their E1 only after it has been loaded with ubiquitin [6, 99, 100]. The structural basis of this effect remains to be determined. 6.2 Interactions with Thiol-linked Ubiquitin

As a consequence of interacting with ubiquitin-loaded E1, the E2 accepts the activated ubiquitin at its active-site cysteine residue. This thiol ester complex, although biochemically detectable [6], has not been crystallized because it is labile in comparison to the requirements of structural biology. However, NMR chemical shift perturbations have been used to map the binding surface of ubiquitin onto human Ubc2b [101], yeast Ubc1 [102], and human Ubc13 [103]. All three models map the ubiquitin-binding surface of the E2 to a common area that includes parts of α-helix H3, the loop between α-helices H3 and H4, and residues around the active-site cysteine in the extended S4-H2 loop, including part of the 310 helix H2 (Figures 2 and 4). The C-terminus of ubiquitin extends around part of the E2 and is constrained in a cleft [102, 103]. Although this contact surface is detectable in the thiol ester, free E2s display a negligible affinity for free ubiquitin [6]. Thus, the covalent E2/ubiquitin bond enables the formation of these non-covalent contacts. 6.3 E3 Interactions

After E2/ubiquitin thiol ester formation, the ubiquitin must be transferred to the substrate, which is sometimes another ubiquitin. An E3 is usually required for this reaction in vitro, and is always required in vivo. There are three known types of E3s: the RING domain, HECT domain, and U-box (UFD2 homology) families. RING and U-box E3s act as bridging factors for E2s and substrates, but HECT E3s use a different mechanism, adding an extra step to the pathway (Section 6.3.3). 6.3.1 RING E3/E2 Interactions The small RING domain coordinates two zinc ions in a cross-brace arrangement [104]. The domain is defined by the presence of eight zinc-binding groups (cysteines and histidines) with a conserved spacing, such that the distance between the two zincs is conserved at 14 Å [104]. Sequence conservation between RING domains is

6 The Biochemistry of E2 Enzymes

Fig. 3 Ubc1/ubiquitin thiol ester complex model (1FXT). The surface of Ubc1 is shown with residues implicated in ubiquitin binding colored purple and the active-site cysteine colored yellow. Ubiquitin is colored green.

otherwise minimal. The RING-domain fold consists of a central α-helix and several small β-strands separated by loops with variable lengths [105]. RING E3s can be either single-subunit or multi-subunit enzymes. The crystal structure of UbcH7 complexed to a single-subunit RING E3, c-Cbl, shows that the RING domain is the main site of contact, although there are a few intermolecular hydrogen bonds to a non-RING helix of the E3 [106] (Figure 5). The structure of the E2 in the c-Cbl/UbcH7 complex is unchanged relative to free E2 structures. The main basis for the interaction is the packing of several hydrophobic residues of UbcH7, notably F63, into a shallow groove on the RING domain surface. These residues come from the S3–S4 and H2–H3 loops (Figure 3). The α-helix H1 of UbcH7 makes the hydrogen-bond contacts to the non-RING α-helix. Interestingly, even though the c-Cbl/UbcH7 structure undoubtedly shows conserved RING/E2 contacts, this complex is catalytically inactive (cited in Ref. [107]). Therefore additional E2/RING contacts may be needed for catalytic competence. The surface of UbcH7 that contacts c-Cbl does not overlap with the E2 surface that contacts ubiquitin (Figures. 4 and 5; see also Ref. [108]), confirming that the

13

14 Ubiquitin-conjugating Enzymes

Fig. 4 UbcH7/c-Cbl complex (1FBV). The surface of UbcH7 is shown with residues interacting with the c-Cbl RING domain shown in red and the active-site cysteine shown in yellow. c-Cbl is colored green.

E2/ubiquitin thiol ester can associate with a RING E3. The E2 surface that contacts cCbl does, however, overlap the E2 surface implicated in E1/E2 interactions (Section 6.1). Thus, the E1 may have to depart from the E2/ubiquitin complex before E2/E3 interactions can take place. The closest approach of a RING-domain residue to the active-site cysteine of UbcH7 is about 15 Å, arguing against a role for RING E3s in chemical catalysis [106]. Instead, RING E3s have been proposed to facilitate ubiquitination by inducing physical proximity of the E2/ubiquitin thiol ester and the substrate [23, 30, 106, 109]. Catalysis would result from the increased local concentrations of the two reactants (discussed further below). RING/E2 interactions have also been studied with BRCA1. This E3 is unique in that it must heterodimerize with a second RING-domain protein, BARD1, in order to display maximal E3 activity [110]. Even though the heterodimer interface leaves both RING domains available for interaction [110], UbcH5c binds exclusively to the BRCA1 RING [107]. The interacting surface of UbcH5c is homologous to the surface of UbcH7 that contacts the c-Cbl RING domain in that several residues of UbcH5c pack into a cleft on the BRCA1 RING domain. But UbcH5c also makes several

6 The Biochemistry of E2 Enzymes

contacts with the C-terminus of the BRCA1 RING domain and with a non-RING region of the heterodimer [107]. These extra contacts are not observed when UbcH7 binds to the BRCA1/BARD1 complex [107]. Because the BRCA1/BARD1/UbcH7 complex is inactive, the extra contacts observed with UbcH5c may help to create a competent E2/E3 complex. Despite the greater complexity of multi-subunit RING E3s, a common theme is evident – all SCF E3s, as well as several other types of cullin-based E3s, utilize a common RING-domain subunit, the small protein Rbx1 (reviewed in Refs. [30, 111]). Four subunits compose the minimal SCF E3 ligase complex: a cullin scaffold, Rbx1, an adaptor protein (Skp1), and a substrate-binding subunit that connects to the adaptor through a conserved domain called the F-box (see Ref. [30]). The cullin acts as a scaffold, with Rbx1 binding to one end to form a cullin/Rbx1 subcomplex that recruits the E2 and, in many cases, displays a substrate-independent ubiquitinligase activity (see Refs. [23, 30]). The crystal structure of the mammalian SCFSkp2 (Skp1/Cul1/F-boxSkp2 /Rbx1) E3 ligase shows a remarkably rigid, elongated complex [112]. The Cul1 scaffold contains three cullin-repeat motifs that span 110Å, with Rbx1 binding to a discrete C-terminal α/β domain. The Skp1/F-boxSkp2 complex binds to the opposite (Nterminal end) of the cullin. Rbx1 displays a hydrophobic groove, as seen previously in the c-Cbl RING domain [106, 112]. In c-Cbl, this groove provides an interaction surface for UbcH7 and it is reasonable to assume a similar mode of interaction in the case of Rbx1. Interestingly, the site where Nedd8 modifies the cullin is close to where the E2 binds, consistent with data which suggest that neddylation modulates E2 binding or activity [113, 114]. A model of the full SCF/E2 complex [112] shows that the end of Skp2 which binds the substrate is pointed toward the Rbx1-bound E2, with a 50-Å gap between the two. Models based on two other SCF structures show similar distances between the F-box protein and the E2 [109, 115]. Whether this gap can be bridged by the bound substrate is currently unclear. It has been suggested that the E2 may bind to Rbx1 somewhat differently than UbcH7 is observed to bind in the c-Cbl RING/UbcH7 complex, but it is not obvious that this can lead to a 20 Å movement of the E2 toward the bound substrate as suggested [109]. One could also imagine that the bound substrate and E2 “meet” through conformational changes of the SCF complex. However, the rigid separation produced by the Cul1 scaffold seems to be important for activity – introducing a flexible linker into the center of Cul1 produced a protein that could still bind an E2, but did not catalyze substrate ubiquitination [112]. An interesting study established a positive correlation between the rate of dissociation of the Ubc3/ubiquitin intermediate from the RING domain and the rate of Sic1 ubiquitination catalyzed by SCFCdc4 [116]. The authors proposed that the role of the RING domain is to bring the charged E2 into the vicinity of the SCF-bound substrate, but that release of the charged E2 is important to bridge the gap and enable multiple substrate lysines to be targeted. However, although these mechanisms may place the substrate’s lysine residue in the general vicinity of the E2’s active site, it is unclear that they can establish an effective orientation of the lysine residue and the thiol ester bond. Since

15

16 Ubiquitin-conjugating Enzymes

there is no known consensus site for ubiquitination [23, 117, 118], it is unlikely that specific molecular contacts in the vicinity of the substrate’s lysine residue are used to position this attacking group. Overall, it remains unclear how the substrate’s lysine residue approachs the E2 active site. 6.3.2 U-box E3/E2 Interactions The U-box family of E3s bind E2s through the small U-box domain [119]. Some U-box E3s do not seem to have their own cognate substrates, but instead promote polyubiquitination of the substrates of other E3s [120]. Other U-box E3s have defined cognate substrates and behave in a canonical manner [121, 122]. An NMR structure [123] confirmed an earlier prediction [124] that the Ubox domain has a RING-domain-like fold. Remarkably, the U-box domain uses hydrogen-bonding networks in place of zinc coordination to support the characteristic cross-brace arrangement. These interactions stabilize a globular fold consisting of a central α-helix surrounded by several β-strands, which are separated by loops of variable length [123]. There is a shallow groove in the surface located in a position homologous to the E2-interacting surface of RING domains. Mutational studies have linked E3 ligase activity to some of the residues in the surface groove [123, 125]. Since these mutations do not disrupt the U-box fold, they are likely to abrogate E2 binding. Although it is likely that E2s bind similarly to the U-box and RING domains, no E2/U-box structure has been reported so far. 6.3.3 HECT E3/E2 Interactions HECT-domain E3s are defined by the presence of a domain of 350 residues that is homologous to the C-terminus of the founding family member, E6AP (E6 Associated Protein [126]). E6AP is known for its role in binding the E6 protein of oncogenic human papilloma viruses and targeting the p53 tumor suppressor for ubiquitination and degradation [127]. HECT-domain E3s possess an active-site cysteine residue positioned 35 residues upstream of the C-terminus; a thiol ester with ubiquitin is formed at this site and is required for substrate ubiquitination [128]. The crystal structure of an E6AP/UbcH7 complex showed that the HECT domain is L-shaped, with a large, mostly α-helical, N-terminal lobe and a small C-terminal lobe with an α/β structure [108] (Figure 6). UbcH7 binds to the end of the Nterminal lobe and somewhat parallel to the C-terminal lobe, forming an overall U-shaped complex. UbcH7 binds in a large hydrophobic groove in the N-terminal lobe [108]. As seen in other E2/E3 structures, neither the E2 enzyme nor the HECT domain changes its overall fold upon binding. UbcH7 contacts its binding groove with residues from the S3–S4 loop and the H2–H3 loop (Figure 3). A few contacts are also made with the C-terminal portion of α-helix H1. Remarkably, these are the same two loops and helix that bind the RING domain in the c-Cbl/UbcH7 structure [106]. That two different E3s contact a largely similar surface on UbcH7 (Figures 5 and 6) can be explained through the nature of key side-chain contacts. The S3–S4 loop seems to provide most of the specificity, as it contains the F63 residue that is present in all E2s that are known to bind both

6 The Biochemistry of E2 Enzymes

Fig. 5 UbcH7/E6AP (1C4Z). The surface of UbcH7 is shown with residues interacting with the E6AP HECT domain shown in blue and the active-site cysteine shown in yellow. E6AP is colored green with the active-site cysteine between the N- and C-lobe shown in yellow.

HECT E3s and c-Cbl [106, 108]. In c-Cbl, the main contacts for F63 are isoleucine and tryptophan residues located in the RING groove [106]. Both F63 and its contact site in c-Cbl are seen to vary in other E2/RING E3 pairs, suggesting that interactions between these three specific residues are needed, but that the nature of the contact can vary [106]. In other words, a different E2 could bind to a different E3 with a similar geometry, but through different types of side-chain contacts. UbcH7-F63 makes specific contacts with six E6AP residues, so this interaction is likely to be important for all HECT/E2 pairs [108]. The H2-H3 loop that makes the other major contacts with the HECT domain is part of the more variable E2 surface (Section 3). The specific contacts made between the H2-H3 loop of UbcH7 and the E6AP HECT domain could be used to correctly predict the E2 preferences of E6AP and Rsp5 [108]. Thus, residues in the S3-S4 and H2-H3 loops play an important role in determining the specificity of E2/E3 interactions. Positioned near the bend between the two lobes of the HECT domain is its activesite cysteine [108]. This residue is 41 Å away from the active-site cysteine of UbcH7 (Figure 6), suggesting that a large conformational change is needed to bring about

17

18 Ubiquitin-conjugating Enzymes

transfer of ubiquitin from E2 to E3. Such a mechanism was confirmed in the crystal structure of another HECT domain [129]. The WWP1-HECT domain resembles the E6AP HECT domain in having two lobes, but their relative positions differ. In the WWP1-HECT domain the two lobes form an inverted T-shape as opposed to the Lshape seen in the E6AP HECT structure [108, 129]. This conformational change can be brought about by a rotation around three residues in a hinge loop connecting the two lobes. Modeling in the E2 in a position homologous to that seen in the E6AP/UbcH7 structure, the distance between the two active-site cysteines decreases to 16 Å [129]. With additional rotation around the hinge loop the WWP1 activesite cysteine can be brought within 5 Å of the E2’s active-site cysteine. Mutational studies suggested that the flexibility of this hinge loop is indeed important for ligase activity [129]. This flexibility requirement points to possible models for ubiquitin transfer and polyubiquitin chain elongation. One possibility is that the HECT domain adds ubiquitin to target substrates one at a time. This would imply that the E3 changes specificity – from recognizing the substrate to recognizing the ubiquitin – following transfer of the first ubiquitin to the substrate. A different possibility is that the chain is built up by the HECT E3 first, and then transferred as a unit to the substrate. This would require two active sites, one to hold the growing polyubiquitin chain, and the other to hold the next ubiquitin to be added. HECT E3/E2 complexes would satisfy this condition. RING E3/E2 complexes cannot, and thus would have to utilize another mechanism, presumably building on their rigid architecture. In an attractive model [129], the HECT cysteine could hold the first ubiquitin (and later the growing chain), while the C-lobe could rotate to position the first ubiquitin’s target lysine near the thiol ester bond of the bound E2/ubiquitin intermediate. Subsequent rounds of ubiquitin addition to the chain terminus would require the C-lobe to keep rotating, ultimately ending in steric problems for the chain which could favor its transfer to the substrate. A third general model for polyubiquitin chain extension is that the initiation and elongation phases of the reaction involve different E2 enzymes, different E3 enzymes, or different E2/E3 complexes. The modification of a substrate with a noncanonical polyubiquitin chain follows the third model. The Rad6/Rad18 complex ligates the first ubiquitin, while the Mms2/Ubc13/Rad5 complex extends the chain [65, 73]. The extension of K48-linked chains from ubiquitin fusion proteins seems to involve the sequential action of two different E3s with the same E2 [120]. In another possible example, two E2s (orthologs of Ubc11 and Ubc4) have been suggested to act sequentially with the APC in the polyubiquitination of mitotic cyclins in fission yeast [82]. 6.4 E2/Substrate Interactions

With HECT domain E3s, all of the chemistry of isopeptide bond formation occurs at the E3 active site. With RING and U-box E3s, however, the E2 participates directly in this chemical reaction, so the substrate’s lysine must closely approach

6 The Biochemistry of E2 Enzymes

the E2’s active site. A crystal structure of the SUMO E2 Ubc9, complexed with a large fragment of RanGAP1 (an efficient sumoylation substrate), reveals the specificity of this interaction [130]. Unlike ubiquitination, sumoylation is site-specific. The target lysine for sumoylation lies within a tetrapeptide sequence motif -K-X-D/E, where is a hydrophobic residue, K is the target lysine, and X is any residue. Ubc9 makes specific interactions with each of these consensus-motif residues in a manner that places the lysine ε-amino group within 3.5 Å of the Ubc9 active-site cysteine [130]. The lysine approaches the cysteine from what is expected to be its unencumbered (by SUMO) side [102]. The interacting surface on Ubc9 involves α-helix H4, the loop preceding it, and the extended S4-H2 loop, including the active-site cysteine [130] (Figure 3). This surface does not overlap with the presumptive binding surface for SUMO. This mode of interaction is unlikely to hold with ubiquitin E2s, since no general consensus site for ubiquitination is known. A model for ubiquitin E2/substrate interactions has also been proposed for the special case in which the substrate is ubiquitin [131]. The crystal structure of the Mms2/Ubc13 complex led to the modeling of an E2/UEV/ubiquitin (donor)/ubiquitin (acceptor) model. As discussed in Section 7, UEV (Ubiquitin E2 Variant) proteins such as Mms2 are homologous to E2s, but lack the active-site cysteine residue. Known UEV/Ubc13 complexes act as E2 enzymes specialized for the synthesis of K63-linked polyubiquitin chains [72, 132]. In the model [131], Ubc13 is bound to the donor ubiquitin through a thiol ester bond in a manner that agrees well with inferences from NMR analysis of the Ubc1/ubiquitin thiol ester [102] (Figure 7). The position of the non-covalently bound acceptor ubiquitin is determined by the orientation of Mms2 on Ubc13 (Figure 7). The acceptor ubiquitin has its K63 side chain placed to enter the active site of Ubc13 to form a diubiquitin conjugate. The model suggests that K63 of ubiquitin is selected as the conjugation site through steric exclusion of other lysines, as determined by an interaction between Mms2 and a region of ubiquitin that is distant from K63 [131]. Recent NMR studies have confirmed and refined this model [103]. Thus, the substrate lysine is presented to the active-site cysteine through an indirect mechanism, in contrast to the Ubc9/RanGAP1 example in which the E2 interacts directly with the lysine residue itself [130]. Unlike most ubiquitination reactions, the modification of ubiquitin itself is often site-specific. The Mms2/Ubc13/ubiquitin model can help to explain this phenomenon. 6.5 E2 Catalysis Mechanism

Chemical catalytic mechanisms in the ubiquitin conjugation cascade have proved difficult to decipher. The reactions leading to E2/ubiquitin thiol ester and isopeptide bond formation would be facilitated by electrostatic stabilization of the oxyanion and deprotonation of the attacking amino group (isopeptide bond formation) by a general base [133, 134]. However, while the sequence conservation around the E2 active site is very high (Figure 2), all E2 structures show a lack of candidate catalytic residues close to the cysteine (see Refs. [23, 135]). Although catalytic residues could

19

20 Ubiquitin-conjugating Enzymes

Fig. 6 Ubc13 interaction surface. The interacting surfaces have been mapped onto Ubc13. The active-site cysteine is shown in yellow. Colored surfaces contact: covalently bound ubiquitin (purple); RING domains (red); E1 (presumptive, green); acceptor ubiquitin involved in K63-linked polyubiqutin-chain synthesis (blue).

be contributed by other enzymes in the cascade or by the E2 backbone, structural data argue strongly against a chemical catalytic contribution where it may be needed most – in reactions involving RING and U-box domain E3s. Recent studies [136] addressed the role of a strictly conserved asparagine positioned just upstream of the active-site cysteine (N79 in Ubc1 numbering, Figure 2). In existing E2 structures the asparagine is hydrogen-bonded to the backbone or a side chain. It is distant from the E3 contact surface and, as expected, it is dispensable for E2 binding to RING domain E3s. However, the asparagine is critical for E2-catalyzed and RING E3/E2-catalyzed ubiquitin conjugation reactions. The similar effect of asparagine mutation on the two types of conjugation reactions is reasonable given that E2s do not experience structural perturbations upon binding to E3s (above). The data suggest that an intrinsic catalytic role of the asparagine side chain is brought into play through RING-mediated recruitment of the catalytically competent E2/ubiquitin thiol ester. The asparagine is dispensable for upstream and downstream thiol transfer reactions, suggesting that catalytic residues for these reactions may be located in the E1 and HECT E3 active sites. A specific proposal for the role of the asparagine was developed in a model which breaks the hydrogen bonds to the backbone and rotates the asparagine toward the active-site cysteine [136]. Molecular modeling suggested that the asparagine can be positioned to donate a hydrogen bond to the oxyanion (Figure 8) [136]. Many

7 Functional Diversification of the E2 Fold 21

Fig. 7 Model for catalytic role of E2 activesite asparagine. The side chain of the asparagine in the conserved “HPN” motif (Figure 2) stabilizes the oxyanion that

forms when the substrate’s lysine attacks the E2/ubiquitin thiol ester bond. N79 is numbering for Ubc1 (Figure 2).

cysteine proteases, including deubiquitinating enzymes, use an amide side chain in this manner [134, 137–139]. Structural studies of a deubiquitinating enzyme have shown that the entry of ubiquitin into the active site causes a histidine and an asparagine to shift their positions so that the histidine becomes the general base and the asparagine provides the oxyanion hole [137]. Similarly, ubiquitin binding in the E2 active site could be a trigger that repositions the asparagine. So far, no general base is evident in E2s, but this group may not be needed due to the lability of the thiol ester bond.

7 Functional Diversification of the E2 Fold

Increasing evidence suggests that evolution has used (and is using) the E2 fold for new purposes. In one apparent example of functional expansion, E2 core domains have been observed to be embedded within much larger polypeptide chains [140, 141]. The functional properties of these massive E2s remain poorly characterized, and it is likely that more of them will be discovered. But the clearest case of functional diversification is provided by the UEV proteins. UEVs are related to E2s in their primary, secondary, and tertiary structures, but they lack an active-site cysteine residue and therefore cannot function as canonical E2s [142]. Nonetheless they play several different roles in ubiquitin-dependent pathways. Mms2 and its close (mammalian) relative Uev1a form heterodimers with Ubc13 [72, 132]. Each complex plays a key role in the synthesis of K63-linked chains, which act as non-proteolytic signals in different cellular pathways. The Mms2/Ubc13 complex participates in the UBC2/RAD6-dependent DNA damage tolerance pathway by polyubiquitinating the DNA polymerase processivity factor called PCNA (Proliferating Cell Nuclear Antigen) [65, 72]. To be activated for this pathway, PCNA is first mono-ubiquitinated by the Rad6/Rad18 complex, and then modified

22 Ubiquitin-conjugating Enzymes

with a K63-linked polyubiquitin chain by the Mms2/Ubc13/Rad5 complex (Rad18 and Rad5 are RING E3s) [65, 73]. The Mms2/Ubc13 complex has a core ubiquitin polymerization activity [72]. Rad5 might stimulate this activity [132] or target the Uev/E2 complex to PCNA [73], or both. The related human Uev1a/Ubc13 complex is involved in NFκB signal transduction [132]. It plays an intermediate role in the signaling cascade that starts with a proinflammatory cytokine signal and culminates in the nuclear translocation of the active NFκB transcription factor. In this pathway the Ubc13/Uev1a complex modifies a RING E3, Traf6, with K63-linked polyubiquitin chains [132]. This modification is linked to Traf6 oligomerization. It instigates a cascade of kinase reactions ultimately cause the ubiquitination and degradation of NFκB’s inhibitory partner, IκBα [74, 143]. The crystal structure of Mms2 has been solved alone and in complex with Ubc13 [131, 144]. The overall fold is similar to that in E2s, containing a central fourstranded anti-parallel β-sheet surrounded by α-helices. Differences include the absence of the C-terminal α-helix H5 in the shorter Mms2 protein. The helical Nterminus of Mms2 is also extended compared to Ubc13, and this region plays the major role in heterodimer formation. Two ubiquitins can bind to the heterodimer (Section 6.4) and the surfaces they contact do not overlap with the surface contacted by Rad5 [131, 145]. Ubiquitin plays a crucial role in a protein-trafficking pathway that delivers specific cargo proteins to regions of the late endosome membrane that invaginate into the lumen, thereby targeting these proteins to the vacuole/lysosome (reviewed in Ref. [146]). A different UEV protein, called Tsg101 in humans (Tumor Susceptibility Gene) and Vps23 in yeast (Vacuolar Protein Sorting), is part of a large complex that plays a critical role in the sorting step. Cargo proteins are selected based on their conjugation to mono-ubiquitin; the specific role of the UEV protein is to bind the cargo-linked mono-ubiquitin moiety [147]. HIV-1 and certain other viruses subvert this function of Tsg101 in order to bud from the plasma membrane [148– 150]. Mechanistically, Tsg101 is recruited to the virus budding sites by binding to a tetrapeptide “PTAP” motif in the late domain of viral proteins such as HIV1-GAG. Tsg101 is essential for virus budding from the plasma membrane [148], so it is possible that the endocytic budding machinery is hijacked to the plasma membrane via the Tsg101/GAG interaction [85]. The solution structure of the Tsg101 UEV domain has been solved alone and in complex with a PTAP-containing peptide [151, 152]. Human Tsg101 contains 390 amino acids, with the UEV domain located at the N-terminus [152]. The UEV domain is the minimal region needed to bind HIV-1 GAG, and is also the domain involved in mono-ubiquitin recognition and binding [153]. The overall fold of the UEV domain is similar to that of E2s. One notable difference is the presence of an extra N-terminal α-helix on Tsg101 [152]. The other major difference is the absence in Tsg101 UEV of the two C-terminal α-helices of the E2s – a truncation that was also seen in the Mms2 structures. This truncation appears to be a special trait of UEV proteins [154]. In Tsg101 UEV, the absence of the C-terminal helices helps to create the binding site for the PTAP peptide [151].

8 Conclusions

When aligning the structures of a canonical E2, Mms2, and the Tsg101 UEV domain, the hydrophobic core and the region surrounding the vestigial active site are quite similar, but the Tsg101 UEV domain differs from Mms2 and canonical E2s in the positions of the first two β-strands [152]. In Tsg101 they are elongated and shifted toward the N-terminus, forming a β-hairpin that extends 11 residues outside the main body of the domain [152]. This loop is important for ubiquitin binding by Tsg101 [152]. As determined by chemical shift mapping and mutagenesis studies, the Tsg101/ubiquitin binding interface involves the bottom half of the four-stranded β-sheet, including the β-hairpin (loop S1–S2). The binding interface for ubiquitin on Tsg101 is distinct from the surface that Mms2 uses to position the acceptor ubiquitin within the Ubc13/Mms2 complex [131, 144]. Thus, two different UEVs bind ubiquitin in two different ways. The structural biology and biochemistry of UEVs illustrates how modest changes to an E2-like module can create new, functionally important interaction sites. The UEV domain is just one of a growing set of small domains that can endow other protein domains with ubiquitin-binding capability (reviewed in Ref. [63]). Such binding elements are likely to play important roles in transducing ubiquitin signals in diverse cellular pathways.

8 Conclusions

We have emphasized the biochemical properties of E2s, particularly interactions with other factors in the conjugation cascade, because these properties are central to the biological actions of E2s. We have tried to give a flavor of the “creativity and economy” [103] with which E2s have evolved to maximize the interaction potential of a relatively small and conserved surface (Figure 7). Owing to the large scope of the relevant literature and the limited length of this chapter, we have not done full justice to the biological breadth of the E2 enzyme family. For example, we have focused on yeast and mammalian enzymes, but ubiquitin conjugation is increasingly being studied in other model organisms, including flies, worms, and plants. These systems offer powerful tools to address outstanding questions about ubiquitin-dependent pathways in general and E2 enzymes in particular. What are some of those questions? Significant uncertainties remain concerning E2 catalysis and mechanism, as discussed in Section 6. Another important question has been largely ignored in this review – exactly why are there so many E2s? One appealing model is that the identity of the E2 can modulate the substrate specificity of the E3, but experimental evidence for this model remains sparse. Another possibility is that the E2 has little or no influence on substrate choice, but rather helps to control the flux of activated ubiquitin to its cognate E3. In view of the remarkable developments in ubiquitin biology over the last decade, we should be prepared for both interesting and unexpected answers to these (and other) questions.

23

24 Ubiquitin-conjugating Enzymes

References 1 HERSHKO, A. and CIECHANOVER, A. The ubiquitin system. Annu. Rev. Biochem. 1998, 67, 425–479. 2 CIECHANOVER, A., HELLER, H., ELIAS, S., HAAS, A. L., and HERSHKO, A. ATP-dependent conjugation of reticulocyte proteins with the polypeptide required for protein degradation. Proc. Natl. Acad. Sci. USA 1980, 77, 1365–1368. 3 HERSHKO, A., CIECHANOVER, A., HELLER, H., HAAS, A. L., and ROSE, I. A. Proposed role of ATP in protein breakdown: conjugation of proteins with multiple chains of the polypeptide of ATP-dependent proteolysis. Proc. Natl. Acad. Sci. USA 1980, 77, 1783–1786. 4 HAAS, A. L., WARMS, J. V. B., HERSHKO, A., and ROSE, I. A. Ubiquitin-activating enzyme. Mechanism and role in protein-ubiquitin conjugation. J. Biol. Chem. 1982, 257, 2543–2548. 5 CANE, D. E. and WALSH, C. T. The parallel and convergent universes of polyketide synthases and nonribosomal peptide synthetases. Chem. Biol. 1999, 6, R319–R325. 6 HERSHKO, A., HELLER, H., ELIAS, S., and CIECHANOVER, A. Components of ubiquitin-protein ligase system. J. Biol. Chem. 1983, 258, 8206–8214. 7 PICKART, C. M. and ROSE, I. A. Functional heterogeneity of ubiquitin carrier proteins. J. Biol. Chem. 1985, 260, 1573–1781. 8 JENTSCH, S., MCGRATH, J. P., and VARSHAVSKY, A. The yeast DNA repair gene RAD6 encodes a ubiquitin-conjugating enzyme. Nature 1987, 329, 131–134. 9 GOEBL, M. G., YOCHEM, J., JENTSCH, S., MCGRATH, J. P., VARSHAVSKY, A., and BYERS, B. The yeast cell cycle gene CDC34 encodes a ubiquitin-conjugating enzyme. Science 1988, 241, 1331–1335. 10 REYNOLDS, P., WEBER, S., and PRAKASH, L. RAD6 gene of Saccharomyces cerevisiae encodes a protein containing a tract of 13 consecutive aspartates. Proc. Natl. Acad. Sci. USA 1985, 82, 168–172.

11 SUNG, P., PRAKASH, S., and PRAKASH, L. Stable ester conjugate between the Saccharomyces cerevisiae RAD6 protein and ubiquitin has no biological activity. J. Mol. Biol. 1991, 221, 745–749. 12 BANERJEE, A., DESHAIES, R. J., and CHAU, V. Characterization of a dominant negative mutant of the cell cycle ubiquitin-conjugating enzyme Cdc34. J. Biol. Chem. 1995, 270, 26209–26215. 13 SEMPLE, C. A. The comparative proteomics of ubiquitination in mouse. Genome Res 2003, 13, 1389–1394. 14 WONG, B. R., PARLATI, F., QUK, K., DEMO, S., PRAY, T., HUANG, J., PAYAN, D. G., and BENNETT, M. K. Drug discovery in the ubiquitin regulatory pathway. Drug Discov. Today 2003, 8, 746–754. 15 KOLMAN, C. J., TOTH, J., and GONDA, D. K. Identification of a portable determinant of cell cycle function within the carboxyl-terminal domain of the yeast CDC34 (UBC3) ubiquitin conjugating (E2) enzyme. EMBO J. 1992, 11, 3081–3090. 16 SILVER, E. T., GWOZD, T. J., PTAK, C., GOEBL, M. G., and ELLISON, M. J. A chimeric ubiquitin conjugating enzyme that combines the cell cycle properties of CDC34 (UBC3) and the DNA repair properties of RAD6 (UBC2): implications for the structure, function, and evolution of the E2s. EMBO J. 1992, 11, 3091–3098. 17 PTAK, C., PRENDERGAST, J. A., HODGINS, R., KAY, C. M., CHAU, V., and ELLISON, M. J. Functional and physical characterization of the cell cycle ubiquitin-conjugating enzyme CDC34 (UBC3). J. Biol. Chem. 1994, 269, 26539–26545. 18 HODGINS, R., GWOZD, C., ARNASON, T., CUMMINGS, M., and ELLISON, M. J. The tail of a ubiquitin-conjugating enzyme redirects multi-ubiquitin chain synthesis from the lysine 48-linked configuration to a novel nonlysine-linked form. J. Biol. Chem. 1996, 271, 28766–28771. 19 MATHIAS, N., STEUSSY, C. N., and GOEBL, M. G. An essential domain within Cdc34p is required for binding to a

References

20

21

22

23

24

25

26

27

28

29

complex containing Cdc4p and Cdc53p in Saccharomyces cerevisiae. J. Biol. Chem. 1998, 273, 4040–4045. COOK, W. J., JEFFREY, L. C., SULLIVAN, M. L., and VIERSTRA, R. D. Three-dimensional structure of a ubiquitin-conjugating enzyme (E2). J. Biol. Chem. 1992, 267, 15116–15121. COOK, W. J., JEFFREY, L. C., XU, Y., and CHAU, V. Tertiary structures of class I ubiquitin-conjugating enzymes are highly conserved: crystal structure of yeast Ubc4. Biochemistry 1993, 32, 13809–13817. HOCHSTRASSER, M. Ubiquitin-dependent protein degradation. Annu. Rev. Genet. 1996, 30, 405–439. PICKART, C. M. Mechanisms underlying ubiquitination. Annu. Rev. Biochem. 2001, 70, 503–533. SEUFERT, W. and JENTSCH, S. Ubiquitin-conjugating enzymes UBC4 and UBC5 mediate selective degradation of short-lived and abnormal proteins. EMBO J. 1990, 9, 543–550. JUNGMANN, J., REINS, H.-A., SCHOBERT, C., and JENTSCH, S. Resistance to cadmium mediated by ubiquitin-dependent proteolysis. Nature 1993, 361, 369–371. HEINEMEYER, W., KLEINSCHMIDT, J. A., SAIDOWSKY, J., ESCHER, C., and WOLF, D. H. Proteinase yscE, the yeast proteasome/multicatalyticmultifunctional proteinase: mutants unravel its function in stress induced proteolysis and uncover its necessity for cell survival. EMBO J. 1991, 10, 555–562. JOHNSON, E. S., MA, P. C., OTA, I. M., and VARSHAVSKY, A. A proteolytic pathway that recognizes ubiquitin as a degradation signal. J. Biol. Chem. 1995, 270, 17442–17456. SEUFERT, W., MCGRATH, J. P., and JENTSCH, S. UBC1 encodes a novel member of an essential subfamily of yeast ubiquitin-conjugating enzymes involved in protein degradation. EMBO J. 1990, 9, 4535–4541. GONEN, H., BERCOVICH, B., ORIAN, A., CARRANO, A., TAKIZAWA, C., YAMANAKA, K., PAGANO, M., IWAI, K., and

30

31

32

33

34

35

36

37

38

CIECHANOVER, A. Identification of the ubiquitin carrier proteins, E2s, involved in signal-induced conjugation and subsequent degradation of IκBα. J. Biol. Chem. 1999, 274, 14823–14830. DESHAIES, R. J. SCF and cullin/RING H2-based ubiquitin ligases. Annu. Rev. Cell Devel. Biol. 1999, 15, 435–467. SCHWOB, E., BOHM, T., MENDENHALL, M. D., and NASMYTH, K. The B-type cyclin kinase inhibitor p40SIC1 controls the G1 to S transition in S. cerevisiae Cell 1994, 79, 233–244. FELDMAN, R. M. R., CORRELL, C. C., KAPLAN, K. B., and DESHAIES, R. J. A complex of Cdc4p, Skp1p, and Cdc53p/cullin catalyzes ubiquitination of the phosphorylated CDK inhibitor Sic1p. Cell 1997, 91, 221–230. SKOWYRA, D., CRAIG, K. L., TYERS, M., ELLEDGE, S. J., and HARPER, J. W. F-box proteins are receptors that recruit phosphorylated substrates to the SCF ubiquitin-ligase complex. Cell 1997, 91, 209–219. PATTON, E. E., WILLEMS, A. R., SA, D., KURAS, L., THOMAS, D., CRAIG, K. L., and TYERS, M. Cdc53 is a scaffold protein for multiple Cdc34/Skp1/F-box protein complexes that regulate cell division and methionine biosynthesis in yeast. Genes Dev. 1998, 12, 692–705. JACKSON, P. K., ELDRIDGE, A. G., FREED, E., FURSTENTHAL, L., HSU, J. Y., KAISER, B. K., and REIMANN, J. D. R. The lore of the RINGs: substrate recognition and catalysis by ubiquitin ligases. Trends Cell Biol. 2000, 10, 429–439. KUS, B., CALDON, C. E., ANDORN-BROZA, R., and EDWARDS, A. M. Functional interactions of 13 yeast SCF complexes with a set of yeast E2 enzymes in vitro. Proteins: Structure, Function, and Bioinformatics 2004, 54, 455–467. BIEDERER, T., VOLKWEIN, C., and SOMMER, T. Role of Cue1p in ubiquitination and degradation at the ER surface. Science 1997, 278, 1806–1809. KOSTOVA, Z. and WOLF, D. H. For whom the bell tolls: protein quality control of the endoplasmic reticulum and the ubiquitin-proteasome connection. EMBO J. 2003, 22, 2309–2317.

25

26 Ubiquitin-conjugating Enzymes

39 BAYS, N. W., GARDNER, R. G., SEELIG, L. P., JOAZEIRO, C. A., and HAMPTON, R. Y. Hrd1p/Der3p is a membrane-anchored ubiquitin ligase required for ER-associated degradation. Nature Cell Biol. 2000, 3, 24–29. 40 FRIEDLANDER, R., JAROSCH, E., URBAN, J., VOLKWEIN, C., and SOMMER, T. A regulatory link between ER-associated protein degradation and the unfolded-protein response. Nature Cell Biol. 2000, 2, 379–384. 41 TRAVERS, K. J., PATIL, C. K., WODICKA, L., LOCKHART, D. J., WEISSMAN, J. S., and WALTER, P. Functional and genomic analyses reveal an essential coordination between the unfolded protein response and ER-associated degradation. Cell 2000, 101, 249–258. 42 SWANSON, R., LOCHER, M., and HOCHSTRASSER, M. A conserved ubiquitin ligase of the nuclear envelope/endoplasmic reticulum that functions in both ER-associated and Matα2 repressor degradation. Genes Dev. 2001, 15, 2660–2674. 43 TIWARI, S. and WEISSMAN, A. M. Endoplasmic reticulum (ER)-associated degradation of T cell receptor subunits. J. Biol. Chem. 2001, 276, 16193–16200. 44 WEBSTER, J. M., TIWARI, S., WEISSMAN, A. M., and WOJCIKIEWICZ, R. J. H. Inositol 1,4,5-trisphosphate receptor ubiquitination is mediated by mammalian Ubc7, a component of the endoplasmic reticulum-associated degradation pathway, and is inhibited by chelation of intracellular Zn2+ . J. Biol. Chem. 2003, 278, 38238–38246. 45 FANG, S., FERRONE, M., YANG, C., JENSEN, J. P., TIWARI, S., and WEISSMAN, A. M. The tumor autocrine motility factor receptor, gp78, is a ubiquitin protein ligase implicated in degradation from the endoplasmic reticulum. Proc. Natl. Acad. Sci. USA 2001, 98, 14422–14427. 46 SOMMER, T. and JENTSCH, S. A protein translocation defect linked to ubiquitin conjugation at the endoplasmic reticulum. Nature 1993, 365, 176–179. 47 BIEDERER, T., VOLKWEIN, C., and SOMMER, T. Degradation of subunits of the Sec61p complex, an integral component of the

48

49

50

51

52

53

54

55

ER membrane, by the ubiquitin-proteasome pathway. EMBO J. 1996, 15, 2069–2076. HILLER, M. M., FINGER, A., SCHWEIGER, M., and WOLF, D. H. ER degradation of a misfolded luminal protein by the cytosolic ubiuqitin-proteasome pathway. Science 1996, 273, 1725–1728. CHEN, P., JOHNSON, P., SOMMER, T., JENTSCH, S., and HOCHSTRASSER, M. Multiple ubiquitin-conjugating enzymes participate in the in vivo degradation of the yeast MATα2 repressor. Cell 1993, 74, 357–369. WALTER, J., URBAN, J., VOLKWEIN, C., and SOMMER, T. Sec61p-independent degradation of the tail-anchored ER membrane protein Ubc6p. EMBO J. 2001, 20, 3124–3131. RAO, H., UHLMANN, F., NASMYTH, K., and VARSHAVSKY, A. Degradation of a cohesin subunit by the N-end rule pathway is essential for chromosome stability. Nature 2001, 410, 955–959. BYRD, C., TURNER, G. C., and VARSHAVSKY, A. The N-end rule pathway controls the import of peptides through degradation of a transcriptional repressor. EMBO J. 1998, 17, 269–277. KAPLUN, L., IVANTSIV, Y., KORNITZER, D., and RAVEH, D. Functions of the DNA damage response pathway target Ho endonuclease of yeast for degradation via the ubiquitin-26S proteasome pathway. Proc. Natl. Acad. Sci. USA 2000, 97, 10077–10082. KOKEN, M. H. M., HOOGERBRUGGE, J. W., JASPERS-DEKKER, I., de WIT, J., WILLEMSEN, R., ROEST, H. P., GROOTEGOED, J. A., and HOEIJMAKERS, J. H. J. Expression of the ubiquitin-conjugating DNA repair enzymes HHR6A and B suggests a role in spermatogenesis and chromatin modification. Dev. Biol. 1996, 173, 119–132. ROEST, H. P., van KLAVEREN, J., de WIT, J., van GURP, C. G., KOKEN, M. H. M., VERMEY, M., van ROIJEN, J. H., HOOGERBRUGGE, J. W., VREEBERG, M. T. M., BAARENDS, W. M., BOOTSMA, D., GROOTEGOED, J. A., and HOEIJMAKERS, J. H. J. Inactivation of the HR6B

References

56

57

58

59

60

61

62

63

64

ubiquitin-conjugating DNA repair enzyme in mice causes male sterility associated with chromatin modification. Cell 1996, 86, 799–810. KWON, Y. T., KASHINA, A. S., DAVYDOV, I. V., HU, R.-G., AN, J. Y., SEO, J. W., DU, F., and VARSHAVSKY, A. An essential role of N-terminal arginylation in cardovascular development. Science 2002, 297, 96–99. SCHULE, T., ROSE, M., ENTIAN, K.-D., THUMM, M., and WOLF, D. H. Ubc8p functions in catabolite degradation of fructose-1,6-bisphosphatase in yeast. EMBO J. 2000, 19, 2161–2167. LI, Y.-P., LECKER, S. H., CHEN, Y., WADDELL, I. D., GOLDBERG, A. L., and REID, M. B. TNF-α increases ubiquitin-conjugating activity in skeletal muscle by up-regulating UbcH2/E220K . FASEB J. 2003, 17, 1048–1057. WEFES, I., MASTRANDREA, L. D., HALDEMAN, M., KOURY, S. T., TAMBURLIN, J., PICKART, C. M., and FINLEY, D. Inducation of ubiquitin-conjugating enzymes during terminal erythroid differentiation. Proc. Natl. Acad. Sci. USA 1995, 92, 4982–4986. GITAN, R. S., SHABABI, M., KRAMER, M., and EIDE, D. J. A cytosolic domain of the yeast Zrt1 zinc transporter is required for its post-translational inactivation in response to zinc and cadmium. J. Biol. Chem. 2003, 278, 39558–39564. HICKE, L. and RIEZMAN, H. Ubiquitination of a yeast plasma membrane receptor signals its ligand-stimulated endocytosis. Cell 1996, 84, 277–287. DUNN, R. and HICKE, L. Domains of the Rsp5 ubiquitin-protein ligase required for receptor-mediated and fluid-phase endocytosis. Mol. Biol. Cell 2001, 12, 421–435. HICKE, L. and DUNN, R. Regulation of membrane protein transport by ubiquitin and ubiquitin-binding proteins. Annu. Rev. Cell Devel. Biol. 2003, 19, 141–172. ULRICH, H. D. Degradation or maintenance: actions of the ubiquitin system on eukaryotic chromatin. Eukaryotic Cell 2002, 1, 1–10.

65 HOEGE, C., PFANDER, B., MOLDOVAN, G.-L., PYROWOLAKIS, G., and JENTSCH, S. RAD6-dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 2002, 419, 135–141. 66 STELTER, P. and ULRICH, H. D. Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature 2003, 425, 188–191. 67 HWANG, W. W., VENKATASUBRAHMANYAM, S., IANCULESCU, A. G., TONG, A., BOONE, C., and MADHANI, H. D. A conserved RING finger protein required for histone H2B monoubiquitination and cell size control. Mol. Cell 2003, 11, 261–266. 68 ROBZYK, K., RECHT, J., and OSLEY, M. A. Rad6-dependent ubiquitination of histone H2B in yeast. Science 2000, 287, 501–504. 69 WOOD, A., KROGAN, N. J., DOVER, J., SCHNEIDER, J., HEIDT, J., BOATENG, M. A., DEAN, K., GOLSHANI, A., ZHANG, Y., GREENBLATT, J. F., JOHNSTON, M., and SHILATIFARD, A. Bre1, an E3 ubiquitin ligase required for recruitment and substrate selection of Rad6 at a promoter. Mol. Cell 2003, 11, 267–274. 70 SCHWARTZ, D. C. and HOCHSTRASSER, M. A superfamily of protein tags: ubiquitin, SUMO and related modifiers. Trends Biochem. Sci. 2003, 28, 321–328. 71 SUN, Z.-W. and ALLIS, C. D. Ubiquitination of histone H2B regulates H3 methylation and gene silencing in yeast. Nature 2002, 418, 104–108. 72 HOFMANN, R. M. and PICKART, C. M. Noncanonical MMS2-encoded ubiquitin-conjugating enzyme functions in assembly of novel polyubiquitin chains for DNA repair. Cell 1999, 96, 645–653. 73 ULRICH, H. D. and JENTSCH, S. Two RING finger proteins mediate cooperation between ubiquitin-conjugating enzymes in DNA repair. EMBO J. 2000, 19, 3388–3397. 74 SUN, L. and CHEN, Z. J. The novel functions of ubiquitination in signaling. Curr. Opin. Cell Biol. 2004, 16, 119–126. 75 WIEBEL, F. F. and KUNAU, W. H. The Pas2 protein essential for peroxisome biogenesis is related to

27

28 Ubiquitin-conjugating Enzymes

76

77

78

79

80

81

82

83

84

ubiquitin-conjugating enzymes. Nature 1992, 359, 73–76. van der KLEI, I. J., HILBRANDS, R. E., KIEL, J. A. K. W., RASMUSSEN, S. W., CREGG, J. M., and VEENHUIS, M. The ubiquitin-conjugating enzyme Pex4p of Hansenula polymorpha is required for efficient functioning of the PTS1 import machinery. EMBO J. 1998, 17, 3608–3618. KOLLER, A., SNYDER, W. B., FABER, K. N., WENZEL, T. J., RANGELL, L., KELLER, G. A., and SUBRAMANI, S. Pex22p of Pichia pastoris, essential for peroxisomal matrix protein import, anchors the ubiquitin-conjugating enzyme, Pex4p, on the peroxisomal membrane. J. Cell Biol. 1999, 146, 99–112. ECKERT, J. H. and JOHNSSON, N. Pex10p links the ubiquitin conjugating enzyme Pex4p to the protein import machinery of the peroxisome. J. Cell Sci. 2003, 116, 3623–3634. ARISTARKHOV, A., EYTAN, E., MOGHE, A., ADMON, A., HERSHKO, A., and RUDERMAN, J. V. E2-C a cyclin-selective ubiquitin carrier protein required for the destruction of mitotic cyclins. Proc. Natl. Acad. Sci. USA 1996, 93, 4294–4299. PETERS, J. M. The anaphase-promoting complex: proteolysis in mitosis and beyond. Mol. Cell 2002, 9, 931–943. YU, H., KING, R. W., PETERS, J.-M., and KIRSCHNER, M. W. Identification of a novel ubiquitin-conjugating enzyme involved in mitotic cyclin degradation. Curr. Biol. 1996, 6, 455–466. SEINO, H., KISHI, T., NISHITANI, H., and YAMAO, F. Two ubiquitin-conjugating enzymes, UbcP1/Ubc4 and UbcP4/Ubc11, have distinct functions for ubiquitination of mitotic cyclin. Mol. Cell. Biol. 2003, 23, 3497–3505. TOWNSLEY, F. M., ARISTARKHOV, A., BECK, S., HERSHKO, A., and RUDERMAN, J. V. Dominant-negative cyclin-selective ubiquitin carrier protein E2-C/UbcH10 blocks cells in metaphase. Proc. Natl. Acad. Sci. USA 1997, 94, 2362–2367. TOWNSLEY, F. M. and RUDERMAN, J. V. Functional analysis of the Saccharomyces cerevisiae UBC11 gene. Yeast 1998, 14, 747–757.

85 PORNILLOS, O., GARRUS, J. E., and SUNDQUIST, W. I. Mechanisms of enveloped RNA virus budding. Trends Cell Biol. 2002, 12, 569–579. 86 FURMAN, M. H. and PLOEGH, H. L. Lessons from viral manipulation of protein disposal pathways. J. Clin. Invest. 2002, 110, 875–879. 87 HINGAMP, P. M., ARNOLD, J. E., MAYER, R. J., and DIXON, L. K. A ubiquitin conjugating enzyme encoded by African swine fever virus. EMBO. J. 1992, 11, 361–366. 88 RODRIGUEZ, J. M., SALAS, M. L., and VINUELA, E. Genes homologous to ubiquitin-conjugating proteins and eukaryotic transcription factor SII in African swine fever virus. Virology 1992, 186, 40–52. 89 HAGGLUND, R. and ROIZMAN, B. Characterization of the novel E3 ubiquitin ligase encoded in exon 3 of herpes simplex virus-1-infected cell protein 0. Proc. Natl. Acad. Sci. USA 2002, 99, 7889–7894. 90 SONG, S., KIM, S.-Y., HONG, Y.-M., JO, D.-G., LEE, J.-Y., SHIM, S. M., CHUNG, C.-W., SEO, S. J., YOO, Y. J., KOH, J.-Y., LEE, M. C., YATES, A. J., ICHIJO, H., and JUNG, Y.-K. Essential role of E2–25K/Hip-2 in mediating amyloid-β toxicity. Mol. Cell 2003, 12, 553–563. 91 LAM, Y. A., PICKART, C. M., ALBAN, A., LANDON, M., JAMIESON, C., RAMAGE, R., MAYER, R. J., and LAYFIELD, R. Inhibition of the ubiquitin-proteasome system in Alzheimer’s disease. Proc. Natl. Acad. Sci. USA 2000, 97, 9902–9906. 92 OKAMOTO, Y., OZAKI, T., MIYAZAKI, K., AOYAMA, M., MIYAZAKI, M., and NAKAGAWARA, A. UbcH10 is the cancer-related E2 ubiquitin-conjugating enzyme. Cancer Res. 2003, 63, 4167–4173. 93 SHEKHAR, M. P. V., LYAKHOVICH, A., VISSCHER, D. W., HENG, H., and KONDRAT, N. Rad6 over-expression induces multinucleation, centrosom amplification, abnormal mitosis, aneuploidy, and transformation. Cancer Res. 2002, 62, 2115–2124. 94 MULLER, S., HOEGE, C., PYROWOLAKIS, G., and JENTSCH, S. SUMO, ubiquitin’s

References

95

96

97

98

99

100

101

102

103

mysterious cousin. Nature Rev. Mol. Cell Biol. 2001, 2, 202–210. HOCHSTRASSER, M. Evolution and function of ubiquitin-like protein-conjugation systems. Nature Cell Biol. 2000, 2, E153–E157. WALDEN, H., PODGORSKI, M. S., and SCHULMAN, B. A. Insights into the ubiquitin transfer cascade from the structure of the E1 for NEDD8. Nature 2003, 422, 330–334. BENCSATH, K. P., PODGORSKI, M. S., PAGALA, V. R., SLAUGHTER, C. A., and SCHULMAN, B. A. Identification of a multifunctional binding site on Ubc9p required for Smt3p conjugation. J. Biol. Chem. 2002, 277, 47938–47945. SULLIVAN, M. L. and VIERSTRA, R. D. Cloning of a 16-kDa ubiquitin carrier protein from wheat and Arabidopsis thaliana. Identification of functional domains by in vitro mutagenesis. J. Biol. Chem. 1991, 266, 23878–23885. HAAS, A. L., BRIGHT, P. M., and JACKSON, V. E. Functional diversity among putative E2 isozymes in the mechanism of ubiquitin-histone ligation. J. Biol. Chem. 1988, 163, 13268–13275. SIEPMANN, T. J., BOHNSACK, R. N., TOKGOZ, Z., BABOSHINA, O., and HAAS, A. L. Protein interactions within the N-end rule ubiquitin ligation pathway. J. Biol. Chem. 2003, 278, 9448–9457. MIURA, T., KLAUS, W., GSELL, B., MIYAMOTO, C., and SENN, H. Characterization of the binding interface between ubiquitin and class I human ubiquitin-conjugating enzyme 2b by multidimensional heteronuclear NMR spectroscopy in solution. J. Mol. Biol. 1999, 290, 213–228. HAMILTON, K. S., ELLISON, M. J., BARBER, K. R., WILLIAMS, R. S., HUZIL, J. T., MCKENNA, S., PTAK, C., GLOVER, M., and SHAW, G. S. Structure of a conjugating enzyme-ubiquitin thiolester intermediate reveals a novel role for the ubiquitin tail. Structure 2001, 9, 897–904. MCKENNA, S., MORAES, T., PASTUSHOK, L., PTAK, C., XIAO, W., SPYRACOPOULOS, L., and ELLISON, M. J. An NMR-based model of the ubiquitin-bound human

104

105

106

107

108

109

110

111

112

ubiquitin conjugation complex Mms2.Ubc13. The structural basis for lysine 63 chain catalysis. J. Biol. Chem. 2003, 278, 13151–13158. BORDEN, K. L. RING domains: master builders of molecular scaffolds? J. Mol. Biol. 2000, 295, 1103–1112. BORDEN, K. L. RING fingers and B-boxes: zinc-binding protein-protein interaction domains. Biochem. Cell Biol. 1998, 76, 351–358. ZHENG, N., WANG, P., JEFFREY, P. D., and PAVLETICH, N. P. Structure of a c-Cbl-UbcH7 complex: RING domain function in ubiquitin-protein ligases. Cell 2000, 102, 533–539. BRZOVIC, P. S., KEEFFE, J. R., NISHIKAWA, H., MAYAMOTO, K., FOX, D., FUKUDA, M., OHTA, T., and KLEVIT, R. Binding and recognition in the assembly of an active BRCA1-BARD1 ubiquitin ligase complex. Proc. Natl. Acad. Sci. USA 2003, 100, 5646–5651. HUANG, L., KINNUCAN, E., WANG, G., BEAUDENON, S., HOWLEY, P. M., HUIBREGTSE, J. M., and PAVLETICH, N. P. Structure of an E6AP-UbcH7 complex: insights into ubiquitination by the E2-E3 enzyme cascade. Science 1999, 286, 1321–1326. WU, G., XU, G., SCHULMAN, B. A., JEFFREY, P. D., HARPER, J. W., and PAVLETICH, N. P. Structure of a β-TrCP1-Skp1-β-catenin complex: destruction motif binding and lysine specificity of the SCFβ-TrCP1 ubiquitin ligase. Mol. Cell 2003, 11, 1145–1456. BRZOVIC, P. S., RAJAGOPAL, P., HOYT, D. W., KING, M.-C., and KLEVIT, R. E. Structure of a BRCA1-BARD1 heterodimeric RING-RING complex. Nature Struct. Biol. 2001, 8, 833–837. CONAWAY, R. C. and CONAWAY, J. W. The von Hippel-Lindau tumor suppressor complex and regulation of hypoxia-inducible transcription. Adv. Cancer Res. 2002, 85, 1–12. ZHENG, N., SCHULMAN, B. A., SONG, L., MILLER, J. J., JEFFREY, P. D., WANG, P., CHU, C., KOEPP, D. M., ELLEDGE, S. J., PAGANO, M., CONAWAY, R. C., CONAWAY, J. W., HARPER, J. W., and PAVLETICH, N. P. Structure of the Cul1-Rbx1-Skp1-F

29

30 Ubiquitin-conjugating Enzymes

113

114

115

116

117

118

119

120

121

boxSkp2 SCF ubiquitin ligase complex. Nature 2002, 416, 703–709. KAWAKAMI, T., CHIBA, T., SUZUKI, T., IWAI, K., YAMANAKA, K., MINATO, N., SUZUKI, H., SHIMBARA, N., HIDAKA, Y., OSAKA, F., OMATA, M., and TANAKA, K. NEDD8 recruits E2-ubiquitin to SCF E3 ligase. EMBO J. 2001, 20, 4003–4012. WU, K., CHEN, A., TAN, P., and PAN, Z.-Q. The Nedd8-conjugated ROC1-CUL1 core ubiquitin ligase utilizes Nedd8 charged surface residues for efficient polyubiquitin chain assembly catalyzed by Cdc34. J. Biol. Chem. 2002, 277, 516–527. ORLICKY, S., TANG, X., WILLEMS, A., TYERS, M., and SICHERI, F. Structural basis for phosphodependent substrate selection and orientation by the SDFCdc4 ubiquitin ligase. Cell 2003, 112, 243–256. DEFFENBAUGH, A. E., SCAGLIONE, K. M., BURANDA, T., SKLAR, L. A., and SKOWYRA, D. Release of ubiquitin-charged Cdc34-S∼Ub from the RING domain is essential for ubiquitination of the SCFCdc4 -bound substrate Sic1. Cell 2003, 114, 611–622. PETROSKI, M. D. and DESHAIES, R. J. Context of multiubiquitin chain attachment influences the rate of Sic1 degradation. Mol. Cell 2003, 11, 1435–1444. PENG, J., SCHWARTZ, D., ELIAS, J. E., THOREEN, C. C., CHENG, D., MARSISCHKY, G., ROELOFS, J., FINLEY, D., and GYGI, S. P. A proteomics approach to understanding protein ubiquitination. Nature Biotechnol. 2003, 21, 921–926. PRINGA, E., MARTINEZ-NOEL, G., MULLER, U., and HARBERS, K. Interaction of the ring finger-related U-box motif of a nuclear dot protein with ubiquitin-conjugating enzymes. J. Biol. Chem. 2001, 276, 19617–19623. KOEGL, M., HOPPE, T., SCHLENKER, S., ULRICH, H. D., MAYER, T. U., and JENTSCH, S. A novel ubiquitination factor, E4, is involved in multiubiquitin chain assembly. Cell 1999, 96, 635–644. JIANG, J., BALLINGER, C. A., WU, Y., DAI, Q., CYR, D. M., HOHFELD, J., and PATTERSON, C. CHIP is a

122

123

124

125

126

127

128

129

130

U-box-dependent E3 ubiquitin ligase: identification of Hsc70 as a target for ubiquitylation. J. Biol. Chem. 2001, 276, 42938–42944. MURATA, S., MINAMI, Y., MINAMI, M., CHIBA, T., and TANAKA, K. CHIP is a chaperone-dependent E3 ligase that ubiquitylates unfolded protein. EMBO Rep. 2001, 2, 1133–1138. OHI, M. D., VANDER KOOI, C. W., ROSENBERG, J. A., CHAZIN, W. J., and GOULD, K. L. Structural insights into the U-box, a domain associated with multi-ubiquitination. Nature Struct. Biol. 2003, 10, 250–255. ARAVIND, L. and KOONIN, E. V. The U box is a modified RING finger – a common domain in ubiquitination. Curr. Biol. 2000, 10, R132–R224. HATAKEYAMA, S., YADA, M., MATSUMOTO, M., ISHIDA, N., and NAKAYAMA, K. I. U box proteins as a new family of ubiquitin-protein ligases. J. Biol. Chem. 2001, 276, 33111–31120. HUIBREGTSE, J. M., SCHEFFNER, M., BEAUDENON, S., and HOWLEY, P. M. A family of proteins structurally and functionally related to the E6-AP ubiquitin-protein ligase. Proc. Natl. Acad. Sci. USA 1995, 92, 2563–2567. HUIBREGTSE, J. M., SCHEFFNER, M., and HOWLEY, P. M. A cellular protein mediates association of p53 with the E6 oncoprotein of human papillomavirus types 16 or 18. EMBO J. 1991, 10, 4129–4135. SCHEFFNER, M., NUBER, U., and HUIBREGTSE, J. M. Protein ubiquitination involving an E1-E2-E3 enzyme ubiquitin thioester cascade. Nature 1995, 373, 81–83. VERDECIA, M. A., JOAZEIRO, C. A. P., WELLS, N. J., FERRER, J.-L., BOWMAN, M. E., HUNTER, T., and NOEL, J. P. Conformational flexibility underlies ubiquitin ligation mediated by the WWP1 HECT domain E3 ligase. Mol. Cell 2003, 11, 249–259. BERNIER-VILLAMOR, V., SAMPSON, D. A., MATUNIS, M. J., and LIMA, C. D. Structural basis for E2-mediated SUMO conjugation revealed by a complex between ubiquitin-conjugating enzyme

References

131

132

133

134

135

136

137

138

139

140

Ubc9 and RanGAP1. Cell 2002, 108, 345–356. VANDEMARK, A. P., HOFMANN, R. M., TSUI, C., PICKART, C. M., and WOLBERGER, C. Molecular insights into polyubiquitin chain assembly: crystal structure of the Mms2/Ubc13 heterodimer. Cell 2001, 105, 711–720. DENG, L., WANG, C., SPENCER, E., YANG, L., BRAUN, A., YOU, J., SLAUGHTER, C., PICKART, C., and CHEN, Z. J. Activation of the IκB kinase complex by TRAF6 requires a dimeric ubiquitin-conjugating enzyme complex and a unique polyubiquitin chain. Cell 2000, 103, 351–361. CARTER, P. and WELLS, J. A. Dissecting the catalytic triad of a serine protease. Nature 1988, 332, 564–568. RAWLINGS, N. D. and BARRETT, A. J. Families of cysteine peptidases. Meth. Enzymol. 1994, 244, 461–486. TONG, H., HATEBOER, G., PERRAKIS, A., BERNARDS, R., and SIXMA, T. K. Crystal structure of murine/human Ubc9 provides insight into the variability of the ubiquitin-conjugating system. J. Biol. Chem. 1997, 272, 21381–21387. WU, P.-Y., HANLON, M., EDDINS, M., TSUI, C., ROGERS, R., JENSEN, J. P., MATUNIS, M. J., WEISSMAN, A. M., WOLBERGER, C., and PICKART, C. M. A conserved catalytic residue in the E2 enzyme family. EMBO J. 2003, 22, 5241–5250. HU, M., LI, P., LI, M., LI, W., YAO, T., WU, J.-W., GU, W., COHEN, R. E., and SHI, Y. Crystal structure of a UBP-family deubiquitinating enzyme in isolation and in complex with ubiquitin aldehyde. Cell 2002, 111, 1041–1054. MOSSESSOVA, E. and LIMA, C. D. Ulp1-SUMO crystal structure and genetic analysis reveal conserved interactions and a regulatory element essential for cell growth in yeast. Mol. Cell 2000, 5, 865–876. JOHNSTON, S. C., RIDDLE, S. M., COHEN, R. E., and HILL, C. P. Structural basis for the specificity of ubiquitin C-terminal hydrolases. EMBO J. 1999, 18, 3877–3887. BERLETH, E. S. and PICKART, C. M. Mechanism of ubiquitin conjugating

141

142

143

144

145

146

147

148

enzyme E2–230K: catalysis involving a thiol relay? Biochemistry 1995, 35, 1664–1671. HAUSER, H.-P., BARDROFF, M., PYROWOLAKIS, G., and JENTSCH, S. A giant ubiquitin-conjugating enzyme related to IAP apoptosis inhibitors. J. Cell Biol. 1998, 141, 1415–1422. SANCHO, E., VILA, M. R., SANCHEZ-PULIDO, L., LOZANO, J. J., PACIUCCI, R., NADAL, M., FOX, M., HARVEY, C., BERCOVICH, B., LOUKILI, N., CIECHANOVER, A., LIN, S. L., SANZ, F., ESTIVILL, X., VALENCIA, A., and THOMSON, T. M. Role of UEV-1, an inactive variant of the E2 ubiquitin-conjugating enzymes, in in vitro differentiation and cell cycle behavior of HT-29-M6 intestinal mucosecretory cells. Mol. Cell Biol. 1998, 18, 576–589. WANG, C., DENG, L., HONG, M., AKKARAJU, G. R., INOUE, J.-I., and CHEN, Z. J. TAK1 is a ubiquitin-dependent kinase of MKK and IKK. Nature 2001, 412, 346–351. MORAES, T. F., EDWARDS, R. A., MCKENNA, S., PASTUSHOK, L., XIAO, W., GLOVER, J. N., and ELLISON, M. J. Crystal structure of the human ubiquitin conjugating enzyme complex, hMms2-hUbc13. Nature Struct. Biol. 2001, 8, 669–673. ULRICH, H. D. Protein-protein interactions in an E2-RING finger complex: implications for ubiquitin-dependent DNA damage repair. J. Biol. Chem. 2003, 278, 7051–7058. KATZMANN, D. J., ODORIZZI, G., and EMR, S. D. Receptor downregulation and multivesticular-body sorting. Nature Rev. Mol. Cell Biol. 2002, 3, 893–905. KATZMANN, D. J., BABST, M., and EMR, S. D. Ubiquitin-dependent sorting into the multivesicular body pathway requires the function of a conserved endosomal protein sorting complex, ESCRT-I. Cell 2001, 106, 145–155. GARRUS, J. E., VON SCHWEDLER, U. K., PORNILLOS, O. W., MORHAM, S. G., ZAVITZ, K. H., WANG, H. E., WETTSTEIN, D. A., STRAY, K. M., COTE, M., RICH, R. L., MYSZKA, D. G., and SUNDQUIST, W. I.

31

32 Ubiquitin-conjugating Enzymes

149

150

151

152

153

154

155

156

157

Tsg101 and the vaculolar protein sorting pathway are essential for HIV-1 budding. Cell 2001, 107, 55–65. MARTIN-SERRANO, J., ZANG, T., and BIENIASZ, P. D. HIV-1 and Ebola virus encode small peptide motifs that recruit Tsg101 to sites of particle assembly to facilitate egress. Nature Med. 2001, 7, 1313–1319. CARTER, C. A. Tsg101: HIV-1’s ticket to ride. Trends Microbiol. 2002, 10, 203–205. PORNILLOS, W., ALAM, S. L., DAVIS, D. R., and SUNDQUIST, W. I. Structure of the Tsg101 UEV domain in complex with the PTAP motif of the HIV-1 p6 protein. Nature Struct. Biol. 2002, 9, 812–817. PORNILLOS, O., ALAM, S. L., RICH, R. L., MYSZKA, D. G., DAVIS, D. R., and SUNDQUIST, W. I. Structure and functional interactions of the Tsg101 UEV domain. EMBO J. 2002, 21, 2397–2406. GOFF, A., EHRLICH, L. S., COHEN, S. N., and CARTER, C. A. Tsg101 control of human immunodeficiency virus type 1 Gag trafficking and release. J. Virol. 2003, 77, 9173–9182. KOONIN, E. V. and ABAGYAN, R. A. TSG101 may be the prototype of a class of dominant negative ubiquitin regulators. Nature Genet. 1997, 16, 330–331. HOFMANN, K. and BUCHER, P. The UBA domain: a sequence motif present in multiple enzyme classes of the ubiquitination pathway. Trends Biochem. Sci. 1996, 21, 172–173. DOHMEN, R. J., MADURA, K., BARTEL, B., and VARSHAVSKY, A. The N-end rule is mediated by the UBC2(RAD6) ubiquitin-conjugating enzyme. Proc. Natl. Acad. Sci. USA 1991, 88, 7351–7355. BAILLY, V., LAMB, J., SUNG, P., PRAKASH, S., and PRAKASH, L. Specific complex

158

159

160

161

162

163

164

165

formation between yeast RAD6 and RAD18 proteins: a potential mechanism for targeting RAD6 ubiquitin-conjugating activity to DNA damage sites. Genes Dev. 1994, 8, 811–820. VARSHAVSKY, A. The N-end rule pathway of protein degradation. Genes Cells 1997, 2, 13–28. BRAUN, S., MATUSCHEWSKI, K., RAPE, M., THOMS, S., and JENTSCH, S. Role of the ubiquitin-selective CDC48UFD1/NPL4 chaperone (segregase) in ERAD of OLE1 and other substrates. EMBO J. 2002, 21, 615–621. QIN, S., NAKAJIMA, B., NOMURA, M., and ARFIN, S. M. Cloning and characterization of a Saccharomyces cerevisiae gene encoding a new member of the ubiquitin-conjugating protein family. J. Biol. Chem. 1991, 266, 15549–15554. SEUFERT, W., FUTCHER, B., and JENTSCH, S. Role of a ubiquitin-conjugating enzyme in degradation of S- and M-phase cyclins. Nature 1995, 373, 78–81. JOHNSON, E. S. and GUPTA, A. A. An E3-like factor that promotes SUMO conjugation to the yeast septins. Cell 2001, 106, 735–744. JOHNSON, E. S. and BLOBEL, G. Ubc9p is the conjugating enzyme for the ubiquitin-like protein Smt3p. J. Biol. Chem. 1997, 272, 26799–26802. LIAKOPOULOS, D., DOENGES, G., MATUSCHEWSKI, K., and JENTSCH, S. A novel protein modification pathway related to the ubiquitin system. EMBO J. 1998, 17, 2208–2214. LAMMER, D., MATHIAS, N., LAPLAZA, J. M., JIANG, W., LIU, Y., CALLIS, J., GOEBL, M., and ESTELLE, M. Modification of yeast Cdc53p by the ubiquitin-related protein Rub1p affects function of the SCFCdc4 complex. Genes Dev. 1998, 12, 914–926.

1

The Deubiquitinating Enzymes

Nathaniel S. Russell and Keith D. Wilkinson Emory University, Atlanta, USA

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8

1 Introduction

In the mid-1970s ubiquitin was found to be a covalent modiﬁer of proteins [1]. At the time, it was quite surprising to ﬁnd a protein that covalently modiﬁed another protein. Since then, the reversible covalent modiﬁcation of proteins by other proteins is known to be commonplace and ubiquitin is used to covalently modify hundreds of proteins, often for the purpose of targeting them to the proteasome for degradation. Protein degradation through the ubiquitin–proteasome system is facilitated by covalently linking ubiquitin to the ε-amino group of a lysine of a substrate protein through its C-terminal glycine [2]. A polyubiquitin (polyUb) chain is formed by linking subsequent ubiquitins to the lysine 48 residue of the preceding ubiquitin in the chain. A chain of four ubiquitins is sufﬁcient for the targeted protein to be recognized and degraded by the proteasome [3]. Conjugation of ubiquitin to other proteins is catalyzed by a three-enzyme cascade [4]. Conjugation begins by activation of ubiquitin by an E1, or Ub-activating enzyme, forming a high-energy thiol ester bond in an ATP-dependent reaction. The ubiquitin is transferred to an E2, or Ub-conjugating enzyme, which then usually pairs with an E3, or Ub-ligase enzyme, to conjugate the ubiquitin to a speciﬁc target protein. The usefulness of ubiquitin conjugation is not limited to the ubiquitin– proteasome pathway. Mono- and polyubiquitin are used as signals in various pathways including endocytosis, DNA repair, apoptosis, and transcriptional regulation [5–8]. Polyubiquitin chains can be formed using lysine residues other than K48, the linkage required for proteasomal degradation [9–11]. In addition, there are a number of other ubiquitin-like proteins that also behave as signaling molecules although they are not involved directly in proteasomal degradation. This group Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 The Deubiquitinating Enzymes

includes SUMO (small ubiquitin-related modiﬁer), Nedd8 (neural precursor cell expressed, developmentally down-regulated 8), ISG15 (interferon-stimulated gene 15), and others [12–14]. These proteins are conjugated to substrates in a similar manner to ubiquitin, using an E1, E2, and E3 cascade of enzymes speciﬁc for the particular ubiquitin-like protein involved [15–17]. Soon after it was shown that ubiquitin is conjugated to proteins, it was determined that this was a reversible process and deubiquitinating enzymes, or DUBs, could remove ubiquitin from ubiquitinated proteins [18, 19]. As the genes for ubiquitin and ubiquitin-like proteins were identiﬁed it became clear that all ubiquitin family members were synthesized as proproteins and processed to reveal the C-terminal glycylglycine of the active proteins [20]. Based on this information, DUBs were deﬁned as proteases that cleave at the C-terminus of ubiquitin or ubiquitin-like proteins to reverse conjugation to target proteins and also process the proproteins. Over 100 DUBs have been identiﬁed (see Table 1 for a list of the DUBs whose roles are known or suspected) and they are used to regulate ubiquitin and ubiquitin-like protein metabolism. Since cells utilize a combination of mono-ubiquitin, polyubiquitin, and ubiquitin-like proteins for a multitude of reasons and conjugate them to thousands of proteins, a regulatory system has evolved that is exceedingly complex and must be exquisitely regulated (see Figure 1). DUBs help regulate this system by processing proubiquitin into a mature form, recycling free polyubiquitin chains into monomeric Ub, assisting the degradation of proteasomal substrates, and regulating the ubiquitination levels of proteins in cellular pathways other than proteolysis. Thus, DUBs play crucial roles in determining the cellular fates of many proteins and regulating cellular function.

Fig. 1 DUB families and substrate speciﬁcity. DUBs can be classiﬁed by genetic relationships (family) or by substrate speciﬁcities (activity). Arrows point towards the substrates that members of each family can

process in a physiologically relevant way. Each family is capable of processing multiple substrates and each activity can be catalyzed by members of more than one family.

1 Introduction 3 Table 1 Physiological roles of DUBs revealed by deletion or knockdown experiments

DUB

Organism

Deletion/knockdown phenotype

Ubiquitin C-terminal hydrolases (UCHs) UCH-L1 mouse gracile axonal dystrophy UCH-L3

mouse

no detectable phenotype

UCH37

human

unknown

BAP1 YUH1

human S. cerevisiae

unknown cannot process proRUB1

Functional role

predominant neuronal UCH [112] undetermined neuronal function [113] edits polyubiquitin chains at proteasome [91] tumor suppressor? [22] processes proRUB1, Ub-adducts [83]

Ubiquitin speciﬁc processing proteases (UBPs) UBP1 S. cerevisiae no phenotype detected undetermined [97] UBP2 S. cerevisiae no phenotype detected undetermined [96] UBP3/ S. cerevisiae, polyubiquitin accumulation transcriptional silencing inhibitor, USP10 human regulates membrane transport [66, 118] UBP6 S. cerevisiae low levels of free ubiquitin processes polyubiquitin chains at proteasome [94] UBP8 S. cerevisiae increase in ubiquitinated transcriptional regulator [88] histone H2B UBP14/IsoT S. cerevisiae, increased polyubiquitin recycles free polyubiquitin to human levels, proteasome defects mono-ubiquitin [33, 86] UBP16 S. cerevisiae no phenotype detected undetermined function at mitochondria [70] DOA4 S. cerevisiae Ub-depletion, defective recycles Ub and polyubiquitin Ub recycling adducts [71, 84] Unp/USP4 mouse, human unknown undetermined USP7 human indirect p53 activation regulates p53 ubiquitination [36] UBPy/USP8 human increase in protein cell-growth regulation [119] ubiquitination USP14 mouse (UBP6 ataxia regulating synaptic activity plus homolog) proteasome [109] USP21 human unknown process Ub and Nedd8 conjugates, growth regulator? [32] USP25 human unknown over-expression has possible role in Down’s Syndrome [120] UBP41 human unknown promotes apoptosis [6] UBP43 mouse accumulation of Isg15 processes Isg15, regulates Isg15 conjugates conjugate levels [31] CYLD human cylindromatosis regulates K63 polyubiquitination of substrates in NF-κB pathway [7, 103, 104] fat facets Drosophila defective germ cell regulates speciﬁc developmental speciﬁcation, eye formation processes [74, 75] DUB1, 2, 2A mouse unknown cytokine speciﬁc growth regulators VDU1 human unknown regulation of Ub-proteasome pathway? [64]

4 The Deubiquitinating Enzymes Table 1 (Continued)

DUB

Organism

Deletion/knockdown phenotype

Functional role

JAMM Isopeptidases Rpn11 Yeast, human Csn5 Yeast, Drosophila

processes polyubiquitin chains at proteasome [46, 47] defects in SCF E3s in yeast, regulation of cullin neddylation lethal in Drosophila levels [45]

OTU DUBs cezanne

human

unknown

A20

mouse

otubain1 otubain2 VCIP 135

human human human

severe inﬂammation, premature death unknown unknown unknown

lethal

Ubiquitin-like proteases (ULPs) Ulp1 S. cerevisiae lethal Ulp2

S. cerevisiae

Den1/SENP8 mouse SENP6 human

increased SUMO conjugates, DNA repair defective unknown unknown

Others Apg4B

mouse

unknown

ataxin-3

human

unknown

negative regulation of NF-κB pathway [43] negative regulation of NF-κB pathway [44, 121] editing DUB? [42, 122] undetermined [42] membrane fusion after mitosis [72] regulates cell cycle progression [39] desumoylating enzyme [123]

deneddylates cullins [26, 37] involved in reproductive function? [124] processes autophagy-related UbLs [116] processes polyubiquitin chains? [125]

The organism listed for each DUB refers either to where it was discovered or where the work characterizing the deletion strain and function was performed. DUBs with multiple organism identiﬁers are either highly similar in sequence in each organism or functional homologs. An unknown deletion phenotype indicates that a deletion, knockout, or knockdown of a particular DUB has yet to be generated. No detectable phenotype indicates that a deletion strain has been made, but no phenotypes were observed.

The study of DUBs has moved at a rapid rate since their initial discovery in the early 1980s. Yet despite all the progress, the total number of DUBs and the substrate speciﬁcity of most DUBs are still undetermined. The discovery of novel DUB families including the JAMM isopeptidases and OTU DUBs has highlighted that there may be still more unidentiﬁed DUBs. Because of the large number of potential DUB substrates and the exquisite speciﬁcity that some individual DUBs exhibit (see Figure 1) study of these enzymes has often been challenging. A burgeoning amount of structural data and recent technical advances are being used to address this challenge. The goal of this chapter is to highlight recent developments in the

2 Structure and In Vitro Speciﬁcity of DUB Families 5

DUB ﬁeld by giving an overview of DUB families, including DUBs that act on ubiquitin-like proteins, to discuss how DUBs achieve their speciﬁcity, and to show how the physiological roles of DUBs and their substrates are being elucidated.

2 Structure and In Vitro Speciﬁcity of DUB Families 2.1 Ubiquitin C-terminal Hydrolases (UCH)

The ﬁrst class of DUBs discovered, the ubiquitin C-terminal hydrolases (UCHs), is a relatively small class with only four members in humans and one in budding yeast. UCHs are cysteine proteases related to the papain family of cysteine proteases. Most UCHs consist entirely of a catalytic core that has a molecular mass of about 25 kDa, although Bap1 and UCH37 have C-terminal extensions [21, 22]. All UCHs have a highly conserved catalytic triad consisting of the active-site cysteine, histidine, and aspartate residues that are absolutely required for function [23]. In vitro studies have determined that UCHs have signiﬁcant activity in removing small adducts from the C-terminus of ubiquitin, including short peptides, ethyl ester groups, and amides [24]. They are also very efﬁcient at cotranslationally processing the primary gene products (proubiquitin or Ub-ribosomal subunit fusions) to expose the C-terminal gly–gly motif required for conjugation of ubiquitin and ubiquitin-like proteins to substrates. However, UCHs are unable to cleave the isopeptide bond between ubiquitins in a polyubiquitin chain or to act on ubiquitin conjugated to a folded protein. They are similarly inefﬁcient in acting upon small peptide substrates based on the sequence of the ubiquitin Cterminus [25]. As described below, the binding of ubiquitin is required for optimal UCH activity. Nedd8, a closely related ubiquitin-like protein, is also a substrate for human UCH-L3, albeit with three orders of magnitude less efﬁciency than ubiquitin [26]. The crystal structures of human and yeast UCHs have been solved, the latter in complex with the inhibitor ubiquitin aldehyde [27, 28]. The UCH fold is closely related to that of the papain family of cysteine proteases. Ubiquitin is bound in a cleft on a surface that is highly conserved in all UCHs. NMR studies on the binding of ubiquitin to human UCH-L3 show a similar mode of interaction and deﬁne three regions on the surface of ubiquitin involved in this binding [29]. As noted below and in Figure 2, the same surface of the ubiquitin fold is also involved in binding to USP7 and ULP1. This is remarkable as these DUBs are from different families and not signiﬁcantly homologous in sequence or structure. A second feature of UCHs is the presence of a “blocking loop” spanning the active site and limiting the size of substrates that can be accommodated. Yuh1 and other UCH DUBs contain a mobile, ∼20-residue loop that is disordered in the unliganded protein, but becomes ordered upon substrate binding. The loop passes directly over the active site and the leaving group attached to the gly–gly at

6 The Deubiquitinating Enzymes

Fig. 2 Substrate binding by DUBs revealed by X-ray crystal structures. In the ribbon diagrams, the DUB is represented in white and the substrate in color. The ubiquitin (yellow or green) or SUMO (red) substrates are shown in the same orientation to highlight the similarity of substrate binding by different DUB classes. (A) Ubiqui-

tin (yellow) bound to YUH1. (B) Ubiquitin (green) bound to USP7. (C) SUMO (red) bound to ULP1. (D) Superimposition of substrates from A-C. The regions of each substrate that are within 3.5Å of the DUB when bound are highlighted in color to demonstrate the conserved regions that are recognized by the different DUB classes.

the ubiquitin C-terminus has to pass directly through this loop in order to access the catalytic cysteine. The maximum diameter of this loop was calculated to be ∼15Å, too small for any folded substrate save a single helix [27]. The loop thus allows small substrates to be efﬁciently cleaved, but excludes larger Ub–protein conjugates. This loop explains, at least in part, the preference of UCHs for small or disordered leaving groups.

2 Structure and In Vitro Speciﬁcity of DUB Families 7

2.2 Ubiquitin-speciﬁc Processing Proteases (UBP/USP)

The ubiquitin speciﬁc processing proteases (referred to as UBPs in yeast and USPs in human and mouse) were the second class of DUBs discovered. Catalytically, the UBPs are very similar to the UCHs in that they also utilize the catalytic triad of an active-site cysteine and a conserved histidine and aspartate. The UBP catalytic core of about 400 amino acids contains blocks of conserved sequences (Cys and His boxes) around these catalytic residues [23]. The UBPs are generally larger and more variable in size than the UCH class, ranging from 50 to 300 kDa. Nterminal extensions to the catalytic core account for most of the increased size although a few UBPs have C-terminal extensions. These N-terminal extensions are highly divergent in sequence, unlike the conserved regions of the catalytic core. The sequence and size variations of these extensions are thought to aid in determining UBP localization and substrate speciﬁcity. There are 16 UBPs in yeast and more than 50 USPs identiﬁed in humans, making the UBP/USP family much larger than the UCH family [30]. UBPs also process a wider variety of substrates than UCH DUBs, including proubiquitin, free polyubiquitin chains of various linkages, and mono-or polyubiquitin conjugated to target proteins in vitro and in vivo. In addition, some family members can act on ubiquitin-like proteins. UBP43 has been demonstrated to act on ISG15 while USP21 cleaves conjugated Nedd8 [31, 32]. The diversity of the UBPs and breadth of substrates they act upon, makes them useful in a wide variety of cellular pathways and locations. UBPs regulate apoptosis, DNA repair, endocytosis, and transcription in addition to the ubiquitin–proteasome pathway (see below). The same diversity presents a challenge in determining the speciﬁcity of UBPs and with the exception of Isopeptidase T (UBP14/USP5), there have been few quantitative studies of in vitro speciﬁcity [33, 34]. In general, speciﬁcity has been described with qualitative “yes or no” assays that are not particularly useful in suggesting in vivo roles. The structure of one UBP catalytic domain has been solved, that of USP7 complexed to ubiquitin aldehyde [35]. The data may be applicable to the way in which other UBPs function because the catalytic core of many UBPs is highly conserved. USP7 (also called HAUSP) is a human ubiquitin-speciﬁc protease that regulates the turnover of p53 [36]. USP7 consists of four structural domains; an N-terminal domain known to bind p53 and EBNA1, a catalytic domain, and two C-terminal domains. The 40-kDa catalytic domain exhibits a three-part architecture comprising Fingers, Palm, and Thumb (see Figure 2). The leaving ubiquitin moiety is speciﬁcally coordinated by the Fingers, with its C-terminus placed in a deep cleft between the Palm and Thumb where the catalytic residues are located. The domains form a pocket ideal for binding ubiquitin. Residues in the structure important for the above functions are conserved amongst UBPs, indicating that many UBPs may utilize the Fingers, Palm, and Thumb architecture to bind and cleave ubiquitinated substrates.

8 The Deubiquitinating Enzymes

Another interesting structural observation is that water molecules cushion ubiquitin in the binding pocket. This is necessary because the binding surfaces of ubiquitin are uncharged, and the USP7 binding pocket is made up of predominantly acidic amino acid residues. These water molecules form extensive networks of hydrogen bonds with the bound ubiquitin and USP7. It is possible that they contribute to USP7’s substrate speciﬁcity by allowing the protein to provide for relatively weak binding of ubiquitin and forcing itself to interact with the target protein to achieve speciﬁcity. This seems to be borne out by the fact that ubiquitin does not form a tightly bound complex with USP7 [35]. 2.3 Ubiquitin-like Speciﬁc Proteases (ULP)

The ubiquitin-like speciﬁc proteases (ULPs) are a third class of DUB ﬁrst thought to act only on SUMO-related ubiquitin-like proteins. There are two yeast ULPs and seven human ULPs (also called sentrin speciﬁc proteases, or SENPs). Further analysis determined that ULPs have little or no activity on ubiquitin substrates, but one (SENP8) acts on Nedd8 [26, 37, 38]. Despite acting on non-ubiquitin substrates, ULPs are still classiﬁed as DUBs because the function and mechanism of catalysis is so similar to those of the DUBs that act on ubiquitin. ULPs lack signiﬁcant sequence homology to other DUBs and are more closely related to viral proteinprocessing proteases [39]. In addition to the lack of sequence homology, ULPs have little structural homology to other DUB classes except in the active site. The structure of ULP1 (see Figure 2) in complex with the C-terminal aldehyde of yeast SUMO (SMT3) illustrates that, like most other DUBs, ULPs are thiol proteases, utilizing a conserved catalytic triad consisting of an active-site cysteine, histidine, and aspartate [40]. Also, they require a gly–gly motif at the C-terminus of their UbL substrate for tight binding. The SUMO binding pocket of ULP1 recognizes SUMO through a number of polar and charged-residue interactions, including multiple salt bridges that are not present in the USP7 ubiquitin-binding site, and does not utilize water molecules or a “blocking loop”. 2.4 OTU DUBs

A class of DUBs only identiﬁed since 2002 is the OTU (ovarian tumor protein) DUB class. The OTU domain was originally identiﬁed in an ovarian tumor protein from Drosophila melanogaster, and over 100 proteins from organisms ranging from bacteria to humans are annotated as having an OTU domain. The members of this protein superfamily were annotated as cysteine proteases, but no speciﬁc function had been demonstrated for any of these proteins. The ﬁrst hint of a role for OTU proteins in the ubiquitin pathway was afforded by the observation that an OTUdomain-containing protein, HSPC263, reacted with ubiquitin vinyl sulfone (an active-site-directed irreversible inhibitor of DUBs) [41].

3 DUB Speciﬁcity 9

Then two groups almost simultaneously discovered that several OTU-containing proteins have DUB activity. Two human OTU DUBs were identiﬁed by puriﬁcation with Ub-aldehyde (a reversible DUB inhibitor) afﬁnity resin [42]. These proteins, named otubain1 and 2 (OTU-domain Ub-aldehyde binding protein) have a mass of approximately 35 kDa and are able to cleave polyubiquitin chains in vitro. However, the cleavage mechanism and their true substrates in vivo have yet to be determined. The other OTU DUB found was Cezanne, a 100-kDa protein that is similar to the A20 negative regulator of NF-κB [43]. Like A20, Cezanne plays a role in regulating NF-κB signaling pathways and has general DUB activity [44]. These OTU DUBs have highly conserved catalytic cysteine and histidine residues, implying that they utilize a catalytic triad to catalyze cleavage of polyubiquitin. It is unclear if most proteins containing OTU domains are DUBs, as analysis of the OTU family for DUB activity is only just beginning. 2.5 JAMM Isopeptidases

JAMM isopeptidases also constitute a recently identiﬁed class of DUBs. The members of this interesting class of DUBs were the ﬁrst non-cysteine protease DUBs identiﬁed. Two JAMM isopeptidases have been conﬁrmed as DUBs: Rpn11, which acts on ubiquitin conjugates, and Csn5, which acts on Nedd8 conjugates [45–47]. A number of other eukaryotic proteins have been annotated as containing the JAMM motif, but whether they have DUB activity has yet to be determined. Instead of cysteine proteases, they are metalloproteases belonging to a family of proteins that contain the Jab1/Csn5 and MPN domains [48]. Their activity depends on the JAMM motif (EXn HS/THX7 SXXD) in the JAMM domain. The two histidines and an aspartic acid act as ligands to bind a metal ion, presumably zinc although this has not been proven, to achieve catalysis through polarization of a bound water molecule. A glutamic acid serves as a general acid-base catalyst. The crystal structure of a JAMM metalloprotease from Archaeoglobolus fulgidus bacteria has been recently been solved, but no structures of a JAMM isopeptidase with DUB activity are yet available [49, 50].

3 DUB Speciﬁcity

Why are there so many DUBs and how do they achieve speciﬁcity? The numerous DUBs identiﬁed to date suggest that DUBs have speciﬁcally evolved to act on distinct cellular substrates rather than to have general deubiquitinating activity (see Figure 1). We can ask what common features of these enzymes deﬁne them as DUBs and what differences allow speciﬁc DUBs to act on mono- vs. polyubiquitin? How have they evolved to cleave only ISG15 or SUMO-modiﬁed substrates, for instance? A body of data has been accumulated that at least partially answers these questions.

10 The Deubiquitinating Enzymes

3.1 Recognition of the Ub-like Domain

All DUBs appear to recognize the body of the ubiquitin fold. UCH-L3, for example, makes contact with three regions of ubiquitin; residues 6–12, 41–48, and 69–74 [29]. These surfaces are highly conserved in Nedd8, but divergent in ISG15 and SUMO. Correspondingly, UCH-L3 can cleave ubiquitin and Nedd8 adducts but not those of the other ubiquitin-like proteins [26]. The same regions appear to be important for interactions of ubiquitin with many other DUBs (see Figure 2) and Ub-binding proteins. Importantly, all ubiquitinbinding domains examined utilize these same surfaces in binding ubiquitin. Recognition of a ubiquitin domain can be accomplished by ubiquitin-associated domains (UBA), which are present in many proteins, including some DUBs, and interact with polyubiquitin up to 1000-fold better than mono-ubiquitin [51]. However, other binding domains such as UIM (ubiquitin-interacting motif) and CUE (coupling of ubiquitin conjugation to ER degradation) domains utilized in endocytic pathways prefer binding mono-ubiquitin [52–54]. Polyubiquitin-binding proteins recognize a subset of this binding surface of ubiquitin, often described as the hydrophobic patch. It is a group of three amino acids, Leu8, Ile44, and Val70, which are oriented in the ubiquitin molecule to form a small hydrophobic patch [55]. Polyubiquitin chains incorporating ubiquitins with mutations at residues 8 and 44 were unable to be disassembled by DUBs present in the 19S subunit of the proteasome [56]. In addition to providing a recognition site for DUBs, the patch is also important in determining the quaternary structure of polyubiquitin, another feature utilized by DUBs in substrate recognition. One UbL protein, ISG15, consists of a fusion of two ubiquitin domains. The crude mimicking of an Ub-dimer could potentially contribute to its speciﬁc recognition by deISGylating enzymes. Polyubiquitin chains linked through all seven lysines in ubiquitin have been detected in vivo, and these poorly characterized forms of non-K48-linked polyubiquitin are also likely to have signiﬁcant roles in the cell [57]. Polyubiquitin chains that are linked through different lysines are expected to be different enough in structure that individual DUBs could distinguish between them. K63-linked polyubiquitin is a well characterized alternative linkage and unlike K48-linked polyubiquitin, is not involved in proteolytic degradation [58, 59]. Structural data conﬁrms the idea that these two types of polyubiquitin can have different structures [60, 61]. The structures of these dimers were solved by NMR analysis and they were found to have quite different conformations. Indicative of this, non-hydrolyzable ubiquitindimer analogs containing different linkages have markedly different effectiveness when used to inhibit the enzymatic activity of Isopeptidase T [62]. Isopeptidase T binds and cleaves polyubiquitin linked through at least four of the seven possible chain linkages found in vivo, although the catalytic efﬁciency of these activities is not known [10]. It is interesting to speculate that Isopeptidase T utilizes its two UBA domains to regulate binding of different polyubiquitin substrates. Mutational analysis of the UBA domains and structural data are needed to determine if this is the case and whether it is applicable to other DUBs as well.

3 DUB Speciﬁcity 11

3.2 Recognition of the gly–gly Linkage

The central feature that deﬁnes all DUBs is that they recognize and act at the C-terminus of the ubiquitin or ubiquitin-like domain. All mature ubiquitin and ubiquitin-like proteins have a C-terminal gly–gly motif and DUB cleavage releases leaving groups attached to the carboxyl group of the C-terminal glycine. With the exception of the JAMM metalloproteases, DUB catalysis starts with the nucleophilic attack of the catalytic cysteine on the carbonyl carbon of the scissile bond to form the tetrahedral intermediate. This is converted to an acyl-enzyme intermediate by expelling the C-terminal leaving group. Attack by a water molecule allows regeneration of the free thiol on the catalytic cysteine and releases free ubiquitin. The JAMM isopeptidases appear to use a classical metalloprotease mechanism [50]. DUB structures have evolved to recognize this C-terminal glycylglycine with exquisite speciﬁcity. Analysis of ubiquitin-fusion proteins lacking the gly–gly motif has clearly shown that they are not cleaved efﬁciently by DUBs [63]. All DUBs exclude larger amino acids at the C-terminus of the ubiquitin domain by having a deep cleft in their respective structures that is only large enough to hold two glycines. The narrowest region of USP7’s catalytic cleft sterically excludes amino acids with any type of side chain, enforcing speciﬁcity for ubiquitin conjugates [35]. However, the end of the cleft is open, which allows USP7 to act on large ubiquitin conjugates like its substrate, ubiquitinated p53. ULP1 uses a similar type of cleft to recognize the gly–gly motif except that it uses a tryptophan residue to restrict access to the catalytic site when a substrate is bound to the enzyme [40]. UCHs have a similarly constrained cleft and also use the previously described “blocking loop” to assist in speciﬁcally recognizing the C-terminus of ubiquitin. 3.3 Recognition of the Leaving Group

In principle, DUBs might also recognize the leaving group to which ubiquitin is attached. In fact, such a mechanism seems likely as several DUBs have little afﬁnity for ubiquitin and several have been shown to bind the un-ubiquitinated target protein (see Table 2). Interactions between DUBs and putative substrates have been shown for the mammalian DUBs VDU1, USP11, and UBPy, as well as UBP3 from yeast and fat facets from Drosophila [64–68]. In other cases, DUBbinding proteins may serve as scaffolds or adaptors that localize DUBs (discussed below). 3.4 Substrate-induced Conformational Changes

DUBs are not general hydrolases for cleaving after a gly–gly sequence even though they recognize the gly–gly motif at the C-terminus of ubiquitin and ubiquitin-like proteins. What is so special about these particular gly–gly sequences that DUBs will only recognize and act on them and not others? The answer comes from the fact

12 The Deubiquitinating Enzymes Table 2 Identiﬁcation of DUBs and DUB-binding partners through physical and genetic inter-

action screens DUB

Afﬁnity

Characterized by MS

UCHs UCH37 UCH-L3 UCH-L1 UBPs DOA4 UBP3

Yeast two-hybrid

Interaction partner(s) Synthetic lethal

X X X

X

UBP6 UBP8 USP5 X USP7 USP11 CYLD fat facets UBPy OTU DUBs cezanne otubain1 and 2 X VCIP 135 X JAMM Isopeptidases Rpn11 Others ataxin 3

X

S14, UIP1 [21] Nedd8 [126] JAB1, p27 [127] X X

X X

X X X X X

ubiquitin [43] ubiquitin aldehyde [42] VCP/P47 [72] X

X

SLA1, SLA2 [128] SIR4, Bre5, Stu1 [66, 118, 129] 19S proteasome [130] SAGA, SLIK acetyl transferases [88] ubiquitin [131] ataxin [132] RanBPM [67] NEMO [7] Vasa [75] CDC25(Mm) [133]

UBP6 [92] RAD23, HHR23A, HHR23B [134]

that DUBs interact with the rest of the ubiquitin or ubiquitin-like substrate, and this interaction causes conformational changes in the DUB that are necessary to achieve catalysis. These changes result in rapid and efﬁcient cleavage of only the particular substrate that the DUB is equipped to bind. It also explains why peptides with a gly–gly in them are not susceptible to cleavage by DUBs as they are lacking the substrate-binding domains that cause the DUB conformational change required for cleavage. The different DUB classes utilize a number of conformational changes that are induced upon substrate binding to assist in promoting efﬁcient cleavage. UCH DUBs have been the most thoroughly analyzed. Comparison of the ubiquitin—UCH complex with unliganded UCH shows two signiﬁcant conformational differences that contribute to keeping the unliganded enzyme in an inactive state. First, the previously described “blocking loop” becomes ordered as it interacts with ubiquitin. Invariant residues form hydrogen bonds with the ubiquitin substrate and other UCH residues, indicating that the loop has functional importance during substrate binding [27]. Second, the side chain of L9 in UCH-L3 intrudes into

4 Localization of DUBs 13

the substrate-binding cleft, occluding the catalytic cysteine and preventing binding of peptide substrates [29]. When ubiquitin binds to the UCH-L3, an interaction between ubiquitin and UCH-L3 repositions L9, allowing access to the active site cleft. Thus, the energy of ubiquitin binding is required to activate UCH-L3, allowing its cleavage. This type of selectivity (where ubiquitin binding is required for activity) may be necessary to prevent deleterious cleavage of other protein substrates by UCHs. A similar situation was observed when the crystal structure of USP7 was solved in the absence of substrate [35]. The catalytic cysteine of the unliganded protein is not in an orientation that would allow catalysis to take place. The histidine residue needed to interact with the active-site cysteine is too distant for a catalytic-triad mechanism to function. Binding of ubiquitin aldehyde induces a signiﬁcant conformational change that realigns the catalytic triad residues so the hydrogen bonding required for catalysis can take place. Thus, like UCH DUBs, the unliganded protease is inactive and only becomes catalytically active when it is binding substrates. ULP1 also uses conformational changes to “clamp down” on the gly–gly motif when a SUMO substrate is bound. Trp448 lies directly above the active site and interacts with the SUMO C-terminus by Van der Waals interactions, sandwiching the gly–gly motif between Trp448 and the active-site cysteine when SUMO binds [40]. Despite the various methods utilized, all DUBs require a conformational change triggered by binding of a speciﬁc substrate to catalyze cleavage. These required conformational changes are driven by the energy of interaction between the DUB and the body of the ubiquitin domain.

4 Localization of DUBs

While many DUBs are cytoplasmic, localization of DUBs is also known to be important in regulating DUB speciﬁcity. The localization of ULP1, for example, is important in determining its substrates. The N-terminal domain of ULP1 is known to localize the enzyme to the nuclear envelope, and truncation mutations lacking this domain remain in the cytoplasm [69]. When the truncated protein is expressed in ULP1 yeast strains, the cells grow at wild-type levels, and the truncated protein is able to cleave SUMO substrates in vitro. However, analysis of ULP1 cells expressing this truncation shows an accumulation of SUMO conjugates. Apparently, the localization of ULP1 to the nuclear envelope is necessary in order for it to act on speciﬁc nuclear-envelope-localized substrates. The localization helps constrain ULP1 isopeptidase activity so ULP1 does not inappropriately act on cytoplasmic substrates. Other examples of DUB activity regulated by localization include UBP6, which is fully active only when bound to the proteasome (see below) and UBP16 residence on the outer membrane of the mitochondria, although its function there is undetermined [70]. Other DUBs have been found to associate with membranes and regulate membrane-associated cellular processes, although they appear not to be membrane

14 The Deubiquitinating Enzymes

anchored like UBP16. The ability of DOA4 to remove ubiquitin from membranebound endocytic substrates promotes their degradation in the vacuole or lysosome [71]. DUBs are also important for membrane fusion events as shown by the fact that an OTU domain DUB, VCIP135 (VCP/p47 complex-interacting protein of 135kd), is necessary for p97-p47-mediated Golgi cisternae reassembly after mitosis [72]. Also, a neuronal DUB, synUSP, was found to localize to post-synaptic lipid rafts (membrane microdomains involved in membrane trafﬁcking and signal transduction) [73]. However, its function at that location has yet to be characterized. A well-studied example of a tissue-speciﬁc DUB activity is fat facets, a UBP originally found in Drosophila[74]. It is important in eye development and germcell speciﬁcation and is active only in speciﬁc cell types during certain stages of development [65, 75]. The lack of fat facets results in defective posterior patterning, germ-cell speciﬁcation, and eye formation. Fat facets activity is required to prevent the inappropriate degradation of vasa and liquid facets. In this case, the role of the DUB appears to be deﬁned by the restricted expression of its known substrates. Temporal regulation of DUB expression also appears important. D’Andrea and colleagues ﬁrst described a small family of DUBs that are induced as immediate early gene products of cytokine stimulation [76]. Different cytokines were shown to induce different DUBs and the expression of these enzymes was short-lived [77]. It appears that these DUBs may be involved in down-regulating cytokine receptors, perhaps by removing the ubiquitin involved in sorting of the receptor at the early endosome. Likewise, UBP43, the short-lived processing protein for ISG15, is present at very low levels in normal cells and highly expressed upon interferon induction [78].

5 Probable Physiological Roles for DUBs 5.1 Proprotein Processing

One important function of DUBs is the processing of ubiquitin or ubiquitin-like proteins to their mature forms. Ubiquitin is expressed in cells as either linear polyubiquitin or N-terminally fused to certain ribosomal proteins [79, 80]. These gene products are processed by DUBs to separate the ubiquitin into monomers and expose the gly–gly motif at the C-terminus. Many DUBs process linear polyubiquitin or Ub-fusion proteins in vitro, but this processing appears to take place cotranslationally in vivo and is extremely rapid. This makes analysis difﬁcult and leaves unanswered the question of which DUBs actually perform this function in vivo. Multiple DUBs may be able to perform this processing at a physiologically relevant level since DUB deletions rarely shows processing defects [81]. Ubiquitin-like proteins are also expressed as proproteins with a short C-terminal extension of a few amino acids that must be removed to make the UbL available for conjugation to target proteins. All ULPs have been shown to metabolize their

5 Probable Physiological Roles for DUBs 15

respective proprotein to an active form in vitro although again it is unclear which ULPs are responsible for this activity in vivo. An exception to the confusion is the ﬁnding that RUB1 (the yeast homolog of Nedd8) is processed by YUH1 in Saccharomyces cerevisiae. Conjugation of RUB1 to Cdc53 is required for efﬁcient assembly of certain SCF (skp1, cullin, F-box) E3 ubiquitin ligases [82]. Yuh1 deletion strains do not process Rub1 or modify Cdc53 with Rub1 [83]. Modiﬁcation of Cdc53 by Rub1 could be reconstituted in a Yuh1 strain by expressing a mature Rub1 construct lacking the C-terminal asparagine normally removed by processing. This demonstrated that Yuh1 processes RUB1 proprotein into the mature form in vivo. It is not known which DUB performs the processing of proNedd8. 5.2 Salvage Pathways: Recovering Mono-ubiquitin Adducts and Recycling Polyubiquitin

It has been speculated that without UCH function, all ubiquitin in the cell would be conjugated with glutathione or other cellular amines and therefore unavailable for conjugation. This would quickly result in the cessation of the ubiquitin–proteasome system function and cell death due to lack of active ubiquitin to conjugate to substrates. The effectiveness of UCH DUBs in liberating ubiquitin from other small adducts makes them likely candidates to act on these particular adducts. In addition, the cell must regenerate mono-ubiquitin from polyubiquitin and various mono-ubiquitinated proteins to maintain levels of mono-ubiquitin for conjugation. Doa4 appears to remove small peptides attached to mono- and di-ubiquitin intermediates resulting from proteasomal degradation as well as removing ubiquitin from proteins targeted for endocytosis [84, 85]. Loss of Doa4 function in yeast results in depleted levels of mono-ubiquitin and increased cell death during stationary phase. Another function for DUBs is regenerating free ubiquitin from unanchored polyubiquitin chains removed from proteasome substrates or proteins targeted for other pathways. Polyubiquitin inhibits the proteasome and lowers the amount of free ubiquitin available for conjugation to proteins. Thus, these chains need to be processed to mono-ubiquitin to prevent polyubiquitin accumulation and inhibition of the proteasome. This type of DUB activity has been well characterized in vivo and in vitro and Isopeptidase T appears to be the DUB that is responsible for the majority of this activity. Deletion of UBP14 in yeast is not lethal, although large amounts of polyubiquitin build up in the cell and proteasome function is impaired [86]. Isopeptidase T seems to serve as a general DUB for regenerating mono-ubiquitin as it cleaves polyubiquitin containing various linkages [59]. 5.3 Regulation of Mono-ubiquitination

DUBs have increasingly been found to be important in regulating the ubiquitination level of proteins not targeted to the proteasome for degradation. Some DUBs are active participants in the regulation of mono-ubiquitin (or mono-UbL) conjugation and others can regulate the conjugation of multiple types of ubiquitin or UbLs

16 The Deubiquitinating Enzymes

to a single substrate. For instance, deneddylating enzymes may regulate the neddylation of cullin proteins both by processing proNedd8 and by removing Nedd8 from neddylated cullins [26, 38]. As a component of the SCF E3 ligase complexes, cullins require neddylation in order for their cognate E3 ligase to be efﬁciently assembled [82]. Regulation of this modiﬁcation indirectly regulates the ubiquitination of a subset of proteins. Defects in deneddylation could lead to inappropriate ubiquitination of substrates owing to inappropriate recruitment of E2s to the SCF E3s [87]. Regulating mono-ubiquitination of proteins by DUBs is important in histone modiﬁcation where ubiquitination is thought to modulate chromatin structure and transcriptional activity. Normally, about 10% of the histone core octomers contain ubiquitinated histones and the ubiquitin is removed at mitosis by DUB activity. UBP8 has been demonstrated to regulate the ubiquitination of histone H2B, which is important in transcriptional activation of many genes [88]. Many cell-surface receptors are ubiquitinated upon internalization and the ubiquitin is removed by DUBs at the early endosome. Properly sorted receptors are then shuttled to the lysosome for degradation. In the absence of Doa4, the ubiquitin is not removed upon sorting and instead is co-degraded in the vacuole, resulting in ubiquitin depletion [84]. Another DUB, UBP3 assists Golgi-ER retrograde transport by deubiquitinating B -COP, thus preventing its degradation [66]. Mono-ubiquitinated B -COP cannot be assembled into the COP1 complex without UBP3/Bre5 complex DUB activity. Disruption of the complex in Bre5 strains reduces the efﬁciency of Golgi-ER transport and facilitates the polyubiquitination and degradation of B -COP by the proteasome. One fascinating observation is that PCNA (proliferating cell nuclear antigen) can be modiﬁed by multiple forms of ubiquitin, demonstrating that DUBs with different speciﬁcities can act at the same location on a speciﬁc substrate. PCNA can be modiﬁed by mono-ubiquitin, 63-linked polyubiquitin, or SUMO at K164 [89]. Modiﬁcation of PCNA by mono- or polyubiquitin determines whether it is utilized in translesion synthesis or error-free DNA repair, respectively. SUMO modiﬁcation prevents PCNA function in DNA repair and instead promotes DNA replication. It is probable that multiple DUBs, as yet unidentiﬁed, are required to regulate PCNA modiﬁcation. 5.4 Processing of Proteasome-bound Polyubiquitin

DUBs play a crucial role in regulating the function of the proteasome. For a long time it was unclear what happens to polyubiquitin conjugated to a proteasome substrate when that substrate is at the proteasome ready for degradation. Was the conjugated polyubiquitin processed by the proteasome and degraded or was it removed by a DUB and released from the proteasome? The small 13-Å diameter entrance to the 20S catalytic core of the proteasome requires all substrates to be fed through as unfolded polypeptides [90]. A branched polypeptide such as a ubiquitin–protein conjugate apparently has difﬁculty ﬁtting through the pore,

6 Finding Substrates and Roles for DUBs 17

greatly reducing proteasome efﬁciency [47]. Thus, it seemed likely that DUBs must remove polyubiquitin from proteasome substrates before they enter the 20S catalytic core of the proteasome. To date, three DUBs are known to perform this function and all are components of the 19S lid of the mammalian proteasome. The ﬁrst described was UCH37, although its exact function is still unclear [91]. UCH37 is thought to be an editing DUB that assists in clearing the proteasome of ubiquitinated proteins. UCH37 slowly cleaves one ubiquitin at a time from the distal end of the polyubiquitin chain. If chain trimming is faster than the degradation process, loss of the polyubiquitin signal could result in partial degradation or release of proteins from the proteasome. UCH37 activity could also be necessary to recover proteasomes that are having difﬁculty degrading ubiquitinated proteins. UCH37 is only found in higher eukaryotes, but the other two proteasome-bound DUBs, RPN11 and UBP6 (USP14), are found in all eukaryotes. RPN11 and UBP6 remove polyubiquitin from substrates that are committed to degradation by the proteasome [92]. The mechanisms for this, and exactly what role each DUB plays in removing ubiquitin, are not fully understood. Interestingly, the Rpn11 DUB activity was ﬁrst detected over 10 years ago when 26S proteasome DUB activity was inhibited by o-phenanthroline, a metal chelator [93]. The metalloprotease DUB activity was not identiﬁed until recently [46]. Rpn11 is thought to remove polyubiquitin chains from proteasome substrates before they are degraded, allowing the unfolded substrate to enter the pore of the 20S subunit of the proteasome. It has been proposed that Rpn11 removes most of the polyubiquitin chain attached to a proteasome substrate and then UBP6 acts to remove the remaining one or two ubiquitin residues. Despite the lack of mechanistic understanding, the DUBs are clearly required for efﬁcient proteasomal degradation to take place. The Rpn11 deletion is lethal in yeast and temperature-sensitive mutants show massive accumulation of polyubiquitin conjugates [46, 47]. UBP6 is approximately 300 times more active when it is associated with the proteasome than in its puriﬁed form [94]. The UBP6 deletion is not lethal in yeast, but a large decrease in the cellular pool of mono-ubiquitin occurs, indicating that ubiquitin is fed into the proteasome and degraded rather than being released from the proteasome and recycled [95].

6 Finding Substrates and Roles for DUBs

Surprisingly, little is known about the in vivo substrate speciﬁcity of DUBs. Difﬁculty in deﬁning the substrate speciﬁcity of individual DUBs often arises from a lack of observable phenotypes in deletion strains. Deletion studies in yeast where up to 4 of the 17 DUBs have been deleted in a single strain have not produced signiﬁcant phenotypes [96]. It is unclear if this is due to the subtle nature of the phenotypes or if the remaining DUBs compensate for the missing ones. However,

18 The Deubiquitinating Enzymes

several tactics have been fruitful in deﬁning the physiological roles of DUBs. First, deﬁnition of in vitro speciﬁcities can be useful in focusing genetic screens. For example, the ﬁrst UBPs were cloned and analyzed after it was discovered that they could cleave ubiquitin-fusion proteins [96, 97]. Second, directed screening of deletions or knockdown studies to identify roles for DUBs have also been successful (see Table 1 for DUB-deletion phenotypes). Study of DUB deletions, including UCH-L1, UCH-L3, and USP14 (see below), in the mouse have demonstrated their importance in neuronal function. Third, potential roles for DUBs have also been identiﬁed by physical and genetic interaction screens. Table 2 shows in more detail the interaction screens that have been used in discovering DUBs and characterizing their in vivo roles by identifying novel binding partners. For example, Cezanne was suggested to be a DUB after two-hybrid studies demonstrated its interaction with ubiquitin and UBP6 was identiﬁed as a component of the 19S subunit of the proteasome by mass spectrometry.

7 Roles of DUBs Revealed in Disease 7.1 NF-jB Pathway

NF-κB is a transcription factor that can be activated by a number of cellular signals, including stress, inﬂammation (via tumor necrosis factor) and antigen receptors among others [98, 99]. After receptor stimulation, a cascade ensues that results in the release of NF-κB from its inhibitor IκB. Released NF-κB translocates to the nucleus and activates transcription of a number of genes. Ubiquitin metabolism plays a signiﬁcant regulatory role in the NF-κB pathway. For NF-κB release from IκB and nuclear translocation to take place, IκB is phosphorylated by IκB kinases, resulting in K48-linked polyubiquitination and proteasomal degradation of IκB [100]. A number of other proteins involved in this pathway such as NEMO, IKKγ , and TRAF6 have K63-linked polyubiquitin chains conjugated to them [101, 102]. It is not clear what purpose the K63-linked chains serve, but they appear to be a regulatory component of the NF-κB pathway. Most of the DUB activity characterized in the NF-κB pathway appears to act on K63-linked polyubiquitin, suggesting that modulation of K63-linked polyubiquitination by DUBs is important for control of the NF-κB pathway. CYLD, a tumor suppressor gene, has been conﬁrmed as a DUB [7, 103, 104]. Loss of CYLD function leads to cylindromatosis, a syndrome characterized by large benign tumors on the face and neck. This is one of the few examples where a defective DUB has been deﬁned as the direct cause of a speciﬁc disease. Preferred in vivo substrates of CYLD are believed to be 63-linked polyubiquitin-protein conjugates of NEMO, TRAF6, and TRAF2 components of the NF-κB pathway, but the exact in vivo regulatory role of CYLD is still unknown.

7 Roles of DUBs Revealed in Disease

OTU family DUBs such as Cezanne and A20 also play signiﬁcant roles as negative regulators of the NF-κB pathway [43, 44]. A20 can cleave K48- and K63-linked polyubiquitin chains in vitro while Cezanne has only been tested on K48-linked chains. Although these DUBs are known to be part of the NF-κB pathway, their in vivo substrates are unknown. It is also unclear as to how these DUBs negatively regulate the NF-κB pathway. 7.2 Neural Function

DUBs, speciﬁcally UCHs, appear to play signiﬁcant roles in neurodegenerative diseases such as Parkinson’s, Alzheimer’s, Huntington’s, and others [105]. A mutant form of UCH-L1 with reduced enzymatic activity has been found in a small family of Parkinson’s patients and the S18Y allele of UCH-L1 has been associated with a reduced risk of sporadic Parkinson’s disease [106, 107]. Many of the inclusion bodies found in patients with Parkinson’s are known to contain high amounts of UCH-L1, ubiquitin, and ubiquitinated proteins, as determined by immunostaining [108]. This suggests that defects in some DUBs or their regulation can cause signiﬁcant harm to the neuronal system, resulting in disease. DUBs have also been implicated in the formation of other neural inclusion bodies. In addition to the case for their involvement in Parkinson’s disease it has been shown that the mutation of USP14 (the mammalian homolog of yeast UBP6) results in Ataxia in the mouse [109]. Many neurological diseases, including Ataxia, result in damaged or mutated proteins aggregating as polyubiquitinated forms at the microtubule organizing center (MTOC) to form inclusions called aggresomes [110]. An adapter, the tubulin deacetylase HDAC6 (histone deacetylase 6), has recently been shown to bind these polyubiquitinated proteins and tether them to the microtubules where they are then transported to the MTOC [111]. The classic Lewy Body of Parkinson’s disease has all the hallmarks of such an aggresome. The formation of an aggresome is thought to be protective and in its absence the aggregated proteins can trigger apoptosis. Thus, the dynamics of ubiquitination and aggregate formation are important responses to this type of cellular stress and several DUBs can modulate this process. Deletion of UCH-L1 and/or UCH-L3 in mice has demonstrated that they are both involved in neuronal regulation, but have separate functions. The GAD (gracile axonal dystrophy) mouse has been shown to lack UCH-L1, the predominant neuronal UCH [112]. These mice show a unique neuronal “dying back” phenotype that results in paralysis of the limbs due to death of nerves originating in the gracile nucleus. Mice lacking UCH-L3 have no obvious abnormalities or defects [113]. However, the double deletion mouse shows more severe defects including reduced weight, a more severe gracile axonal dystrophy than the L1 deletion, and earlier lethality caused by a loss of the ability to swallow resulting in starvation [114]. This demonstrates that the two DUBs are not redundant and have separate neuronal functions.

19

20 The Deubiquitinating Enzymes

8 New Tools for DUB Analysis

Despite all the DUB structures and substrates previously described, in most cases the in vivo substrate for a particular DUB is unknown. Structural and localization data can provide clues to determine in vivo DUB speciﬁcity, especially if one knows what ubiquitin or ubiquitin-like protein it acts upon. Genomic databases have helped, but many annotated DUBs have never been tested for DUB activity and some DUBs thought to act on one type of substrate (based on their homology) are found to act on another when tested. The characterization of hundreds of potential DUBs is a daunting task and in vivo characterization is even more difﬁcult. To make headway, novel tools are needed to conclusively identify potential DUBs and their substrates to help direct appropriate in vivo studies.

8.1 Active-site-directed Irreversible Inhibitors and Substrates

The most promising tools developed for this sort of analysis are active-site-directed irreversible inhibitors of DUBs. These inhibitors are ubiquitin or ubiquitin-like proteins chemically modiﬁed at the C-terminus by an electrophilic moiety such as a Michael acceptor or alkyl halide. The modiﬁed ubiquitin can be incubated with a puriﬁed DUB or a cell lysate containing DUB activity. Ubiquitin vinyl sulfone (UbVS) is one such irreversible inhibitor because the vinyl sulfone moiety reacts with the active-site cysteine of the DUB, forming a thioether linkage. The covalent adduct is stable and can be detected in a variety of ways. Labeling of DUBs is speciﬁc, as only a DUB active-site cysteine will efﬁciently react with the vinyl sulfone moiety. To create these inhibitors, an N-terminally tagged ubiquitin or ubiquitin-like protein (lacking the C-terminal glycine) is expressed using the intein expression system (New England Biolabs). Brieﬂy, in this system a fusion protein consisting of a ubiquitin or a ubiquitin-like protein lacking the C-terminal glycine, an intein linker, and a chitin-binding domain (CBD) is expressed in E. coli. Clariﬁed cell lysate is incubated with chitin beads to bind the ubiquitin-fusion protein. The ubiquitin is then cleaved from the CBD and intein linker by adding mercaptoethanesulfonic acid (MESNA). After MESNA elution, the resulting truncated ubiquitin C-terminal thioester is reacted with glycine vinyl methyl sulfone to create the Ub or UbL vinyl sulfone derivative. The N-terminal tag on the ubiquitin molecule allows analysis of DUB labeling by immunoprecipitation and Western blotting. This labeling has been used with success in yeast where 6 of the 17 known DUBs were labeled with UbVS [115]. Incomplete labeling likely results from DUBs that do not act on mono-ubiquitin or where the UbVS could not access the active site. The labeling has also been used with great success in mammalian cell lysates to identify novel ubiquitin DUBs [41]. A novel deneddylating enzyme and a novel DUB that acts on autophagy-related UbL proteins have also been identiﬁed using vinyl sulfone labeled probes [26, 37, 116].

8 New Tools for DUB Analysis 21

This ubiquitin intein system can also be utilized to make a DUB substrate rather than inhibitors by attaching a C-terminal ﬂuorescent tag such as 7amidomethylcoumarin (AMC) instead of vinyl sulfone. DUBs cleave the ubiquitin derivative and release the ﬂuorescent tag, a process that can be followed ﬂuorometrically. Fluorometric assays can then be used to determine a particular DUB’s preferred substrate or to quantitate DUB activity in crude lysates. AMC substrates have turned out to be excellent tools for identifying the substrates of individual DUBs. Den1, for example, was shown to cleave Nedd8-AMC 60 000-fold faster than it cleaves ubiquitin–AMC, and the ratio was even higher when compared to SUMO–AMC [26]. Clearly, these reagents are powerful tools for identifying novel DUBs and identifying potential DUB substrates.

8.2 Non-hydrolyzable Polyubiquitin Analogs

Other modiﬁed ubiquitin reagents that are useful in analyzing DUBs are nonhydrolyzable polyubiquitin analogs. These analogs are polyubiquitin chains where the ubiquitins are linked by cross-linking reagents. To synthesize them, one ubiquitin is mutated to cysteine at the C-terminal glycine and another has cysteine introduced at a particular lysine residue. These ubiquitins can then be linked through their cysteine residues with a bifunctional thiol reagent such as dichloroacetone (DCA). As the native ubiquitin sequence contains no cysteines, the ubiquitins will only be linked through the introduced cysteine residues. The result is a ubiquitin dimer analog that mimics physiological dimers. The isopeptide bond is replaced by a DCA linkage, but the ubiquitin subunits retain the appropriate spatial orientation. Thus, DUBs should bind these dimers, but will be unable to cleave them because they cannot hydrolyze the DCA linkage. To make longer polyubiquitin chains, cysteine residues or sulfhydryl groups must be introduced at the desired lysine and the C-terminal glycine on the same ubiquitin molecule. These chain analogs have been used to characterize DUBs in two fashions. First, they can be used as inhibitors of DUBs [62]. Cleavage of Ub–AMC by Isopeptidase T, a polyubiquitin-binding DUB, was inhibited by the addition of differently linked dimer analogs and kinetic inhibition constants were determined. The K i values of dimer analogs were all much lower than the K i for mono-ubiquitin. Further, there was considerable selectivity, as inhibition constants varied depending on the linkage present in the dimer [62]. This demonstrated that the analogs act as faithful mimics of native polyubiquitin. The other way these chain analogs are used is to synthesize them on afﬁnity supports and analyze cell lysates for DUBs that bind a speciﬁc type of polyubiquitin chain. These afﬁnity resins have proven useful in identifying a number of binding proteins, including DUBs, from yeast cell lysates [117]. Analogs with different linkages bind a different subset of proteins, perhaps suggesting a way to identify DUBs acting upon polyubiquitin with linkage speciﬁcity. Thus, these analogs are excellent tools for characterizing the substrate preferences of known DUBs and discovering novel ones.

22 The Deubiquitinating Enzymes

9 Conclusion

DUBs are clearly an essential cellular component needed for a variety of pathways including protein degradation, DNA repair, apoptosis, membrane trafﬁcking, stress response, and transcriptional regulation. Not only do they act on various ubiquitin substrates, but they are also needed to process ubiquitin-like substrates. Over 100 DUBs from ﬁve major families have been identiﬁed and the number is likely to increase. Factors that enhance DUB speciﬁcity are the presence of a binding pocket that only accommodates the physiological substrate, the requirement for a substrate-induced conformational change that prevents undesired catalysis, and the recognition of the ubiquitous C-terminal gly–gly motif of all DUB substrates. Subcellular localization to a speciﬁc organelle or protein complex and tissue-speciﬁc as well as temporal expression are also important components of DUB speciﬁcity and function. In spite of all that has been learned about DUBs and their function, much remains to be discovered. Future studies are likely to focus on identifying in vivo DUB substrates, novel DUBs, DUB-binding partners, and phenotypes of DUB deletions. Further development of new reagents, such as the non-hydrolyzable polyubiquitin analogs and active-site-directed inhibitors or substrates will help greatly. Directed proteomics studies should assist in identifying DUBs, loss-offunction phenotypes, and potential binding partners. Despite the major gaps that remain in our understanding of DUBs, our knowledge of their roles and importance has progressed amazingly rapidly. Novel DUB gene families have been identiﬁed, new ubiquitin-like DUB substrates have been uncovered, and structural data has been analyzed to elucidate how DUBs perform catalysis and speciﬁcally recognize their substrates. In vivo substrates of DUBs are beginning to be identiﬁed and the tools and techniques needed to search for novel DUBs and analyze known ones for their speciﬁcity are rapidly being created. With so much discovered, and yet so much remaining to be found, deubiquitinating enzymes are a vibrant ﬁeld of study.

References 1 SCHLESINGER, D. H., GOLDSTEIN, G., and NIALL, H. D. The complete amino acid sequence of ubiquitin, an adenylate cyclase stimulating polypeptide probably universal in living cells, Biochemistry, 1975, 14, 2214–2218. 2 HOCHSTRASSER, M. Ubiquitin-dependent protein degradation, Annu Rev Genet, 1996, 30, 405–439. 3 THROWER, J. S., HOFFMAN, L., RECHSTEINER, M., and PICKART, C. M. Recognition of the polyubiquitin

proteolytic signal, Embo J, 2000, 19, 94–102. 4 WILKINSON, K. D. Ubiquitination and deubiquitination: targeting of proteins for degradation by the proteasome, Semin Cell Dev Biol, 2000, 11, 141– 148. 5 WANG, Q., GOH, A. M., HOWLEY, P. M., and WALTERS, K. J. Ubiquitin recognition by the DNA repair protein hHR23a, Biochemistry, 2003, 42, 13529–13535.

References 6 GEWIES, A. and GRIMM, S. UBP41 is a proapoptotic ubiquitin-speciﬁc protease, Cancer Res, 2003, 63, 682–688. 7 KOVALENKO, A., CHABLE-BESSIA, C., CANTARELLA, G., ISRAEL, A., WALLACH, D., and COURTOIS, G. The tumour suppressor CYLD negatively regulates NF-kappaB signalling by deubiquitination, Nature, 2003, 424, 801–805. 8 HICKE, L. A new ticket for entry into budding vesicles-ubiquitin, Cell, 2001, 106, 527–530. 9 MASTRANDREA, L. D., YOU, J., NILES, E. G., and PICKART, C. M. E2/E3-mediated assembly of lysine 29-linked polyubiquitin chains, J Biol Chem, 1999, 274, 27299–27306. 10 YOU, J. and PICKART, C. M. A HECT domain E3 enzyme assembles novel polyubiquitin chains, J Biol Chem, 2001, 276, 19871–19878. 11 BABOSHINA, O. V. and HAAS, A. L. Novel multiubiquitin chain linkages catalyzed by the conjugating enzymes E2EPF and RAD6 are recognized by 26 S proteasome subunit 5, J Biol Chem, 1996, 271, 2823–2831. 12 MAHAJAN, R., DELPHIN, C., GUAN, T., GERACE, L., and MELCHIOR, F. A small ubiquitin-related polypeptide involved in targeting RanGAP1 to nuclear pore complex protein RanBP2, Cell, 1997, 88, 97–107. 13 KAMITANI, T., KITO, K., NGUYEN, H. P., and YEH, E. T. Characterization of NEDD8, a developmentally down-regulated ubiquitin-like protein, J Biol Chem, 1997, 272, 28557–28562. 14 NARASIMHAN, J., POTTER, J. L., and HAAS, A. L. Conjugation of the 15-kDa interferon-induced ubiquitin homolog is distinct from that of ubiquitin, J Biol Chem, 1996, 271, 324–330. 15 WALDEN, H., PODGORSKI, M. S., HUANG, D. T., MILLER, D. W., HOWARD, R. J., MINOR, D. L., Jr., HOLTON, J. M., and SCHULMAN, B. A. The structure of the APPBP1-UBA3-NEDD8-ATP complex reveals the basis for selective ubiquitin-like protein activation by an E1, Mol Cell, 2003, 12, 1427–1437.

16 JOHNSON, E. S., SCHWIENHORST, I., DOHMEN, R. J., and BLOBEL, G. The ubiquitin-like protein Smt3p is activated for conjugation to other proteins by an Aos1p/Uba2p heterodimer, Embo J, 1997, 16, 5509–5519. 17 HEMELAAR, J., BORODOVSKY, A., KESSLER, B. M., REVERTER, D., COOK, J., KOLLI, N., GAN-ERDENE, T., WILKINSON, K. D., GILL, G., LIMA, C. D., PLOEGH, H. L., and OVAA, H. Speciﬁc and covalent targeting of conjugating and deconjugating enzymes of ubiquitin-like proteins, Mol Cell Biol, 2004, 24, 84–95. 18 ANDERSEN, M. W., GOLDKNOPF, I. L., and BUSCH, H. Protein A24 lyase is an isopeptidase, FEBS Lett, 1981, 132, 210–214. 19 PICKART, C. M. and ROSE, I. A. Ubiquitin carboxyl-terminal hydrolase acts on ubiquitin carboxyl-terminal amides, J Biol Chem, 1985, 260, 7903–7910. 20 JENTSCH, S. and PYROWOLAKIS, G. Ubiquitin and its kin: how close are the family ties?, Trends Cell Biol, 2000, 10, 335–342. 21 LI, T., DUAN, W., YANG, H., LEE, M. K., BTE MUSTAFA, F., LEE, B. H., and TEO, T. S. Identiﬁcation of two proteins, S14 and UIP1, that interact with UCH37, FEBS Lett, 2001, 488, 201–205. 22 JENSEN, D. E., PROCTOR, M., MARQUIS, S. T., GARDNER, H. P., HA, S. I., CHODOSH, L. A., ISHOV, A. M., TOMMERUP, N., VISSING, H., SEKIDO, Y., MINNA, J., BORODOVSKY, A., SCHULTZ, D. C., WILKINSON, K. D., MAUL, G. G., BARLEV, N., BERGER, S. L., PRENDERGAST, G. C., and RAUSCHER, F. J., 3rd. BAP1: a novel ubiquitin hydrolase which binds to the BRCA1 RING ﬁnger and enhances BRCA1-mediated cell growth suppression, Oncogene, 1998, 16, 1097–1112. 23 WILKINSON, K. D. Regulation of ubiquitin-dependent processes by deubiquitinating enzymes, Faseb J, 1997, 11, 1245–1256. 24 LARSEN, C. N., KRANTZ, B. A., and WILKINSON, K. D. Substrate speciﬁcity of deubiquitinating enzymes: ubiquitin C-terminal hydrolases, Biochemistry, 1998, 37, 3358–3368.

23

24 The Deubiquitinating Enzymes

25 DANG, L. C., MELANDRI, F. D., and STEIN, R. L. Kinetic and mechanistic studies on the hydrolysis of ubiquitin C-terminal 7-amido-4-methylcoumarin by deubiquitinating enzymes, Biochemistry, 1998, 37, 1868–1879. 26 GAN-ERDENE, T., KOLLI, N., YIN, L., WU, K., PAN, Z. Q., and WILKINSON, K. D. Identiﬁcation and characterization of DEN1, a deneddylase of the ULP family, J Biol Chem, 2003, 31, 28892–28900. 27 JOHNSTON, S. C., RIDDLE, S. M., COHEN, R. E., and HILL, C. P. Structural basis for the speciﬁcity of ubiquitin C-terminal hydrolases, Embo J, 1999, 18, 3877–3887. 28 JOHNSTON, S. C., LARSEN, C. N., COOK, W. J., WILKINSON, K. D., and HILL, C. P. Crystal structure of a deubiquitinating enzyme (human UCH-L3) at 1.8 A resolution, Embo J, 1997, 16, 3787–3796. 29 WILKINSON, K. D., LALELI-SAHIN, E., URBAUER, J., LARSEN, C. N., SHIH, G. H., HAAS, A. L., WALSH, S. T., and WAND, A. J. The binding site for UCH-L3 on ubiquitin: mutagenesis and NMR studies on the complex between ubiquitin and UCH-L3, J Mol Biol, 1999, 291, 1067–1077. 30 WING, S. S. Deubiquitinating enzymes – the importance of driving in reverse along the ubiquitin-proteasome pathway, Int J Biochem Cell Biol, 2003, 35, 590–605. 31 MALAKHOV, M. P., MALAKHOVA, O. A., KIM, K. I., RITCHIE, K. J., and ZHANG, D. E. UBP43 (USP18) speciﬁcally removes ISG15 from conjugated proteins, J Biol Chem, 2002, 277, 9976–9981. 32 GONG, L., KAMITANI, T., MILLAS, S., and YEH, E. T. Identiﬁcation of a novel isopeptidase with dual speciﬁcity for ubiquitin- and NEDD8-conjugated proteins, J Biol Chem, 2000, 275, 14212–14216. 33 WILKINSON, K. D., TASHAYEV, V. L., O’CONNOR, L. B., LARSEN, C. N., KASPEREK, E., and PICKART, C. M. Metabolism of the polyubiquitin degradation signal: structure, mechanism, and role of isopeptidase T, Biochemistry, 1995, 34, 14535–14546. 34 MELANDRI, F., GRENIER, L., PLAMONDON, L., HUSKEY, W. P., and STEIN, R. L.

35

36

37

38

39

40

41

42

43

Kinetic studies on the inhibition of isopeptidase T by ubiquitin aldehyde, Biochemistry, 1996, 35, 12893–12900. HU, M., LI, P., LI, M., LI, W., YAO, T., WU, J. W., GU, W., COHEN, R. E., and SHI, Y. Crystal structure of a UBP-family deubiquitinating enzyme in isolation and in complex with ubiquitin aldehyde, Cell, 2002, 111, 1041–1054. LI, M., CHEN, D., SHILOH, A., LUO, J., NIKOLAEV, A. Y., QIN, J., and GU, W. Deubiquitination of p53 by HAUSP is an important pathway for p53 stabilization, Nature, 2002, 416, 648–653. WU, K., YAMOAH, K., DOLIOS, G., GAN-ERDENE, T., TAN, P., CHEN, A., LEE, C. G., WEI, N., WILKINSON, K. D., WANG, R., and PAN, Z. Q. DEN1 is a dual function protease capable of processing the C-terminus of Nedd8 deconjugating hyper-neddylated CUL1, J Biol Chem, 2003, 31, 28882–28891. MENDOZA, H. M., SHEN, L. N., BOTTING, C., LEWIS, A., CHEN, J., INK, B., and HAY, R. T. NEDP1, a highly conserved cysteine protease that deNEDDylates Cullins, J Biol Chem, 2003, 278, 25637–25643. LI, S. J. and HOCHSTRASSER, M. A new protease required for cell-cycle progression in yeast, Nature, 1999, 398, 246–251. MOSSESSOVA, E. and LIMA, C. D. Ulp1-SUMO crystal structure and genetic analysis reveal conserved interactions and a regulatory element essential for cell growth in yeast, Mol Cell, 2000, 5, 865–876. BORODOVSKY, A., OVAA, H., KOLLI, N., GAN-ERDENE, T., WILKINSON, K. D., PLOEGH, H. L., and KESSLER, B. M. Chemistry-based functional proteomics reveals novel members of the deubiquitinating enzyme family, Chem Biol, 2002, 9, 1149–1159. BALAKIREV, M. Y., TCHERNIUK, S. O., JAQUINOD, M., and CHROBOCZEK, J. Otubains: a new family of cysteine proteases in the ubiquitin pathway, EMBO Rep, 2003, 4, 517–522. EVANS, P. C., SMITH, T. S., LAI, M. J., WILLIAMS, M. G., BURKE, D. F., HEYNINCK, K., KREIKE, M. M., BEYAERT,

References

44

45

46

47

48

49

50

51

52

53

R., BLUNDELL, T. L., and KILSHAW, P. J. A novel type of deubiquitinating enzyme, J Biol Chem, 2003, 278, 23180–23186. EVANS, P. C., OVAA, H., HAMON, M., KILSHAW, P. J., HAMM, S., BAUER, S., PLOEGH, H. L., and SMITH, T. S. A20, a regulator of inﬂammation and cell survival, has deubiquitinating activity, Biochem J, 2004, Pt 3, 727–734. COPE, G. A., SUH, G. S., ARAVIND, L., SCHWARZ, S. E., ZIPURSKY, S. L., KOONIN, E. V., and DESHAIES, R. J. Role of predicted metalloprotease motif of Jab1/Csn5 in cleavage of Nedd8 from Cul1, Science, 2002, 298, 608–611. VERMA, R., ARAVIND, L., OANIA, R., MCDONALD, W. H., YATES, J. R., 3rd, KOONIN, E. V., and DESHAIES, R. J. Role of Rpn11 metalloprotease in deubiquitination and degradation by the 26S proteasome, Science, 2002, 298, 611–615. YAO, T. and COHEN, R. E. A cryptic protease couples deubiquitination and degradation by the proteasome, Nature, 2002, 419, 403–407. BERNDT, C., BECH-OTSCHIR, D., DUBIEL, W., and SEEGER, M. Ubiquitin system: JAMMing in the name of the lid, Curr Biol, 2002, 12, R815–R817. TRAN, H. J., ALLEN, M. D., LOWE, J., and BYCROFT, M. Structure of the Jab1/MPN domain and its implications for proteasome function, Biochemistry, 2003, 42, 11460–11465. AMBROGGIO, X. I., REES, D. C., and DESHAIES, R. J. JAMM: A Metalloprotease-Like Zinc Site in the Proteasome and Signalosome, PLoS Biol, 2004, 2, E2. MUELLER, T. D., KAMIONKA, M., and FEIGON, J. Speciﬁcity of the interaction between UBA domains and ubiquitin, J Biol Chem, 2004, 12, 11926–11936. KANG, R. S., DANIELS, C. M., FRANCIS, S. A., SHIH, S. C., SALERNO, W. J., HICKE, L., and RADHAKRISHNAN, I. Solution structure of a CUE-ubiquitin complex reveals a conserved mode of ubiquitin binding, Cell, 2003, 113, 621–630. PRAG, G., MISRA, S., JONES, E. A., GHIRLANDO, R., DAVIES, B. A.,

54

55

56

57

58

59

60

61

62

HORAZDOVSKY, B. F., and HURLEY, J. H. Mechanism of ubiquitin recognition by the CUE domain of Vps9p, Cell, 2003, 113, 609–620. FISHER, R. D., WANG, B., ALAM, S. L., HIGGINSON, D. S., ROBINSON, H., SUNDQUIST, W. I., and HILL, C. P. Structure and ubiquitin binding of the ubiquitin interacting motif, J Biol Chem, 2003, 31, 28976–28984. BEAL, R. E., TOSCANO-CANTAFFA, D., YOUNG, P., RECHSTEINER, M., and PICKART, C. M. The hydrophobic effect contributes to polyubiquitin chain recognition, Biochemistry, 1998, 37, 2925–2934. LAM, Y. A., DEMARTINO, G. N., PICKART, C. M., and COHEN, R. E. Speciﬁcity of the ubiquitin isopeptidase in the PA700 regulatory complex of 26 S proteasomes, J Biol Chem, 1997, 272, 28438–28446. PENG, J., SCHWARTZ, D., ELIAS, J. E., THOREEN, C. C., CHENG, D., MARSISCHKY, G., ROELOFS, J., FINLEY, D., and GYGI, S. P. A proteomics approach to understanding protein ubiquitination, Nat Biotechnol, 2003, 21, 921–926. SPENCE, J., SADIS, S., HAAS, A. L., and FINLEY, D. A ubiquitin mutant with speciﬁc defects in DNA repair and multiubiquitination, Mol Cell Biol, 1995, 15, 1265–1273. HOFMANN, R. M. and PICKART, C. M. In vitro assembly and recognition of Lys-63 polyubiquitin chains, J Biol Chem, 2001, 276, 27936–27943. VARADAN, R., WALKER, O., PICKART, C., and FUSHMAN, D. Structural properties of polyubiquitin chains in solution, J Mol Biol, 2002, 324, 637–647. VARADAN, R., ASSFALG, M., HARIRINIA, A., RAASI, S., PICKART, C., and FUSHMAN, D. Solution conformation of Lys63-linked di-ubiquitin chain provides clues to functional diversity of polyubiquitin signaling, J Biol Chem, 2003, 8, 7055–7063. YIN, L., KRANTZ, B., RUSSELL, N. S., DESHPANDE, S., and WILKINSON, K. D. Nonhydrolyzable diubiquitin analogues are inhibitors of ubiquitin conjugation and deconjugation, Biochemistry, 2000, 39, 10001–10010.

25

26 The Deubiquitinating Enzymes

63 JOHNSON, E. S., MA, P. C., OTA, I. M., and VARSHAVSKY, A. A proteolytic pathway that recognizes ubiquitin as a degradation signal, J Biol Chem, 1995, 270, 17442–17456. 64 LI, Z., NA, X., WANG, D., SCHOEN, S. R., MESSING, E. M., and WU, G. Ubiquitination of a novel deubiquitinating enzyme requires direct binding to von Hippel-Lindau tumor suppressor protein, J Biol Chem, 2002, 277, 4656–4662. 65 CHEN, X., ZHANG, B., and FISCHER, J. A. A speciﬁc protein substrate for a deubiquitinating enzyme: Liquid facets is the substrate of Fat facets, Genes Dev, 2002, 16, 289–294. 66 COHEN, M., STUTZ, F., and DARGEMONT, C. Deubiquitination, a new player in Golgi to endoplasmic reticulum retrograde transport, J Biol Chem, 2003, 278, 51989–51992. 67 IDEGUCHI, H., UEDA, A., TANAKA, M., YANG, J., TSUJI, T., OHNO, S., HAGIWARA, E., AOKI, A., and ISHIGATSUBO, Y. Structural and functional characterization of the USP11 deubiquitinating enzyme which interacts with the RanGTP-associated protein RanBPM, Biochem J, 2002, 367, 87–95. 68 KATO, M., MIYAZAWA, K., and KITAMURA, N. A deubiquitinating enzyme UBPY interacts with the Src homology 3 domain of Hrs-binding protein via a novel binding motif PX(V/I) (D/N)RXXKP, J Biol Chem, 2000, 275, 37481–37487. 69 LI, S. J. and HOCHSTRASSER, M. The Ulp1 SUMO isopeptidase: distinct domains required for viability, nuclear envelope localization, and substrate speciﬁcity, J Cell Biol, 2003, 160, 1069–1081. 70 KINNER, A. and KOLLING, R. The yeast deubiquitinating enzyme Ubp16 is anchored to the outer mitochondrial membrane, FEBS Lett, 2003, 549, 135–140. 71 DUPRE, S. and HAGUENAUER-TSAPIS, R. Deubiquitination step in the endocytic pathway of yeast plasma membrane proteins: crucial role of Doa4p ubiquitin

72

73

74

75

76

77

78

79

80

81

isopeptidase, Mol Cell Biol, 2001, 21, 4482–4494. WANG, Y., SATOH, A., WARREN, G., and MEYER, H. H. VCIP135 acts as a deubiquitinating enzyme during p97-p47-mediated reassembly of mitotic Golgi fragments, J Cell Biol, 2004, 164, 973–978. TIAN, Q. B., OKANO, A., NAKAYAMA, K., MIYAZAWA, S., ENDO, S., and SUZUKI, T. A novel ubiquitin-speciﬁc protease, synUSP, is localized at the post-synaptic density and post-synaptic lipid raft, J Neurochem, 2003, 87, 665–675. FISCHER-VIZE, J. A., RUBIN, G. M., and LEHMANN, R. The fat facets gene is required for Drosophila eye and embryo development, Development, 1992, 116, 985–1000. LIU, N., DANSEREAU, D. A., and LASKO, P. Fat facets interacts with vasa in the Drosophila pole plasm and protects it from degradation, Curr Biol, 2003, 13, 1905–1909. ZHU, Y., PLESS, M., INHORN, R., MATHEY-PREVOT, B., and D’ANDREA, A. D. The murine DUB-1 gene is speciﬁcally induced by the betac subunit of interleukin-3 receptor, Mol Cell Biol, 1996, 16, 4808–4817. ZHU, Y., LAMBERT, K., CORLESS, C., COPELAND, N. G., GILBERT, D. J., JENKINS, N. A., and D’ANDREA, A. D. DUB-2 is a member of a novel family of cytokine-inducible deubiquitinating enzymes, J Biol Chem, 1997, 272, 51–57. LOEB, K. R. and HAAS, A. L. The interferon-inducible 15-kDa ubiquitin homolog conjugates to intracellular proteins, J Biol Chem, 1992, 267, 7806–7813. FINLEY, D., OZKAYNAK, E., and VARSHAVSKY, A. The yeast polyubiquitin gene is essential for resistance to high temperatures, starvation, and other stresses, Cell, 1987, 48, 1035–1046. FINLEY, D., BARTEL, B., and VARSHAVSKY, A. The tails of ubiquitin precursors are ribosomal proteins whose fusion to ubiquitin facilitates ribosome biogenesis, Nature, 1989, 338, 394–401. AMERIK, A. Y., LI, S. J., and HOCHSTRASSER, M. Analysis of the

References

82

83

84

85

86

87

88

89

90

deubiquitinating enzymes of the yeast Saccharomyces cerevisiae, Biol Chem, 2000, 381, 981–992. LAMMER, D., MATHIAS, N., LAPLAZA, J. M., JIANG, W., LIU, Y., CALLIS, J., GOEBL, M., and ESTELLE, M. Modiﬁcation of yeast Cdc53p by the ubiquitin-related protein rub1p affects function of the SCFCdc4 complex, Genes Dev, 1998, 12, 914–926. LINGHU, B., CALLIS, J., and GOEBL, M. G. Rub1p processing by Yuh1p is required for wild-type levels of Rub1p conjugation to Cdc53p, Eukaryot Cell, 2002, 1, 491–494. SWAMINATHAN, S., AMERIK, A. Y., and HOCHSTRASSER, M. The Doa4 deubiquitinating enzyme is required for ubiquitin homeostasis in yeast, Mol Biol Cell, 1999, 10, 2583–2594. AMERIK, A. Y., NOWAK, J., SWAMINATHAN, S., and HOCHSTRASSER, M. The Doa4 deubiquitinating enzyme is functionally linked to the vacuolar protein-sorting and endocytic pathways, Mol Biol Cell, 2000, 11, 3365–3380. AMERIK, A., SWAMINATHAN, S., KRANTZ, B. A., WILKINSON, K. D., and HOCHSTRASSER, M. In vivo disassembly of free polyubiquitin chains by yeast Ubp14 modulates rates of protein degradation by the proteasome, Embo J, 1997, 16, 4826–4838. KAWAKAMI, T., CHIBA, T., SUZUKI, T., IWAI, K., YAMANAKA, K., MINATO, N., SUZUKI, H., SHIMBARA, N., HIDAKA, Y., OSAKA, F., OMATA, M., and TANAKA, K. NEDD8 recruits E2-ubiquitin to SCF E3 ligase, Embo J, 2001, 20, 4003–4012. DANIEL, J. A., TOROK, M. S., SUN, Z. W., SCHIELTZ, D., ALLIS, C. D., YATES, J. R., 3rd, and GRANT, P. A. Deubiquitination of histone H2B by a yeast acetyltransferase complex regulates transcription, J Biol Chem, 2004, 279, 1867–1871. HOEGE, C., PFANDER, B., MOLDOVAN, G. L., PYROWOLAKIS, G., and JENTSCH, S. RAD6-dependent DNA repair is linked to modiﬁcation of PCNA by ubiquitin and SUMO, Nature, 2002, 419, 135–141. GROLL, M., DITZEL, L., LOWE, J., STOCK, D., BOCHTLER, M., BARTUNIK, H. D., and HUBER, R. Structure of 20S proteasome

91

92

93

94

95

96

97

98

99

100

from yeast at 2.4 A resolution, Nature, 1997, 386, 463–471. LAM, Y. A., XU, W., DEMARTINO, G. N., and COHEN, R. E. Editing of ubiquitin conjugates by an isopeptidase in the 26S proteasome, Nature, 1997, 385, 737–740. GUTERMAN, A. and GLICKMAN, M. H. Complementary roles for Rpn11 and Ubp6 in deubiquitination and proteolysis by the proteasome, J Biol Chem, 2004, 279, 1729–1738. EYTAN, E., ARMON, T., HELLER, H., BECK, S., and HERSHKO, A. Ubiquitin C-terminal hydrolase activity associated with the 26 S protease complex, J Biol Chem, 1993, 268, 4668–4674. LEGGETT, D. S., HANNA, J., BORODOVSKY, A., CROSAS, B., SCHMIDT, M., BAKER, R. T., WALZ, T., PLOEGH, H., and FINLEY, D. Multiple associated proteins regulate proteasome structure and function, Mol Cell, 2002, 10, 495–507. CHERNOVA, T. A., ALLEN, K. D., WESOLOSKI, L. M., SHANKS, J. R., CHERNOFF, Y. O., and WILKINSON, K. D. Pleiotropic effects of Ubp6 loss on drug sensitivities and yeast prion are due to depletion of the free ubiquitin pool, J Biol Chem, 2003, 278, 52102–52115. BAKER, R. T., TOBIAS, J. W., and VARSHAVSKY, A. Ubiquitin-speciﬁc proteases of Saccharomyces cerevisiae. Cloning of UBP2 and UBP3, and functional analysis of the UBP gene family, J Biol Chem, 1992, 267, 23364–23375. TOBIAS, J. W. and VARSHAVSKY, A. Cloning and functional analysis of the ubiquitin-speciﬁc protease gene UBP1 of Saccharomyces cerevisiae, J Biol Chem, 1991, 266, 12021–12028. KARIN, M. and BEN-NERIAH, Y. Phosphorylation meets ubiquitination: the control of NF-[kappa]B activity, Annu Rev Immunol, 2000, 18, 621–663. ZHOU, H., WERTZ, I., O’ROURKE, K., ULTSCH, M., SESHAGIRI, S., EBY, M., XIAO, W., and DIXIT, V. M. Bcl10 activates the NF-kappaB pathway through ubiquitination of NEMO, Nature, 2004, 427, 167–171. YARON, A., GONEN, H., ALKALAY, I., HATZUBAI, A., JUNG, S., BEYTH, S.,

27

28 The Deubiquitinating Enzymes

101

102

103

104

105

106

107

MERCURIO, F., MANNING, A. M., CIECHANOVER, A., and BEN-NERIAH, Y. Inhibition of NF-kappa-B cellular function via speciﬁc targeting of the I-kappa-B-ubiquitin ligase, Embo J, 1997, 16, 6486–6494. WANG, C., DENG, L., HONG, M., AKKARAJU, G. R., INOUE, J., and CHEN, Z. J. TAK1 is a ubiquitin-dependent kinase of MKK and IKK, Nature, 2001, 412, 346–351. TANG, E. D., WANG, C. Y., XIONG, Y., and GUAN, K. L. A role for NF-kappaB essential modiﬁer/IkappaB kinase-gamma (NEMO/IKKgamma) ubiquitination in the activation of the IkappaB kinase complex by tumor necrosis factor-alpha, J Biol Chem, 2003, 278, 37297–37305. TROMPOUKI, E., HATZIVASSILIOU, E., TSICHRITZIS, T., FARMER, H., ASHWORTH, A., and MOSIALOS, G. CYLD is a deubiquitinating enzyme that negatively regulates NF-kappaB activation by TNFR family members, Nature, 2003, 424, 793–796. BRUMMELKAMP, T. R., NIJMAN, S. M., DIRAC, A. M., and BERNARDS, R. Loss of the cylindromatosis tumour suppressor inhibits apoptosis by activating NF-kappaB, Nature, 2003, 424, 797– 801. CIECHANOVER, A. and BRUNDIN, P. The ubiquitin proteasome system in neurodegenerative diseases: sometimes the chicken, sometimes the egg, Neuron, 2003, 40, 427–446. LIU, Y., FALLON, L., LASHUEL, H. A., LIU, Z., and LANSBURY, P. T., Jr. The UCH-L1 gene encodes two opposing enzymatic activities that affect alpha-synuclein degradation and Parkinson’s disease susceptibility, Cell, 2002, 111, 209–218. LEROY, E., BOYER, R., AUBURGER, G., LEUBE, B., ULM, G., MEZEY, E., HARTA, G., BROWNSTEIN, M. J., JONNALAGADA, S., CHERNOVA, T., DEHEJIA, A., LAVEDAN, C., GASSER, T., STEINBACH, P. J., WILKINSON, K. D., and POLYMEROPOULOS, M. H. The ubiquitin pathway in Parkinson’s disease, Nature, 1998, 395, 451–452.

108 LOWE, J., MCDERMOTT, H., LANDON, M., MAYER, R. J., and WILKINSON, K. D. Ubiquitin carboxyl-terminal hydrolase (PGP 9.5) is selectively present in ubiquitinated inclusion bodies characteristic of human neurodegenerative diseases, J Pathol, 1990, 161, 153–160. 109 WILSON, S. M., BHATTACHARYYA, B., RACHEL, R. A., COPPOLA, V., TESSAROLLO, L., HOUSEHOLDER, D. B., FLETCHER, C. F., MILLER, R. J., COPELAND, N. G., and JENKINS, N. A. Synaptic defects in ataxia mice result from a mutation in Usp14, encoding a ubiquitin-speciﬁc protease, Nat Genet 2002, 32, 420–425. 110 GARCIA-MATA, R., GAO, Y. S., and SZTUL, E. Hassles with taking out the garbage: aggravating aggresomes, Trafﬁc, 2002, 3, 388–396. 111 KAWAGUCHI, Y., KOVACS, J. J., MCLAURIN, A., VANCE, J. M., ITO, A., and YAO, T. P. The deacetylase HDAC6 regulates aggresome formation and cell viability in response to misfolded protein stress, Cell, 2003, 115, 727–738. 112 SAIGOH, K., WANG, Y. L., SUH, J. G., YAMANISHI, T., SAKAI, Y., KIYOSAWA, H., HARADA, T., ICHIHARA, N., WAKANA, S., KIKUCHI, T., and WADA, K. Intragenic deletion in the gene encoding ubiquitin carboxy-terminal hydrolase in gad mice, Nat Genet 1999, 23, 47–51. 113 KURIHARA, L. J., SEMENOVA, E., LEVORSE, J. M., and TILGHMAN, S. M. Expression and functional analysis of Uch-L3 during mouse development, Mol Cell Biol, 2000, 20, 2498–2504. 114 KURIHARA, L. J., KIKUCHI, T., WADA, K., and TILGHMAN, S. M. Loss of Uch-L1 and Uch-L3 leads to neuro-degeneration, posterior paralysis and dysphagia, Hum Mol Genet, 2001, 10, 1963–1970. 115 BORODOVSKY, A., KESSLER, B. M., CASAGRANDE, R., OVERKLEEFT, H. S., WILKINSON, K. D., and PLOEGH, H. L. A novel active site-directed probe speciﬁc for deubiquitylating enzymes reveals proteasome association of USP14, Embo J, 2001, 20, 5187–5196. 116 HEMELAAR, J., LELYVELD, V. S., KESSLER, B. M., and PLOEGH, H. L. A single protease,

References

117

118

119

120

121

122

123

124

Apg4B, is speciﬁc for the autophagy-related ubiquitin-like proteins GATE-16, MAP1-LC3, GABARAP, and Apg8L, J Biol Chem, 2003, 278, 51841–51850. RUSSELL, N. S. and WILKINSON, K. D. Identiﬁcation of a novel 29-linked polyubiquitin binding protein, ufd3, using polyubiquitin chain analogues, Biochemistry, 2004, 43, 4844–4854. MOAZED, D. and JOHNSON, D. A deubiquitinating enzyme interacts with SIR4 and regulates silencing in S. cerevisiae, Cell, 1996, 86, 667–677. NAVIGLIO, S., MATTECUCCI, C., MATOSKOVA, B., NAGASE, T., NOMURA, N., DI FIORE, P. P., and DRAETTA, G. F. UBPY: a growth-regulated human ubiquitin isopeptidase, Embo J, 1998, 17, 3241–3250. VALERO, R., BAYES, M., FRANCISCA SANCHEZ-FONT, M., GONZALEZ-ANGULO, O., GONZALEZ-DUARTE, R., and MARFANY, G. Characterization of alternatively spliced products and tissue-speciﬁc isoforms of USP28 and USP25, Genome Biol, 2001, 2, RESEARCH0043. LEE, E. G., BOONE, D. L., CHAI, S., LIBBY, S. L., CHIEN, M., LODOLCE, J. P., and MA, A. Failure to regulate TNF-induced NF-kappaB and cell death responses in A20-deﬁcient mice, Science, 2000, 289, 2350–2354. SOARES, L., SEROOGY, C., SKRENTA, H., ANANDASABAPATHY, N., LOVELACE, P., CHUNG, C. D., ENGLEMAN, E., and FATHMAN, C. G. Two isoforms of otubain 1 regulate T cell anergy via GRAIL, Nat Immunol 2004, 5, 45–54. LI, S. J. and HOCHSTRASSER, M. The yeast ULP2 (SMT4) gene encodes a novel protease speciﬁc for the ubiquitin-like Smt3 protein, Mol Cell Biol, 2000, 20, 2367–2377. KIM, K. I., BAEK, S. H., JEON, Y. J., NISHIMORI, S., SUZUKI, T., UCHIDA, S., SHIMBARA, N., SAITOH, H., TANAKA, K., and CHUNG, C. H. A new SUMO-1-speciﬁc protease, SUSP1, that is highly expressed in reproductive

125

126

127

128

129

130

131

132

133

organs, J Biol Chem, 2000, 275, 14102–14106. BURNETT, B., LI, F., and PITTMAN, R. N. The polyglutamine neuro-degenerative protein ataxin-3 binds polyubiquitylated proteins and has ubiquitin protease activity, Hum Mol Genet, 2003, 12, 3195–3205. WADA, H., KITO, K., CASKEY, L. S., YEH, E. T., and KAMITANI, T. Cleavage of the C-terminus of NEDD8 by UCH-L3, Biochem Biophys Res Commun, 1998, 251, 688–692. CABALLERO, O. L., RESTO, V., PATTURAJAN, M., MEERZAMAN, D., GUO, M. Z., ENGLES, J., YOCHEM, R., RATOVITSKI, E., SIDRANSKY, D., and JEN, J. Interaction and colocalization of PGP9.5 with JAB1 and p27(Kip1), Oncogene, 2002, 21, 3003–3010. FIORANI, P., REID, R. J., SCHEPIS, A., JACQUIAU, H. R., GUO, H., THIMMAIAH, P., BENEDETTI, P., and BJORNSTI, M. A. The deubiquitinating enzyme Doa4p protects cells from DNA topoisomerase I poisons, J Biol Chem, 2004, 20, 21271–21281. BREW, C. T. and HUFFAKER, T. C. The yeast ubiquitin protease, Ubp3p, promotes protein stability, Genetics, 2002, 162, 1079–1089. VERMA, R., CHEN, S., FELDMAN, R., SCHIELTZ, D., YATES, J., DOHMEN, J., and DESHAIES, R. J. Proteasomal proteomics: identiﬁcation of nucleotide-sensitive proteasome-interacting proteins by mass spectrometric analysis of afﬁnity-puriﬁed proteasomes, Mol Biol Cell, 2000, 11, 3425–3439. HADARI, T., WARMS, J. V., ROSE, I. A., and HERSHKO, A. A ubiquitin C-terminal isopeptidase that acts on polyubiquitin chains. Role in protein degradation, J Biol Chem, 1992, 267, 719–727. HONG, S., KIM, S. J., KA, S., CHOI, I., and KANG, S. USP7, a ubiquitin-speciﬁc protease, interacts with ataxin-1, the SCA1 gene product, Mol Cell Neurosci, 2002, 20, 298–306. GNESUTTA, N., CERIANI, M., INNOCENTI, M., MAURI, I., ZIPPEL, R., STURANI, E., BORGONOVO, B., BERRUTI, G., and

29

30 The Deubiquitinating Enzymes

MARTEGANI, E. Cloning and characterization of mouse UBPy, a deubiquitinating enzyme that interacts with the ras guanine nucleotide exchange factor CDC25(Mm)/ Ras-GRF1, J Biol Chem, 2001, 276, 39448–39454.

134 WANG, G., SAWAI, N., KOTLIAROVA, S., KANAZAWA, I., and NUKINA, N. Ataxin-3, the MJD1 gene product, interacts with the two human homologs of yeast DNA repair protein RAD23, HHR23A and HHR23B, Hum Mol Genet, 2000, 9, 1795–1803.

1

The 26S Proteasome Martin Rechsteiner University of Utah Medical School, Salt Lake City, USA

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8 The 26S proteasome is a large ATP-dependent protease composed of more than 30 different polypeptide chains. Like the ribosome, the 26S proteasome is assembled from two “subunits”, the 19S regulatory complex and the 20S proteasome. The 19S regulatory complex confers the ability to recognize and unfold protein substrates, and the 20S proteasome provides the proteolytic activities needed to degrade the substrates. The 26S proteasome is the only enzyme known to degrade ubiquitylated proteins, and it also degrades intracellular proteins that have not been marked by ubiquitin. The 26S proteasome is located in the nucleus and cytosol of eukaryotic cells, where the enzyme is responsible for the selective degradation of a vast number of important cellular proteins. Because rapid proteolysis is a pervasive regulatory mechanism, the 26S proteasome is essential for the proper functioning of many physiological processes.

1 Introduction

It has become apparent since the mid-1990s that the ubiquitin-proteasome system (UPS) plays a major regulatory role in eukaryotic cells. The UPS helps to control such important physiological processes as the cell cycle [1, 2], circadian rhythms [3], axon guidance [4], synapse formation [5] and transcription [6–8], to name just a few. In view of the growing family of ubiquitin-like proteins [9, 10], it is possible that covalent attachment of ubiquitin and its relatives will even surpass phosphorylation as a regulatory mechanism. Although ubiquitin serves non-proteolytic roles, such as histone modification [11, 12] the activation of cell signaling components [13], endocytosis [14], or viral budding [15], the protein’s principal function appears to be targeting proteins for destruction [16]. To do this, the C-terminus of ubiquitin is activated by an ATP-consuming enzyme (E1) and transferred to one of several

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 The 26S Proteasome

Fig. 1 Schematic representation of the Ubiquitin–Proteasome pathway. Ubiquitin molecules are activated by an E1 enzyme (shown green at 1/3 scale) in an ATPdependent reaction, transferred to a cysteine residue (yellow) on an E2 or Ub carrier protein and subsequently attached to

amino groups (NH2 ) on a substrate protein (lysozyme shown in purple) by an E3 or ubiquitin ligase, (the multicolored SCF complex). Note that chains of Ub are generated on the substrate, and these are recognized by the 26S proteasome depicted in the upper right at 1/20 scale.

dozen or more small carrier proteins (E2s) in the form of a reactive thiol ester. The E2s collaborate with members of several large families of ubiquitin ligases or E3s, and ubiquitin is transferred once again to lysine amino groups on the proteolytic substrates (S) and to itself, thereby generating chains of ubiquitin. The substrate bearing the ubiquitin chains is subsequently recognized and degraded by the 26S proteasome, a large, complex ATP-dependent protease (see Figure 1). Eukaryotic genomes contain information for more than 20 E2s and hundreds of E3s. In contrast to the wealth of components devoted to marking protein substrates for destruction, only one enzyme, the 26S proteasome, has been found to degrade ubiquitylated proteins. However, there is complexity here as well, since the 26S proteasome is an assemblage of at least 30 different subunits. Moreover, there is

2 The 20S Proteasome

a growing list of proteins that act as proteasome activators, adapters, or accessory factors. In this article I focus on basic biochemical and physiological properties of the 26S proteasome, drawing occasionally from findings on structurally similar prokaryotic, ATP-dependent proteases [17].

2 The 20S Proteasome 2.1 Structure

We know the molecular anatomy of archaebacterial, yeast and bovine proteasomes in great detail since high-resolution crystal structures have been determined for all three enzymes [18–20]. The archaebacterial proteasome is composed of two kinds of subunits, called α and β. Each subunit forms heptameric rings that assemble into the 20S proteasome by stacking four deep on top of one another to form a “hollow” cylinder. Catalytically inactive α rings form the ends of the cylinder while proteolytic β subunits occupy the two central rings. The quaternary structure of the 20S proteasome can therefore be described as α7β7β7α7. The active sites of the β-subunits face a large central chamber about the size of serum albumin. The αrings seal off the central proteolytic chamber and two smaller antechambers from the external solvent. Archaebacterial proteasomes, with their fourteen identical β subunits, preferentially hydrolyze peptide bonds following hydrophobic amino acids and are therefore said to have chymotrypsin-like activity [21]. Eukaryotic proteasomes maintain the overall structure of the archaebacterial enzyme, but they exhibit a more complicated subunit composition. There are seven different α-subunits and at least seven distinct β-subunits arranged in a precise order within their respective rings. Although current evidence indicates that only three of its seven β-subunits are catalytically active, the eukaryotic proteasome cleaves a wider range of peptide bonds, containing, as it does, two copies each of trypsin-like, chymotrypsin-like and post-glutamyl-hydrolyzing subunits. For this reason it is capable of cleaving almost any peptide bond, having difficulty only with proline–X, glycine–X and to a lesser extent with glutamine–X bonds [22]. 2.2 Enzyme Mechanism and Proteasome Inhibitors

Whereas standard proteases use serine, cysteine, aspartate, or metals to cleave pep-tide bonds, the proteasome employs an unusual catalytic mechanism. Nterminal threonine residues are generated by self-removal of short peptide extensions from the active β-subunits and act as nucleophiles during peptide-bond hydrolysis [23]. Given its unusual catalytic mechanism, it is not surprising that there are highly specific inhibitors of the proteasome. The fungal metabolite lactacystin

3

4 The 26S Proteasome

and the bacterial product epoxomicin covalently modify the active-site threonines and inhibit the enzyme [24, 25]. Other inhibitors include vinylsulfones [26] and various peptide aldehydes, which are generally less specific. A peptide boronate inhibitor of the proteasome, Velcade, has been approved for the treatment of multiple myeloma [27]. 2.3 Immunoproteasomes

Interferon γ (IFNγ ) is an immune cytokine that increases expression of a number of cellular components involved in Class I antigen presentation [28]. Among the IFNγ -inducible components are three catalytically active β-subunits of the proteasome, called LMP2, LMP7 and MECL1 [29]. Each replaces its corresponding constitutive subunit resulting in altered peptide-bond cleavage preferences of 20S immunoproteasomes. For example, immunoproteasomes exhibit much reduced cleavage after acidic residues and enhanced hydrolysis of peptide bonds following branch-chain amino acids such as isoleucine or valine. Class I molecules preferentially bind peptides with hydrophobic or positive C-termini, and proteasomes generate the vast majority of Class I peptides [28, 30]. Hence, the observed β-subunit exchanges are well suited for producing peptides able to bind Class I molecules.

3 The 26S Proteasome 3.1 The Ubiquitin–Proteasome System

Bacteria express as many as five ATP-dependent proteases, all of which contain nucleotide-binding domains that belong to the AAA+ family of ATPases [31]. By contrast, the 26S proteasome is the only ATP-dependent protease discovered so far in the nuclear and cytosolic compartments of eukaryotic cells. Because the 20S proteasome’s internal cavities are inaccessible to intact proteins, openings must be generated in the enzyme’s outer surface for proteolysis to occur. A number of protein complexes have been found to bind the proteasome and stimulate peptide hydrolysis (see Figure 2). The most important of the proteasome-associated components is the 19S regulatory complex (RC) for it is a major part of the 26S ATP-dependent enzyme that degrades ubiquitin-tagged proteins in eukaryotic cells. In Figure 2 the 20S proteasome is shown binding only one 19S RC although doublycapped 26S proteasomes also exist (see Figure 3 below). The 20S proteasome also binds activators such as PA28 or PA200. Each of these activators can be present in 26S proteasome complexes forming what are called hybrid proteasomes. Finally a protein called Ecm29p has been found associated with the 26S proteasome. Ecm29p is thought to act as an adapter coupling the 26S enzyme to secretory vesicles.

3 The 26S Proteasome

Fig. 2 Interaction of the 20S proteasome with other cellular components.

Fig. 3 Electron-microscopic reconstructions of the 26S proteasome. Three images of a doubly-capped 26S proteasome are presented to illustrate the positions of the lid and base subcomplexes of the 19S RC and to identify the most probable location of the RC ATPases.

5

6 The 26S Proteasome

3.2 Ultrastructure of the 26S Proteasome and Regulatory Complex

Electron micrographs of purified 26S proteasomes by Baumeister and colleagues [32] reveal a dumbbell-shaped particle approximately 40 nm in length in which the central 20S proteasome cylinder is capped at one or both ends by asymmetric RCs looking much like Chinese dragonheads (Figure 3). In doubly capped 26S proteasomes the regulatory complexes face in opposite directions, indicating that contact between the proteasome’s α-rings and the RC is highly specific. However, the contacts may not be especially tight since image analysis of Drosophila 26S proteasomes suggests movement of the RCs relative to the 20S proteasome [33]. Electron microscopy (EM) images of the 26S proteasome from several organisms appear similar, indicating that the overall architecture of the enzyme has been conserved in evolution. This conclusion is also supported by sequence conservation among RC subunits (see below). A yeast mutant lacking the RC subunit Rpn10 contains a salt-labile RC that dissociates into two subcomplexes called the lid and the base [34]. The base contains nine RC subunits, which include six ATPases described below, the two largest RC subunits S1 and S2, and S5a; the lid contains the remaining RC subunits. Thus the RC is composed of two sub-complexes separated on one side by a cavity, i.e. the dragon’s mouth (see Figure 3). Ultrastructural studies have also been performed on the lid and on a related protein complex called the COP9 signalosome [35]. Both particles lack obvious symmetry. Some particles exhibit a negative stainfilled, central groove; other classes of particles exhibit seven or eight lobes in a disclike arrangement. Since both the lid and the COP9 signalosome are composed of eight subunits, the lobes may represent individual subunits. 3.3 The 19S Regulatory Complex

The regulatory complex is also called the 19S cap, PA700, and the µ particle. As its most commonly used name suggests, the 19S RC is roughly the same size as the 20S proteasome. In fact it is a more complicated protein assembly containing 17 different subunits ranging in size from 25 kDa to about 110 kDa. In animal cells the subunits are designated S1 through S15. Homologs for each of these subunits are present in budding yeast where an alternate nomenclature has been adopted (see Table 1). Sequences for the 17 RC subunits permit their classification into a group of 6 ATPases and another group containing the 11 non-ATPases. 3.4 ATPases of the RC

The six ATPases belong to the rather large family of AAA ATPases (for ATPases Associated with a variety of cellular Activities) whose eukaryotic members include the motor protein dynein, the membrane fusion factor NSF, and the chaperone VCP/Cdc48 and whose prokaryotic members include five ATP-dependent pro-

3 The 26S Proteasome Table 1 Subunits of the 19S regulatory complex

Mammalian nomenclature Yeast nomenclature Function

Motifs

S1 S2 S3 p55 S4 S5a S5b S6 S6 S7 S8 S9 S10a S10b S11 S12 S13 S14

Leu-rich repeats, KEKE Leu-rich repeats, KEKE PCI PCI AAA nucleotidase UIM, KEKE

Rpn2 Rpn1 Rpn3 Rpn5 Rpt2 Rpn10 none Rpt3 Rpt5 Rpt1 Rpt6 Rpn6 Rpn7 Rpt4 Rpn9 Rpn8 Rpn11 Rpn12

Ub/UbL binding Ub/UbL binding ? ? ATPase polyubiquitin binding ? ATPase ATPase ATPase ATPase ? ? ATPase ? ? Isopeptidase ?

AAA nucleotidase AAA nucleotidase AAA nucleotidase AAA nucleotidase PCI PCI, KEKE AAA nucleotidase PCI MPN, KEKE MPN PCI

teases [31]. The six ATPases, denoted S4, S6, S6 , S7, S8, and S10b in mammals, are about 400 amino acids in length and homologous to one another. Based on their sequences, one can distinguish three major regions: (1) A central nucleotidebinding domain of about 200 amino acids, which is roughly 60% identical among members of the RC subfamily; (2) the C-terminal region, approximately 100 amino acids in length and with a lesser, though significant, degree of conservation (40%); and (3) a highly divergent N-terminal region (<20% identity) around 120 amino acids in length; this region contains heptad repeats characteristic of coiled-coil proteins. Despite sequence differences among RC ATPases within an organism, each ATPase has been conserved during evolution with specific subunits being almost 75% identical between yeast and humans. The high degree of conservation encompasses the entire sequence, making it likely that even the divergent N-terminal regions play an important role in RC function. Conceivably they are used to select substrates for degradation by the 26S proteasome [36]. Alternatively the variable N-terminal regions in the RC ATPases may promote assembly of the RC by the specific placement of the ATPase subunits within the complex. In this regard, the six ATPases associate with one another in highly specific pairs: S4 binds S7, S6 binds S8, and S6 binds to S10b. Moreover, the N-terminal regions of RC ATPases are required for partner-specific binding [37]. Staining patterns of two-dimensional gels show the six RC ATPases to be present at comparable levels, and affinity capture of yeast 26S proteasomes indicate the presence of one copy of each in the regulatory complex. Mutational analysis in yeast demonstrates that the ATPases are not functionally redundant since mutation of yeast S4 has a particularly profound effect on peptide hydrolysis [38]. It is probable that the ATPases form a hexameric ring like other members of the AAA

7

8 The 26S Proteasome

family of ATPases such as NSF or VCP/Cdc48. But this assumption has not been experimentally verified, and pentameric or heptameric arrangements of AAA ATPase complexes have been reported [39, 40]. Finally it is quite likely that the ATP-ases directly bind the α-ring of the 20S proteasome (see Figure 3). Evidence favoring this arrangement comes from chemical cross-linking experiments [41] and the presence of the ATPases in the base subcomplex of the yeast 26S proteasome [34]. 3.5 The non-ATPase Subunits

Whereas the six RC ATPases are homologous and relatively uniform in size, the non-ATPases are heterogeneous in size and sequence. Nonetheless, they can be grouped on the basis of their location, on the presence or absence of certain sequence motifs, and on their affinity for ubiquitin chains or ubiquitin-like proteins. Eight RC subunits are found in the lid subcomplex, each being homologous to a subunit in a separate protein complex called the COP9 Signalosome [42–44]. One of the lid subunits, S13, is a metalloisopeptidase that removes ubiquitin chains from the tagged substrate prior to its translocation into the proteasome for degradation. Six of the eight lid subunits contain PCI domains, stretches of about 200 residues so named from their occurrence in Proteasome, Cop9 signalosome, and the eukaryotic Initiation factor 3 subunits. The PCI domains are thought to mediate subunit-subunit interactions. Two lid subunits contain 140 amino acid-long MPN domains (Mpr1p and Pad1p N-terminal regions) with one of these subunits being the S13 isopeptidase. Although several models have been proposed [45–48], the arrangement of subunits within the lid subcomplex is not known. Functionally, only S13 stands out because of its isopeptidase activity. The presence of S13 in the lid explains why the lid is necessary for degradation of ubiquitylated proteins even though the RC base complex supports the ATP-dependent degradation of some small non-ubiquitylated proteins [34]. Interestingly the COP9 signalosome also exhibits isopeptidase activity that removes the ubiquitin-like protein, NEDD8, from certain ubiquitin ligases [49]. Biochemical activity has not been assigned to any of the remaining seven lid subunits, although two lid subunits, S3 and S9, are critical for the degradation of specific substrates [50, 51]. In addition to the six ATPases the base sub-complex contains the two largest RC subunits (S1, S2) and a smaller subunit called S5a [52]. Besides their common location, these three subunits share the property of binding polyubiquitin chains or ubiquitin-like (UbL) domains. S5a binds polyubiquitin chains even after it has been transferred from SDS-PAGE gels and displays many features that match polyubiquitin recognition by the 26S proteasome [52]. However, S5a cannot be the only ubiquitin-recognition component in the 26S proteasome because deletion of the gene encoding yeast S5a (Rpn10) has only a modest impact on proteolysis [53]. This strongly suggests that there are other ubiquitin-recognition components in the RC, with S1 and S2 being prime candidates. S1 and S2 display significant homology to each other, and both can be modeled as α-helical toroids [54]. They have been shown to bind the UbL domains of RAD23 and Dsk2, adapter proteins

3 The 26S Proteasome

that target ubiquitylated proteins to the 26S proteasome [55, 56]. It has also been found that the S6 ATPase can be cross-linked to ubiquitin [57]. Currently then it appears that the RC contains three or possibly four subunits able to recognize ubiquitin or UbL proteins. As discussed below there are other ways in which the RC can select proteins for destruction. 3.6 Biochemical Properties of the Regulatory Complex 3.6.1 Nucleotide Hydrolysis Both the 26S proteasome and the RC hydrolyze all four nucleotide triphosphates, with ATP and CTP preferred over GTP and UTP [58]. Although ATP hydrolysis is required for conjugate degradation, the two processes are not strictly coupled. Complete inhibition of the peptidase activity of the 26S proteasome by calpain inhibitor I has little effect on the ATPase activity of the enzyme. The nucleotidase activities of the RC and the 26S proteasome closely resemble those of E. coli Lon protease, which is composed of identical subunits that possess both proteolytic and nucleotidase activities in the same polypeptide chain. Like the regulatory complex and 26S proteasome, Lon hydrolyzes all four ribonucleotide triphosphates, but not ADP or AMP [18]. 3.6.2 Chaperone-like Activity AAA nucleotidases share the common property of altering the conformation or association state of proteins, so it is not surprising that the RC has been shown to prevent aggregation of several denatured proteins including citrate synthase and ribonuclease A [59–61]. The chaperone activity of the RC may explain why the RC plays a role in transcription apparently in the absence of an attached 20S proteasome [62]. 3.6.3 Proteasome Activation The 20S proteasome is a latent protease owing to the barrier imposed by the αsubunit rings on peptide entry. Consequently, a readily measured activity of the RC is activation of fluorogenic peptide hydrolysis by the 20S proteasome. The extent of activation is generally found to be in the range 3- to 20-fold [63]. Activation is relatively uniform for all three proteasome catalytic subunits and presumably reflects opening by the attached RC of a channel leading to the proteasome’s central chamber. 3.6.4 Ubiquitin Isopeptide Hydrolysis The channel through the proteasome α-ring into the central chamber measures 1.3 nm in diameter, a size too small to permit passage of a folded protein, even one as small as ubiquitin. This consideration, coupled with the fact that ubiquitin is recycled intact upon substrate degadation, requires an enzyme to remove the polyubiquitin chain prior to or concomitant with proteolysis. Several isopeptidases that remove ubiquitin from substrates have been found associated with the 26S

9

10 The 26S Proteasome

proteasome [64–66]. Of these the ATP-stimulated metalloisopeptidase S13 is an integral component of the enzyme. 3.6.5 Substrate Recognition It is clear that the RC plays a predominant role in selecting proteins for degradation. This important topic is covered below in the context of substrate recognition by both 20S and 26S proteasomes.

4 Substrate Recognition by Proteasomes 4.1 Degradation Signals (Degrons)

The discovery that proteins possess built-in signals targeting them to specific locations within cells was a major success of twentieth-century cell biology [67]. Selective proteolysis can be considered targeting out of existence, and a number of short peptide motifs have been discovered to confer rapid destruction on proteins that bear them. These motifs include PEST sequences [68], the N-terminal amino acid [69], and destruction and KEN boxes [70]. These motifs are recognized by one or more ubiquitin ligases that mark the substrate protein by addition of a polyubiquitin chain. However some proteins are degraded by the 26S proteasome without prior marking by ubiquitin [71]. Denatured proteins are also selectively degraded by both 26S and 20S proteasomes [72]. It is not clear what features of denatured proteins are recognized by proteasomes or by components of the ubiquitin proteolytic system. 4.2 Ubiquitin-dependent Recognition of Substrates

Most well characterized substrates of the 26S proteasome are ubiquitylated proteins so our discussion starts with them. Ubiquitin contains seven lysine residues, and proteomic studies in yeast indicate that chains (or dimers) can be formed using any of them [73]. In higher eukaryotes polyubiquitin chains formed via Lys6, Lys27, Lys29, Lys48, and Lys63 have been observed. Lys6 chains are formed by BRCA/BARD heterodimers where they presumably play a role in DNA repair [74]. Ubiquitin monomers linked to each other through Lys63 are involved in endocytosis and DNA repair, but not apparently in targeting proteins to the 26S proteasome [75, 76]. Lys27 chains have been found on the co-chaperone BAG1, and they target degradation of misfolded proteins bound by the Hsp70 chaperone to the 26S proteasome [77]. Lys29 and Lys48 chains form directly on proteolytic substrates and target them for destruction [78, 79]. Efficient proteolysis of ubiquitylated proteins by the 26S proteasome requires a chain containing at least four ubiquitin monomers [80]. This matches well the ubiquitin-binding characteristics of the RC subunit S5a. It too selectively binds

4 Substrate Recognition by Proteasomes

ubiquitin polymers composed of four or more ubiquitin moieties and exhibits increased affinity for longer chains [52]. S5a molecules from a number of higher eukaryotes contain two independent polyubiquitin-binding sites; each is approximately 30 residues long and characterized by five hydrophobic residues that consist of alternating large and small hydrophobic residues, e.g. Leu-Ala-Leu-Ala-Leu [81]. Similar motifs have been found in other proteins of the ubiquitin system and are now called UIMs, (Ubiquitin Interacting Motifs) [82]. Two recent NMR studies have demonstrated direct interaction between UIMs and the hydrophobic patch on ubiquitin [83–85]. Whereas S5a provides for direct recognition of polyubiquitylated substrates, a second mechanism involves adapter proteins possessing both a UbL domain and one or more UbA (ubiquitin associated) domains [86]. UbA domains are polyubiquitin-binding domains found in several presumed adapter proteins of the ubiquitin system. The adapter proteins include RAD23 and Dsk2 in yeast and recruit substrates to the 26S proteasome. The UbL domains of these proteins bind to 26S proteasome subunits while their UbA domains bind substrate-tethered Ub chains. In yeast the RC subunits S1 and S2 serve as UbL-binding components [55, 87]; in mammals S5a serves this purpose [88–90]. The co-chaperone BAG1 illustrates a third way in which polyubiquitin can target substrate proteins to the 26S proteasome. In this case the substrate is not polyubiquitylated; rather it is bound to the chaperone Hsp70. A polyubiquitin chain, linked through Lys27, is attached to the Hsp70-associated co-chaperone BAG1 [77]. Apparently the Lys27 chain promotes association of the chaperone–substrate complex with the 26S proteasome, after which the substrate is degraded while BAG1, Hsp70, and ubiquitin are recycled. Direct interaction between E3 ubiquitin ligases and RC subunits can also deliver ubiquitylated substrates to the protease. The yeast E3 ligase called UFD4 binds RC ATPases [91] and UFD4-mediated delivery of substrates bypasses the requirement for S5a. In what appears to be a similar delivery system, the mammalian E3 Parkin uses a UbL domain to bind the 26S proteasome [89], and the E3 component pVHL binds a 26S proteasome ATPase [92]. Mutational analyses in yeast have shown that whereas deletion of either S5a or Rad23 has a mild impact on proteolysis, loss of both proteins produces a severe phenotype [93]; furthermore, yeasts lacking S5a, RAD23 and Dsk2 are not viable, indicating that direct delivery by E3 ligases cannot compensate for the absence of all three proteins.

4.3 Substrate Selection Independent of Ubiquitin

The 26S proteasome also degrades non-ubiquitylated proteins [71]. The short-lived enzyme ornithine decarboxylase (ODC) and the cell-cycle regulator p21Cip provide well documented examples of ubiquitin-independent proteolysis by the 26S enzyme [94, 95]. ODC degradation is stimulated by antizyme, a polyamine-induced protein that binds both ODC and the 26S proteasome. Antizyme functions as an adapter much like RAD23 and Dsk2 except that polyubiquitin chains are not involved. However, free ubiquitin chains do compete with antizyme–ODC for degradation

11

12 The 26S Proteasome

[96]. Other potential adapters, such as gankyrin, may target proteins to the 26S proteasome in the absence of ubiquitin marking [97]. The CDK inhibitor p21Cip is degraded in a nonubiquitin-dependent reaction, as clearly demonstrated by substitution of arginines for all the lysine residues in p21Cip. These modifications prevented ubiquitylation of p21Cip, but the lysineless protein was still degraded in human fibroblasts by the proteasome [95]. The Cterminal region of p21Cip binds to the proteasome α-subunit C8, and in vitro p21Cip is degraded by the 20S proteasome alone [98, 99]. Direct binding of p21Cip to the 20S proteasome may open a channel through the α-ring allowing the loosely structured protein to enter the central proteolytic chamber. c-Jun, “aged” calmodulin, troponin C, and p53 are other proteins that can be degraded by the 26S proteasome absent marking by ubiquitin [71]. Thus, other 20S proteasome substrates, in vitro at least, include oxidized proteins, small, denatured proteins and loosely folded proteins such as casein. Whether the 20S proteasome degrades proteins within cells is an unresolved problem.

5 Proteolysis by the 26S Proteasome 5.1 Presumed Mechanism

Proteolysis of ubiquitylated proteins by the 26S proteasome can be thought to consist of seven steps: (1) chaperone-mediated substrate presentation; (2) substrate association with RC subunits; (3) substrate unfolding; (4) detachment of polyubiquitin from the substrate; (5) translocation of the substrate into the 20S proteasome central chamber; (6) peptide bond cleavage; and (7) release of peptide products as well as polyubiquitin (see Figure 4). Step 1 is optional depending on the substrate, and in principle steps 3 and 4 could occur in either order. Step 4 is unnecessary for substrates like ODC that are not ubiquitylated. The other steps almost have to occur as presented. Although it is easy to conceptualize the reaction sequence, few experimental findings bear directly on any of the proposed subreactions, and virtually nothing is known about molecular movements within the 26S proteasome. However several studies on prokaryotic ATP-dependent proteases permit some informed speculation, and it has been shown that step 6 is not required for sequestration of ODC by the 26S proteasome [100]. 5.2 Contribution of Chaperones to Proteasome-mediated Degradation

Chaperones are connected to proteasomes in at least four ways. First, chaperones can deliver substrates to the proteasome as described above for the co-chaperone BAG1 [77]. In a similar fashion the chaperone VCP/Cdc48 is required for the degradation of several ubiquitin-pathway substrates. VCP, a member of the AAA

5 Proteolysis by the 26S Proteasome

Fig. 4 Hypothetical reaction cycle for the 26S proteasome. A polyubiquitylated substrate is delivered to a 26S hybrid proteasome in some cases by chaperones such as VCP/p97 (step 1). Substrate is bound by polyubiquitin recognition components within the regulatory complex (RC) until the polypeptide chain is engaged by the ATPases (step 2). As the polypeptide chain

is unfolded and presumably pumped down the central pore of the proteasome, a signal is conveyed to the S13 metallo-isopeptidase to remove the polyubiquitin chain (step 3). The unfolded polypeptide is eventually degraded within the inner chamber of the proteasome (step 4) and peptide fragments exit the enzyme.

family of ATPases, is a large hexameric ATPase that appears to function as a protein separase able to remove ubiquitylated monomers from multisubunit complexes [101–104]. In some cases the liberated proteins are degraded by the 26S proteasome; in other cases the separated proteins may change their intracellular location. The proteasome also degrades proteins embedded in the endoplasmic reticulum (ER) membrane. If these ER membrane proteins possess a large cytoplasmic domain, their proteasomal degradation can require Hsps 40, 70, and 90 as well as VCP [105, 106]. Hsp90 is required to assemble and stabilize the yeast 26S proteasome providing a third connection between chaperones and proteasomes; Hsp90 is also able to bind and suppress peptide hydrolysis by the 20S proteasome (see below).

13

14 The 26S Proteasome

Finally both chaperones and proteasomes are induced by the accumulation of denatured proteins within eukaryotic cells. 5.2.1 Substrate Binding to the 26S Proteasome Although chaperones may provide for the recognition of some 26S proteasome substrates [107], there is little doubt that the 26S proteasome recognizes substrates directly. As mentioned, ODC is the best-characterized substrate recognized by the 26S proteasome in the absence of a polyubiquitin chain [96]. Which RC subunits actually recognize ODC–antizyme complexes has not been discovered. Presumably one or more subunits in the RC recognize both the C-terminal degron in ODC and some feature of antizyme. The apparent dual recognition of elements in ODC and antizyme may reflect a general mechanism by which ATP-dependent proteases process substrates. For example a number of studies on substrate recognition by E. coli ATP-dependent proteases indicate that substrate adapters provide one recognition site while the substrate provides another [108–110]. Similarly, for the 26S proteasome one recognition element, the attached polyubiquitin chains, may be seen by RC subunits S5a or S2, while the substrate’s N- or C-termini are recognized by RC ATPases. Interestingly, the location of the polyubiquitin chain on the substrate can affect rates of degradation as much as five-fold [111] perhaps by altering the rates at which the RC ATPases engage substrate termini. 5.2.2 Translocation of the Polypeptide Substrate to the Central Proteolytic Chamber Translocation is thought to proceed by the six ATPases threading the polypeptide through a channel in the 20S proteasome’s α-ring. It is also thought that the RC ATPases processively unravel substrates from degrons within the polypeptide chain and are able to “pump” the polypeptide chain in either the N-terminal to C-terminal direction or the opposite [112–114]. Several studies have estimated that hundreds of ATP molecules are needed to degrade small to medium-sized proteins [115, 116]. Rates of translocation range between 10 and 30 amino acids per second [116, 117]. These values are similar to DNA helicases where rates of 50 bases per second and 1ATP per base have been reported [118]. Current models would suggest that the RC ATPases hydrolyze ATP in a sequential rotary fashion essentially screwing the polypeptide chain into the 20S proteasome [119, 120], but the possibility that convulsive movements transfer the substrate has not been ruled out. The ATPases may even be capable of transferring loops into the 20S enzyme. However, a proteomic screen for ClpXP substrates revealed that degrons were either C-terminal or N-terminal [121], so it is likely that the RC ATPases usually engage polypeptide termini. The peptide fragments generated in the central chamber are generally 5 to 10 residues in length, but fragments as long as 35 amino acids can be present [64]. How these fragments exit the central chamber is not known. They could diffuse back through the RC, through small side panels in the 20S proteasome or in the case of hybrid proteasomes (see below) through the end capped by a proteasome activator.

6 Proteasome Biogenesis

5.3 Processing by the 26S Proteasome

In some cases the 26S proteasome partially degrades the substrate protein, releasing processed functional domains. The best-studied example of processing involves the transcriptional activator NFκB. The C-terminal half of a 105-kDa precursor is degraded by the 26S proteasome to yield a 50-kDa N-terminal domain that is the active transcription component [122]. A glycine-rich stretch of amino acids at the C-terminal boundary of p50 is an important factor in limiting proteolysis [123]. It is possible that polypeptide translocation by the RC starts at the Gly-rich region and proceeds in only one direction owing to the presence of the tightly folded N-terminal domain. Or the RC may start translocation at the C-terminus and stop when the ATPases encounter the Gly-rich region. Insertion of a Gly-Ala stretch as small as seven residues into the C-terminal degron of ODC is sufficient to prevent complete destruction of ODC leading instead to partial processing of the enzyme. This has led Coffino and colleagues to suggest that the Gly-Ala stretch impairs substrate transfer by the RC ATPases [124]. Another example of partial processing involves SPT23, a yeast protein embedded in the endoplasmic reticulum membrane [125]. SPT23 controls unsaturated fatty acid levels, and membrane fluidity regulates 26S proteasomal generation of a freely diffusible transcription factor from the SPT23 precursor. Partial processing may be a more widespread regulatory mechanism than is currently thought.

6 Proteasome Biogenesis 6.1 Subunit Synthesis

The synthesis of proteasome subunits is markedly affected by proteasome function. For example, inhibition of proteasome activity by lactacystin induces coordinate expression of both RC and 20S proteasome subunits [66]. Likewise, impaired synthesis of a given RC subunit results in over-expression of all RC subunits [126, 127]. Proteasome subunit synthesis in yeast is controlled by Rpn4p, a short-lived positive transcription factor that binds PACE elements upstream of proteasome genes [128]. Rpn4p is a substrate of the 26S proteasome suggesting that the transcription factor functions in a feedback loop in which proteasome activity limits its concentration thereby regulating proteasome levels [129]. To date an Rpn4-like factor has not been identified in higher eukaryotes, but such a factor is likely to exist. 6.2 Biogenesis of the 20S Proteasome

Proteasome β-subunits are synthesized with N-terminal extensions and are inactive because a free N-terminal threonine is required for peptide-bond hydrolysis [130].

15

16 The 26S Proteasome

The precursor β-subunits assemble with α-subunits to form half proteasomes composed of one α- and one β-ring, which then dimerize to form the 20S particle [131]. The N-terminal extensions are removed thereby generating a new unblocked N-terminal threonine in the catalytically active β-subunits. A small accessory protein called Ump1 in yeast or proteassemblin in mammalian cells assists in the final assembly of the 20S proteasome [132]. Interestingly Ump1/POMP is apparently trapped in the proteasome’s central chamber and degraded upon maturation of the enzyme [133]. 6.3 Biogenesis of the RC

Assembly pathways for the RC are virtually unknown. Asmentioned above, the ATP-ases interact with one another and complexes containing all six S4 subfamily members have been observed following in vitro synthesis. Impaired synthesis of the yeast lid subunit Rpn6 results in the absence of the entire lid [134], so presumably lid and base subcomplexes assemble independently and associate in the final stages of RC formation cells. In mammalian cells, 26S proteasomes assemble from preformed regulatory complexes and 20S proteasomes [135]. 6.4 Post-translational Modification of Proteasome Subunits

Proteasome and RC subunits are subjected to a variety of post-translational modifications including phosphorylation, acetylation, myristoylation, and even Oglycosylation [136–140]. In yeast all seven α-subunits are acetylated as well as two β-subunits. Since acetylation of the N-terminal threonine in an active β-subunit would poison catalysis, it has been suggested that the propeptide extensions function to prevent acetylation [130]. Three members of the S4 ATPase subfamily (S4, S6, and S10b) and two 20S α-subunits (C8 and C9) are known to be phosphorylated. Phosphorylation appears to be particularly important for 26S proteasome assembly and stability. The kinase inhibitor staurosporine reduces 26S proteasome levels in mouse lymphoma cells [135] and interferon γ results in reduced phosphorylation of 20S proteasome α-subunits and decreased 26S proteasome levels [141]. 6.5 Assembly of the 26S Proteasome

The RC and 20S proteasome associate to form the 26S proteasome in the presence of ATP [63]. Comparison of the cross-linking patterns of RC and assembled 26S proteasomes indicates that this association is accompanied by subunit rearrangement [142]. In yeast two proteins play a special role in 26S proteasome assembly or stability. Nob1p is a nuclear protein required for biogenesis of the 26S proteasome and is degraded following assembly of the 26S enzyme. Thus Nob1p suffers the same fate as Ump1 does following 20S maturation [143]. In fission yeast the protein Yin6 regulates both the nuclear localization and the stability of the 26S proteasome

7 Proteasome Activators

[144]. In budding yeast the chaperone Hsp90 also plays a role in the assembly and maintenance of yeast 26S proteasomes, since functional loss of Hsp90 results in 26S proteasome dissociation, indeed even dissociation of the lid subcomplex [145]. The 26S proteasome also dissociates into RC and 20S proteasomes when budding yeast is subjected to long periods of starvation [146].

7 Proteasome Activators

In addition to the RC there are two protein complexes, REGαβ and REGγ , and a single polypeptide chain, PA200, that bind the 20S proteasome and stimulate peptide hydrolysis but not protein degradation. Like the RC, proteasome activators bind the ends of the 20S proteasome and, importantly, they can form mixed or hybrid 26S proteasomes in which one end of the 20S proteasome is associated with a 19S RC and the other is bound to a proteasome activator [147–150]. This latter property raises the possibility that proteasome activators serve to localize the 26S proteasome within eukaryotic cells. 7.1 REGs or PA28s 7.1.1 REGs There are three distinct REG subunits called αβγ [151, 152]. REGsα and β form donut-shaped heteroheptamers found principally in the cytoplasm, whereas REGγ forms a homoheptamer located in the nucleus. REGαβ is abundantly expressed in immune tissues, while REGγ expression is highest in brain. The REGs also differ in their activation properties. REGαβ activates all three proteasome active sites; REGγ only activates the trypsin-like subunit. There is reasonably solid evidence that REGαβ plays a role in Class I antigen presentation [153], but we have little knowledge concerning REGγ function since REGγ knockout mice have almost no phenotype [154]. The crystal structure of REGα reveals that the seven subunits form a donut-shaped structure with a central aqueous channel, and the structure of a REG-proteasome complex provides important insight into the mechanism by which REGα activates the proteasome. The carboxyl tail on each REG subunit fits into a corresponding cavity on the α-ring of the proteasome. Loops on the REG subunits displace N-terminal strands on several proteasome α-subunits reorienting them upward into the aqueous channel of the REG heptamer thereby opening a continuous channel from the exterior solvent to the proteasome central chamber [155]. 7.1.2 PA200 A new proteasome activator called PA200 was recently purified from bovine testis [156]. Human PA200 is a nuclear protein of 1843 amino acids that activates all three catalytic subunits with some preference for the PGPH active site. Homologs

17

18 The 26S Proteasome

of PA200 are present in budding yeast, worms, and plants. A single chain of PA200 can bind each end of the proteasome and, when bound, PA200 molecules look like volcanos in negatively stained EM images. PA200 is thought to play a role in DNA repair, perhaps by recruiting proteasomes to DNA double-strand breaks. 7.1.3 Hybrid Proteasomes As the α-rings at each end of the 20S proteasome are equivalent, the 20S proteasome is capable of binding two RCs, two PA28s, two PA200s, or combinations of these components. In fact 20S proteasomes simultaneously bound to RC and PA28 or PA200 have been observed, and are called hybrid proteasomes [147, 148, 150]. In HeLa cells the levels of hybrid proteasomes containing PA28 at one end and an RC at the other are two-fold higher than 26S proteasomes capped at both ends by 19S RCs [149]. Hybrid 26S proteasomes containing PA200 appear to be much less abundant in HeLa cells [156]. 7.2 ECM29

Another proteasome-associated protein, called Ecm29, has been identified in several proteomic screens [157, 158]. Ecm29p clearly associates with 26S proteasomes; whether it activates proteasomal peptide hydrolysis is currently unknown. Ecm29p is reported to stabilize the yeast 26S proteasome by clamping the RC to the 20S cylinder [159]. However, in mammalian cells Ecm29p is found mainly associated to secretory and endocytic organelles, a location suggesting a role in secretion rather than 26S proteasome stability. Moreover, the levels of Ecm29p vary markedly among mouse organs, so if mammalian Ecm29p serves as a clamp, some tissues either do not require a clamped 26S proteasome or other proteins function as RC-20S clamps.

8 Protein Inhibitors of the Proteasome

A number of proteins have been found to suppress proteolysis by the proteasome. One of these is PI31 [160, 161]; another is the abundant cytosolic chaperone Hsp90 [162], and a third is a proline/arginine-rich 39-residue peptide called PR39 [163]. Both PI31 and Hsp90 may affect how the proteasome functions in Class I antigen presentation. PI31 is a 30-kDa proline-rich protein that inhibits peptide hydrolysis by the 20S proteasome and can block activation by both RC and REGαβ. Although surveys of various cell lines show PI31 to be considerably less abundant than RC or REGαβ, when over-expressed, PI31 is reported to inhibit Class I antigen presentation by interfering with the assembly of immuno-proteasomes [161]. A number of studies have shown that Hsp90 can bind the 20S proteasome and inhibit its chymotrypsin-like and PGPH activities. Interestingly inhibition by Hsp90 is observed with constitutive but not with immuno-proteasomes, a finding consistent

9 Physiological Aspects 19

with proposals that Hsp90 shuttles immuno-proteasome-generated peptides to the endoplasmic reticulum for Class I presentation [164]. PR39 was originally isolated from bone marrow as a factor able to induce angiogenesis and inhibit inflammation. Two hybrid screens showed that PR39 binds the 20S proteasome. Apparently PR39 affects angiogenesis and inflammation by inhibiting respectively the degradation of HIF1 or IκBα, the latter being an inhibitor of NFκB. Finally HIV’s Tat protein inhibits the 20S proteasome’s peptidase activity [165]. Tat also competes with REGαβ for proteasome binding and, by doing so, Tat can inhibit Class I presentation of certain epitopes [166].

9 Physiological Aspects 9.1 Tissue and Subcellular Distribution of Proteasomes

Proteasomes are found in all organs of higher eukaryotes, but the degree to which the composition of proteasomes and its activators varies among tissues is largely unexplored territory. Proteasomes are very abundant in testis which contains almost five-fold more 20S subunits than skeletal muscle [167]. At the cellular level there are about 800 000 proteasomes in a HeLa cell and roughly 20000 proteasomes in a yeast cell. At the subcellular level, 26S proteasomes are present in cytosol and nucleus where they appear to be freely diffusible [168, 169]. They are not usually found in the nucleolus [170] and have not been reported in membrane-bound organelles other than the nucleus. When large amounts of misfolded proteins are synthesized by a cell, the aberrant polypeptides often accumulate around the centrosome in what are called “aggresomes” [171]. Under these conditions 26S proteasomes, chaperones, and proteasome activators also redistribute to the aggresomes, presumably to refold and/or degrade the misfolded polypeptides [172].

9.2 Physiological Importance

Deletion of yeast genes encoding 20S proteasome and 19S RC subunits is usually lethal, indicating that the 26S proteasome is required for eukaryotic cell viability. Known substrates of the 26S proteasome include transcription factors, cell-cycle regulators, protein kinases, etc., essentially most of the cell’s important regulatory proteins. Even proteins secreted into the endoplasmic reticulum are returned to the cytosol for degradation by the 26S proteasome [173, 174]. Given the scope of its substrates it is hardly surprising that in higher eukaryotes the ubiquitin–proteasome system contributes to the regulation of a vast array of physiological processes ranging from cell-cycle traverse to circadian rhythms to learning.

20 The 26S Proteasome

10 Conclusions

The 20S proteasome was discovered in 1980 and the 26S proteasome six years later. Research since the mid-1980s has made it abundantly clear that the ubiquitinproteasome system is of central importance in eukaryotic cell physiology. Yet there is much more to discover. A crystal structure of the 19S RC, or better still the 26S proteasome, would surely provide insight into the mechanism by which the 26S proteasome degrades its substrates. How the 26S proteasome is itself regulated and the extent to which proteasomal components vary among tissues in higher eukaryotes are other important unresolved problems. Hopefully, these and other unanswered questions will spark further interest in the proteasome.

References 1 REED, S. I. Ratchets and clocks: the cell cycle, ubiquitylation and protein turnover. Nat Rev Mol Cell Biol 2003, 4, 855–864. 2 BASHIR, T. and PAGANO, M. Aberrant ubiquitin-mediated proteolysis of cell cycle regulatory proteins and oncogenesis. Adv Cancer Res 2003, 88, 101–144. 3 VIERSTRA, R. D. The ubiquitin/26S proteasome pathway, the complex last chapter in the life of many plant proteins. Trends Plant Sci 2003, 8, 135–142. 4 CAMPBELL, D. S. and HOLT, C. E. Chemotropic responses of retinal growth cones mediated by rapid local protein synthesis and degradation. Neuron 2001, 32, 1013–1026. 5 HEGDE, A. N. and DIANTONIO, A. Ubiquitin and the synapse. Nat Rev Neurosci 2002, 3, 854–861. 6 MURATANI, M. and TANSEY, W. P. How the Ubiquitin–Proteasome system controls transcription. Nat Rev Mol Cell Biol 2003, 4, 192–201. 7 CONAWAY, R. C., BROWER, C. S., and CONAWAY, J. W. Emerging roles of ubiquitin in transcription regulation. Science 2002, 296, 1254–1258. 8 LIPFORD, J. R. and DESHAIES, R. J. Diverse roles for ubiquitin-dependent proteolysis in transcriptional activation. Nat Cell Biol 2003, 5, 845–850.

9 SEELER, J. S. and DEJEAN, A. Nuclear and unclear functions of SUMO. Nat Rev Mol Cell Biol 2003, 4, 690–699. 10 SCHWARTZ, D. C. and HOCHSTRASSER, M. A superfamily of protein tags: ubiquitin, SUMO and related modifiers. Trends Biochem Sci 2003, 28, 321–328. 11 JASON, L. J., MOORE, S. C., LEWIS, J. D., LINDSEY, G., and AUSIO, J. Histone ubiquitination: a tagging tail unfolds? Bioessays 2002, 24, 166–174. 12 ZHANG, Y. Transcriptional regulation by histone ubiquitination and deubiquitination. Genes Dev 2003, 17, 2733–2740. 13 DENG, L. et al. Activation of the IkappaB kinase complex by TRAF6 requires a dimeric ubiquitin-conjugating enzyme complex and a unique polyubiquitin chain. Cell 2000, 103, 351–361. 14 HICKE, L. and DUNN, R. Regulation of membrane protein transport by ubiquitin and ubiquitin-binding proteins. Annu Rev Cell Dev Biol 2003, 19, 141–172. 15 PORNILLOS, O., GARRUS, J. E., and SUNDQUIST, W. I. Mechanisms of enveloped RNA virus budding. Trends Cell Biol 2002, 12, 569–579. 16 HERSHKO, A., CIECHANOVER, A., and VARSHAVSKY, A. Basic Medical Research Award. The ubiquitin system. Nat Med 2000, 6, 1073–1081.

References 17 KESSEL, M. et al. Homology in structural organization between E. coli ClpAP protease the eukaryotic 26S proteasome. J Mol Biol 1995, 250, 587–594. 18 LOWE, J. et al. Crystal structure of the 20S proteasome from the archaeon T. acidophilum at 3.4 A resolution. Science 1995, 268, 533–539. 19 GROLL, M. et al. Structure of 20S proteasome from yeast at 2.4 A resolution. Nature 1997, 386, 463–471. 20 UNNO, M. et al. The structure of the mammalian 20S proteasome at 2.75 A resolution. Structure (Camb) 2002, 10, 609–618. 21 DAHLMANN, B. et al. The multicatalytic proteinase (prosome, proteasome): comparison of the eukaryotic and archaebacterial enzyme. Biomed Biochim Acta 1991, 50, 465–469. 22 HARRIS, J. L., ALPER, P. B., LI, J., RECHSTEINER, M., and BACKES, B. J. Substrate specificity of the human proteasome. Chem Biol 2001, 8, 1131–1141. 23 SEEMULLER, E. et al. Proteasome from Thermoplasma acidophilum: a threonine protease. Science 1995, 268, 579–582. 24 FENTEANY, G. et al. Inhibition of proteasome activities and subunit-specific amino-terminal threonine modification by lactacystin. Science 1995, 268, 726–731. 25 MENG, L. et al. Epoxomicin, a potent and selective proteasome inhibitor, exhibits in vivo anti-inflammatory activity. Proc. Natl. Acad. Sci. USA 1999, 96, 10403–10408. 26 BOGYO, M. et al. Covalent modification of the active site threonine of proteasomal β subunits and the Escherichia coli homolg HsIV by a new class of inhibitors. Proc. Natl. Acad. Sci. USA 1997, 94, 6629–6634. 27 KANE, R. C., BROSS, P. F., FARRELL, A. T., and PAZDUR, R. Velcade: U.S. FDA approval for the treatment of multiple myeloma progressing on prior therapy. Oncologist 2003, 8, 508–513. 28 YEWDELL, J. W. and HILL, A. B. Viral interference with antigen presentation. Nat Immunol 2002, 3, 1019–1025.

29 GOLDBERG, A. L., CASCIO, P., SARIC, T., and ROCK, K. L. The importance of the proteasome and subsequent proteolytic steps in the generation of antigenic peptides. Mol Immunol 2002, 39, 147–164. 30 RAMMENSEE, H.-G., FALK, K., and R¨OTZSCHKE, O. Peptides naturally presented by MHC class I molecules. Annu. Rev. Immunol. 1993, 11, 213–244. 31 DOUGAN, D. A., MOGK, A., ZETH, K., TURGAY, K., and BUKAU, B. AAA+ proteins and substrate recognition, it all depends on their partner in crime. FEBS Lett 2002, 529, 6–10. 32 PETERS, J. M., CEJKA, Z., HARRIS, J. R., KLEINSCHMIDT, J. A., and BAUMEISTER, W. Structural features of the 26S proteasome complex. J Mol Biol 1993, 234, 932–937. 33 WALZ, J. et al. 26S proteasome structure revealed by three-dimensional electron microscopy. J Struct Biol 1998, 121, 19–29. 34 GLICKMAN, M. H. et al. A subcomplex of the proteasome regulatory particle required for ubiquitin-conjugate degradation and related to the COP9-signalosome and elF3. Cell 1998, 94, 615–623. 35 KAPELARI, B. et al. Electron microscopy and subunit-subunit interaction studies reveal a first architecture of COP9 signalosome. J Mol Biol 2000, 300, 1169–1178. 36 RECHSTEINER, M., HOFFMAN, L., and DUBIEL, W. The multicatalytic and 26S proteases. J. Biol. Chem. 1993, 268, 6065–6068. 37 RICHMOND, C., GORBEA, C., and RECHSTEINER, M. Specific interactions between ATPase subunits of the 26S protease. J. Biol. Chem. 1997, 272, 13403–13411. 38 RUBIN, D. M., GLICKMAN, M. H., LARSEN, C. N., DHRUVAKUMAR, S., and FINLEY, D. Active site mutants in the six regulatory particle ATPases reveal multiple roles for ATP in the proteasome. EMBO J. 1998, 17, 4909–4919. 39 DAVEY, M. J., JERUZALMI, D., KURIYAN, J., and O’DONNELL, M. Motors and switches: AAA+ machines within the

21

22 The 26S Proteasome

40

41

42

43

44

45

46

47

48

49

50

replisome. Nat Rev Mol Cell Biol 2002, 3, 826–835. LEE, S. Y. et al. Regulation of the transcriptional activator NtrC1: structural studies of the regulatory and AAA+ ATPase domains. Genes Dev 2003, 17, 2552–2563. HARTMANN-PETERSEN, R., TANAKA, K., and HENDIL, K. B. Quaternary structure of the ATPase complex of human 26S proteasomes determined by chemical cross-linking. Arch Biochem Biophys 2001, 386, 89–94. BERNDT, C., BECH-OTSCHIR, D., DUBIEL, W., and SEEGER, M. Ubiquitin system: JAMMing in the name of the lid. Curr Biol 2002, 12, R815–R817. COPE, G. A. and DESHAIES, R. J. COP9 signalosome: a multifunctional regulator of SCF and other cullin-based ubiquitin ligases. Cell 2003, 114, 663–671. LI, L. and DENG, X. W. The COP9 signalosome: an alternative lid for the 26S proteasome? Trends Cell Biol 2003, 13, 507–509. FERRELL, K., WILKINSON, C. R., DUBIEL, W., and GORDON, C. Regulatory subunit interactions of the 26S proteasome, a complex problem. Trends Biochem Sci 2000, 25, 83–88. FU, H., REIS, N., LEE, Y., GLICKMAN, M. H., and VIERSTRA, R. D. Subunit interaction maps for the regulatory particle of the 26S proteasome and the COP9 signalosome. Embo J 2001, 20, 7096–7107. CAGNEY, G., UETZ, P., and FIELDS, S. Two-hybrid analysis of the Saccharomyces cerevisiae 26S proteasome. Physiol Genomics 2001, 7, 27–34. DAVY, A. et al. A protein-protein interaction map of the Caenorhabditis elegans 26S proteasome. EMBO Rep 2001, 2, 821–828. COPE, G. A. et al. Role of predicted metalloprotease motif of Jab1/Csn5 in cleavage of Nedd8 from Cul1. Science 2002, 298, 608–611. BAILLY, E. and REED, S. I. Functional characterization of rpn3 uncovers a distinct 19S proteasomal subunit requirement for ubiquitin-dependent proteolysis of cell cycle regulatory

51

52

53

54

55

56

57

58

59

60

proteins in budding yeast. Mol Cell Biol 1999, 19, 6872–6890. FONG, A., ZHANG, M., NEELY, J., and SUN, S. C. S9, a 19 S proteasome subunit interacting with ubiquitinated NF-kappaB2/p100. J Biol Chem 2002, 277, 40697–40702. DEVERAUX, Q., USTRELL, V., PICKART, C., and RECHSTEINER, M. A 26S protease subunit that binds ubiquitin conjugates. J. Biol. Chem. 1994, 269, 7059–7061. van NOCKER, S. et al. The Multiubiquitin-Chain-Binding Protein McB1 Is a Component of the 26S Proteasome in Saccharomyces cerevisiae and Plays a Nonessentail, Substrate-Specific Role in Protein Turnover. Mol. Cell Biol. 1996, 16, 6020–6028. KAJAVA, A. V. What curves alphasolenoids? Evidence for an alpha-helical toroid structure of Rpn1 and Rpn2 proteins of the 26S proteasome. J Biol Chem 2002, 277, 49791–49798. ELSASSER, S. et al. Proteasome subunit Rpn1 binds ubiquitin-like protein domains. Nat Cell Biol 2002, 4, 725–730. SAEKI, Y., SONE, T., TOHE, A., and YOKOSAWA, H. Identification of ubiquitin-like protein-binding subunits of the 26S proteasome. Biochem Biophys Res Commun 2002, 296, 813–819. LAM, Y. A., LAWSON, T. G., VELAYUTHAM, M., ZWEIER, J. L., and PICKART, C. M. A proteasomal ATPase subunit recognizes the polyubiquitin degradation signal. Nature 2002, 416, 763–767. HOFFMAN, L. and RECHSTEINER, M. Activation of the multicatalytic protease. The 11 S regulator and 20S ATPase complexes contain distinct 30-kilodalton subunits. J. Biol. Chem. 1994, 269, 16890–16895. BRAUN, B. C. et al. The base of the proteasome regulatory particle exhibits chaperone-like activity. Nat Cell Biol 1999, 1, 221–226. STRICKLAND, E., HAKALA, K., THOMAS, P. J., and DEMARTINO, G. N. Recognition of misfolding proteins by PA700, the regulatory subcomplex of the 26S

References

61

62

63

64

65

66

67 68

69

70

71

proteasome. J Biol Chem 2000, 275, 5565–5572. LIU, C. W. et al. Conformational remodeling of proteasomal substrates by PA700, the 19 S regulatory complex of the 26S proteasome. J Biol Chem 2002, 277, 26815–26820. GONZALEZ, F., DELAHODDE, A., KODADEK, T., and JOHNSTON, S. A. Recruitment of a 19S proteasome subcomplex to an activated promoter. Science 2002, 296, 548–550. HOFFMAN, L., PRATT, G., and RECHSTEINER, M. Multiple forms of the 20 S multicatalytic and the 26S ubiquitin/ATP-dependent proteases from rabbit reticulocyte lysate. J. Biol. Chem. 1992, 267, 22362–22368. VERMA, R. et al. Proteasomal proteomics: identification of nucleotide-sensitive proteasome-interacting proteins by mass spectrometric analysis of affinity-purified proteasomes. Mol Biol Cell 2000, 11, 3425–3439. GUTERMAN, A. and GLICKMAN, M. H. Complementary roles for rpn11 and ubp6 in deubiquitination and proteolysis by the proteasome. J Biol Chem 2004, 279, 1729–1738. MEINERS, S. et al. Inhibition of proteasome activity induces concerted expression of proteasome genes and de novo formation of Mammalian proteasomes. J Biol Chem 2003, 278, 21517–21525. BLOBEL, G. Protein targeting (Nobel lecture). Chembiochem 2000, 1, 86–102. RECHSTEINER, M. and ROGERS, S. W. PEST sequences and regulation by proteolysis. Trends Biochem. Sci. 1996, 21, 267–271. VARSHAVSKY, A. The N-end rule: Functions, mysteries, uses. Proc. Natl. Acad. Sci. USA 1996, 93, 12142–12149. ZUR, A. and BRANDEIS, M. Timing of APC/C substrate degradation is determined by fzy/fzr specificity of destruction boxes. Embo J 2002, 21, 4500–4510. ORLOWSKI, M. and WILK, S. Ubiquitin-independent proteolytic functions of the proteasome. Arch Biochem Biophys 2003, 415, 1–5.

72 GRUNE, T., MERKER, K., SANDIG, G., and DAVIES, K. J. Selective degradation of oxidatively modified protein substrates by the proteasome. Biochem Biophys Res Commun 2003, 305, 709–718. 73 PENG, J. et al. A proteomics approach to understanding protein ubiquitination. Nat Biotechnol 2003, 21, 921–926. 74 WU-BAER, F., LAGRAZON, K., YUAN, W., and BAER, R. The BRCA1/BARD1 heterodimer assembles polyubiquitin chains through an unconventional linkage involving lysine residue K6 of ubiquitin. J Biol Chem 2003, 278, 34743–34746. 75 SPENCE, J., SADIS, S., HAAS, A. L., and FINLEY, D. A ubiquitin mutant with specific defects in DNA repair and multiubiquitination. Mol Cell Biol 1995, 15, 1265–1273. 76 SPRINGAEL, J. Y., GALAN, J. M., HAGUENAUER-TSAPIS, R., and ANDRE, B. NH4+-induced down-regulation of the Saccharomyces cerevisiae Gap1p permease involves its ubiquitination with lysine-63-linked chains. J Cell Sci 1999, 112 (Pt 9), 1375–1383. 77 ALBERTI, S. et al. Ubiquitylation of BAG-1 suggests a novel regulatory mechanism during the sorting of chaperone substrates to the proteasome. J Biol Chem 2002, 277, 45920–45927. 78 CHAU, V. et al. A multiubiquitin chain is confined to specific lysine in a targeted short-lived protein. Science 1989, 243, 1576–1583. 79 JOHNSON, E. S., MA, P. C., OTA, I. M., and VARSHAVSKY, A. A proteolytic pathway that recognizes ubiquitin as a degradation signal. J. Biol. Chem. 1995, 270, 17442–17456. 80 THROWER, J. S., HOFFMAN, L., RECHSTEINER, M., and PICKART, C. M. Recognition of the polyubiquitin proteolytic signal. EMBO J. 2000, 19, 94–102. 81 YOUNG, P., DEVERAUX, Q., BEAL, R., PICKART, C., and RECHSTEINER, M. Characterization of two polyubiquitin binding sites in the 26S protease subunit 5a. J. Biol. Chem. 1998, 273, 5461–5467. 82 HOFMANN, K. and FALQUET, L. A ubiquitin-interacting motif conserved in

23

24 The 26S Proteasome

83

84

85

86

87

88

89

90

91

92

components of the proteasomal and lysosomal protein degradation systems. Trends Biochem Sci 2001, 26, 347–350. SWANSON, K. A., KANG, R. S., STAMENOVA, S. D., HICKE, L., and RADHAKRISHNAN, I. Solution structure of Vps27 UIM-ubiquitin complex important for endosomal sorting and receptor downregulation. Embo J 2003, 22, 4597–4606. MUELLER, T. D. and FEIGON, J. Structural determinants for the binding of ubiquitin-like domains to the proteasome. Embo J 2003, 22, 4634–4645. RYU, K. S. et al. Binding surface mapping of intra- and interdomain interactions among hHR23B, ubiquitin, and polyubiquitin binding site 2 of S5a. J Biol Chem 2003, 278, 36621–36627. HARTMANN-PETERSEN, R., SEEGER, M., and GORDON, C. Transferring substrates to the 26S proteasome. Trends Biochem Sci 2003, 28, 26–31. SAEKI, Y., SAITOH, A., TOHE, A., and YOKOSAWA, H. Ubiquitin-like proteins and Rpn10 play cooperative roles in ubiquitin-dependent proteolysis. Biochem Biophys Res Commun 2002, 293, 986–992. HIYAMA, H. et al. Interaction of hHR23 with S5a. The ubiquitin-like domain of hHR23 mediates interaction with S5a subunit of 26S proteasome. J Biol Chem 1999, 274, 28019–28025. SAKATA, E. et al. Parkin binds the Rpn10 subunit of 26S proteasomes through its ubiquitin-like domain. EMBO Rep 2003, 4, 301–306. KLEIJNEN, M. F., ALARCON, R. M., and HOWLEY, P. M. The ubiquitin-associated domain of hPLIC-2 interacts with the proteasome. Mol Biol Cell 2003, 14, 3868–3875. XIE, Y. and VARSHAVSKY, A. UFD4 lacking the proteasome-binding region catalyses ubiquitination but is impaired in proteolysis. Nat Cell Biol 2002, 4, 1003–1007. CORN, P. G., MCDONALD, E. R., 3rd, HERMAN, J. G., and EL-DEIRY, W. S. Tat-binding protein-1, a component of the 26S proteasome, contributes to the

93

94

95

96

97

98

99

100

101

102

103

E3 ubiquitin ligase function of the von Hippel-Lindau protein. Nat Genet 2003, 35, 229–237. LAMBERTSON, D., CHEN, L., and MADURA, K. Pleiotropic defects caused by loss of the proteasome-interacting factors Rad23 and Rpn10 of Saccharomyces cerevisiae. Genetics 1999, 153, 69–79. MURAKAMI, Y. et al. Ornithine decarboxylase is degraded by the 26S proteasome without ubiquitination. Nature 1992, 360, 597–599. SHEAFF, R. J. et al. Proteasome turnover of p21Cip1 does not require p21Cip1 ubiquitination. Molec. Cell. 2000, 5, 403–410. ZHANG, M., PICKART, C. M., and COFFINO, P. Determinants of proteasome recognition of ornithine decarboxylase, a ubiquitin-independent substrate. Embo J 2003, 22, 1488–1496. REZVANI, K. et al. Proteasomal interactors control activities as diverse as the cell cycle and glutaminergic neurotransmission. Biochem Soc Trans 2003, 31, 470–473. TOUITOU, R. et al. A degradation signal located in the C-terminus of p21WAF1/CIP1 is a binding site for the C8 alpha-subunit of the 20S proteasome. Embo J 2001, 20, 2367–2375. COLEMAN, M. L., MARSHALL, C. J., and OLSON, M. F. Ras promotes p21(Waf1/Cip1) protein stability via a cyclin D1-imposed block in proteasome-mediated degradation. Embo J 2003, 22, 2036–2046. MURAKAMI, Y., MATSUFUJI, S., HAYASHI, S. I., TANAHASHI, N., and TANAKA, K. ATP-Dependent inactivation and sequestration of ornithine decarboxylase by the 26S proteasome are prerequisites for degradation. Mol Cell Biol 1999, 19, 7216–7227. BAYS, N. W. and HAMPTON, R. Y. Cdc48-Ufd1-Npl4: stuck in the middle with Ub. Curr Biol 2002, 12, R366–R371. BRUNGER, A. T. and DELABARRE, B. NSF and p97/VCP: similar at first, different at last. FEBS Lett 2003, 555, 126–133. RABINOVICH, E., KEREM, A., FROHLICH, K. U., DIAMANT, N., and BAR-NUN, S. AAA-ATPase p97/Cdc48p, a cytosolic

References

104

105

106

107

108

109

110

111

112

chaperone required for endoplasmic reticulum-associated protein degradation. Mol Cell Biol 2002, 22, 626–634. WOJCIK, C., YANO, M., and DEMARTINO, G. N. RNA interference of valosin-containing protein (VCP/p97) reveals multiple cellular roles linked to ubiquitin/proteasome-dependent proteolysis. J Cell Sci 2004, 117, 281–292. IMAMURA, T. et al. Involvement of heat shock protein 90 in the degradation of mutant insulin receptors by the proteasome. J Biol Chem 1998, 273, 11183–11188. TAXIS, C. et al. Use of modular substrates demonstrates mechanistic diversity and reveals differences in chaperone requirement of ERAD. J Biol Chem 2003, 278, 35903–35913. TOKUMOTO, T. et al. Regulated interaction between polypeptide chain elongation factor-1 complex with the 26S proteasome during Xenopus oocyte maturation. BMC Biochem 2003, 4, 6. GONCIARZ-SWIATEK, M. et al. Recognition, targeting, and hydrolysis of the lambda O replication protein by the ClpP/ClpX protease. J Biol Chem 1999, 274, 13999–14005. STUDEMANN, A. et al. Sequential recognition of two distinct sites in sigma(S) by the proteolytic targeting factor RssB and ClpX. Embo J 2003, 22, 4111–4120. NEHER, S. B., SAUER, R. T., and BAKER, T. A. Distinct peptide signals in the UmuD and UmuD subunits of UmuD/D mediate tethering and substrate processing by the ClpXP protease. Proc Natl Acad Sci USA 2003, 100, 13219–13224. PETROSKI, M. D. and DESHAIES, R. J. Context of multiubiquitin chain attachment influences the rate of Sic1 degradation. Mol Cell 2003, 11, 1435–1444. LEE, C., SCHWARTZ, M. P., PRAKASH, S., IWAKURA, M., and MATOUSCHEK, A. ATP-dependent proteases degrade their substrates by processively unraveling them from the degradation signal. Mol Cell 2001, 7, 627–637.

113 HOSKINS, J. R., YANAGIHARA, K., MIZUUCHI, K., and WICKNER, S. ClpAP and ClpXP degrade proteins with tags located in the interior of the primary sequence. Proc Natl Acad Sci USA 2002, 99, 11037–11042. 114 LEE, C., PRAKASH, S., and MATOUSCHEK, A. Concurrent translocation of multiple polypeptide chains through the proteasomal degradation channel. J Biol Chem 2002, 277, 34760–34765. 115 BENAROUDJ, N., ZWICKL, P., SEEMULLER, E., BAUMEISTER, W., and GOLDBERG, A. L. ATP hydrolysis by the proteasome regulatory complex PAN serves multiple functions in protein degradation. Mol Cell 2003, 11, 69–78. 116 KENNISTON, J. A., BAKER, T. A., FERNANDEZ, J. M., and SAUER, R. T. Linkage between ATP consumption and mechanical unfolding during the protein processing reactions of an AAA+ degradation machine. Cell 2003, 114, 511–520. 117 REID, B. G., FENTON, W. A., HORWICH, A. L., and WEBER-BAN, E. U. ClpA mediates directional translocation of substrate proteins into the ClpP protease. Proc Natl Acad Sci USA 2001, 98, 3768–3772. 118 DILLINGHAM, M. S., WIGLEY, D. B., and WEBB, M. R. Demonstration of unidirectional single-stranded DNA translocation by PcrA helicase: measurement of step size and translocation speed. Biochemistry 2000, 39, 205–212. 119 LLORCA, O. et al. The ‘sequential allosteric ring’ mechanism in the eukaryotic chaperonin-assisted folding of actin and tubulin. Embo J 2001, 20, 4065–4075. 120 LASKEY, R. A. and MADINE, M. A. A rotary pumping model for helicase function of MCM proteins at a distance from replication forks. EMBO Rep 2003, 4, 26–30. 121 FLYNN, J. M., NEHER, S. B., KIM, Y. I., SAUER, R. T., and BAKER, T. A. Proteomic discovery of cellular substrates of the ClpXP protease reveals five classes of ClpX-recognition signals. Mol Cell 2003, 11, 671–683.

25

26 The 26S Proteasome

122 PALOMBELLA, V. J., RANDO, O. J., GOLDBERG, A. L., and MANIATIS, T. The Ubiquitin–Proteasome pathway is required for processing the NF-kappa B1 precursor protein and the activation of NF-kappa B. Cell 1994, 78, 773–785. 123 CIECHANOVER, A. et al. Mechanisms of ubiquitin-mediated, limited processing of the NF-kappaB1 precursor protein p105. Biochimie 2001, 83, 341–349. 124 ZHANG, M. and COFFINO, P. Repeat sequence of Epstein-Barr virus EBNA1 protein interrupts proteasome substrate processing. J Biol Chem 2004, 279, 8635–8641. 125 HOPPE, T. et al. Activation of a membrane-bound transcription factor by regulated ubiquitin/proteasomedependent processing. Cell 2000, 102, 577–586. 126 WOJCIK, C. and DEMARTINO, G. N. Analysis of Drosophila 26S proteasome using RNA interference. J Biol Chem 2002, 277, 6188–6197. 127 SZLANKA, T. et al. Deletion of proteasomal subunit S5a/Rpn10/p54 causes lethality, multiple mitotic defects and overexpression of proteasomal genes in Drosophila melanogaster. J Cell Sci 2003, 116, 1023–1033. 128 MANNHAUPT, G., SCHNALL, R., KARPOV, V., VETTER, I., and FELDMANN, H. Rpn4p acts as a transcription factor by binding to PACE, a nonamer box found upstream of 26S proteasomal and other genes in yeast. FEBS Lett 1999, 450, 27–34. 129 XIE, Y. and VARSHAVSKY, A. RPN4 is a ligand, substrate, and transcriptional regulator of the 26S proteasome: a negative feedback circuit. Proc Natl Acad Sci USA 2001, 98, 3056–3061. 130 ARENDT, C. S. and HOCHSTRASSER, M. Eukaryotic 20S proteasome catalytic subunit propeptides prevent active site inactivation by N-terminal acetylation and promote particle assembly. Embo J 1999, 18, 3575–3585. 131 NANDI, D., WOODWARD, E., GINSBURG, D. B., and MONACO, J. J. Intermediates in the formation of mouse 20S proteasomes: implications for the

132

133

134

135

136

137

138

139

140

assembly of precursor beta subunits. Embo J 1997, 16, 5363–5375. GRIFFIN, T. A., SLACK, J. P., MCCLUSKEY, T. S., MONACO, J. J., and COLBERT, R. A. Identification of proteassemblin, a mammalian homologue of the yeast protein, Ump1p, that is required for normal proteasome assembly. Mol Cell Biol Res Commun 2000, 3, 212–217. RAMOS, P. C., HOCKENDORFF, J., JOHNSON, E. S., VARSHAVSKY, A., and DOHMEN, R. J. Ump1p is required for proper maturation of the 20S proteasome and becomes its substrate upon completion of the assembly. Cell 1998, 92, 489–499. SANTAMARIA, P. G., FINLEY, D., BALLESTA, J. P., and REMACHA, M. Rpn6p, a proteasome subunit from Saccharomyces cerevisiae, is essential for the assembly and activity of the 26S proteasome. J Biol Chem 2003, 278, 6687–6695. YANG, Y., FRU¨ H, K., AHN, K., and PETERSON, P. A. In vivo assembly of the proteasomal complexes, implications for antigen processing. J. Biol. Chem. 1995, 270, 27687–27694. MASON, G. G., HENDIL, K. B., and RIVETT, A. J. Phosphorylation of proteasomes in mammalian cells. Identification two phosphorylated subunits and the effect of phosphorylation on activity. Eur. J. Biochem. 1996, 238, 453–462. MASON, G. G., MURRAY, R. Z., PAPPIN, D., and RIVETT, A. J. Phosphorylation of ATPase subunits of the 26S proteasome. FEBS Lett 1998, 430, 269–274. KIMURA, Y. et al. N(alpha)-acetylation and proteolytic activity of the yeast 20 S proteasome. J Biol Chem 2000, 275, 4635–4639. KIMURA, Y. et al. N-Terminal modifications of the 19S regulatory particle subunits of the yeast proteasome. Arch Biochem Biophys 2003, 409, 341–348. SUMEGI, M., HUNYADI-GULYAS, E., MEDZIHRADSZKY, K. F., and UDVARDY, A. 26S proteasome subunits are O-linked N-acetylglucosamine-modified in Drosophila melanogaster. Biochem Biophys Res Commun 2003, 312, 1284–1289.

References 141 BOSE, S., STRATFORD, F. L., BROADFOOT, K. I., MASON, G. G., and RIVETT, A. J. Phosphorylation of 20S proteasome alpha subunit C8 (alpha7) stabilizes the 26S proteasome and plays a role in the regulation of proteasome complexes by gamma-interferon. Biochem J Pt 2004, 378, 177–184. 142 KURUCZ, E. et al. Assembly of the Drosophila 26S proteasome is accompanied by extensive subunit rearrangements. Biochem J 2002, 365, 527–536. 143 TONE, Y. and TOH, E. A. Nob1p is required for biogenesis of the 26S proteasome and degraded upon its maturation in Saccharomyces cerevisiae. Genes Dev 2002, 16, 3142–3157. 144 YEN, H. C., GORDON, C., and CHANG, E. C. Schizosaccharomyces pombe Int6 and Ras homologs regulate cell division and mitotic fidelity via the proteasome. Cell 2003, 112, 207–217. 145 IMAI, J., MARUYA, M., YASHIRODA, H., YAHARA, I., and TANAKA, K. The molecular chaperone Hsp90 plays a role in the assembly and maintenance of the 26S proteasome. Embo J 2003, 22, 3557–3567. 146 BAJOREK, M., FINLEY, D., and GLICKMAN, M. H. Proteasome disassembly and downregulation is correlated with viability during stationary phase. Curr Biol 2003, 13, 1140–1144. 147 HENDIL, K. B., KHAN, S. and TANAKA, K. Simultaneous binding of PA28 and PA700 activators to 20 S proteasomes. Biochem. J. 1998, 332, 749–754. 148 KOPP, F., DAHLMANN, B., and KUEHN, L. Reconstitution of hybrid proteasomes from purified PA700–20 S complexes and PA28alphabeta activator: ultrastructure and peptidase activities. J Mol Biol 2001, 313, 465–471. 149 TANAHASHI, N. et al. Hybrid proteasomes. Induction by interferon-gamma and contribution to ATP-dependent proteolysis. J. Biol. Chem. 2000, 275, 14336–14345. 150 CASCIO, P., CALL, M., PETRE, B. T. W., and GOLDBERG, A. L. Properties of the hybrid form of the 26S proteasome containing

151

152

153

154

155

156

157

158

159

160

161

162

both 19S and PA28 complexes. EMBO J. 2002, 21, 2636–2645. RECHSTEINER, M., REALINI, C., and USTRELL, V. The proteasome activator 11S REG (PA28) and class I antigen presentation. Biochem. J. 2000, 345, 1–15. HILL, C. P., MASTERS, E. I., and WHITBY, F. G. The 11S regulators of 20S proteasome activity. Curr Top Microbiol Immunol 2002, 268, 73–89. MURATA, S. et al. Immunoproteasome assembly and antigen presentation in mice lacking both PA28alpha and PA28beta. Embo J 2001, 20, 5898–5907. MURATA, S. et al. Growth retardation in mice lacking the proteasome activator PA28gamma. J Biol Chem 1999, 274, 38211–38215. WHITBY, F. G. et al. Structural basis for the activation of 20 S proteasomes by 11 S regulators. Nature 2000, 408, 115–120. USTRELL, V., HOFFMAN, L., PRATT, G., and RECHSTEINER, M. PA200, a nuclear proteasome activator involved in DNA repair. EMBO J. 2002, 21, 3403–3412. HO, Y., GRUHLER, A., HEILBUT, A., BADER, G. D., and MOORE, L. Systematic identification of protein complexes in Saccharomyces cerevisiaex by mass spectrometry. Nature 2002, 415, 180–183. GAVIN, A. C. et al. Functional organization of the yeast proteome by systematic analysis of protein complexes. Nature 2002, 415, 141–147. LEGGETT, D. S. et al. Multiple associated proteins regulate proteasome structure and function. Mol Cell 2002, 10, 495–507. MCCUTCHEN-MALONEY, S. L. et al. cDNA cloning, expression, and functional characterization of PI31, a proline-rich inhibitor of the proteasome. J Biol Chem 2000, 275, 18557–18565. ZAISS, D. M., STANDERA, S., KLOETZEL, P. M., and SIJTS, A. J. PI31 is a modulator of proteasome formation and antigen processing. Proc Natl Acad Sci USA 2002, 99, 14344–14349. LU, D. C. et al. A second cytotoxic proteolytic peptide derived from amyloid beta-protein precursor. Nat. Medicine 2000, 6, 397–404.

27

28 The 26S Proteasome

163 GACZYNSKA, M., OSMULSKI, P. A., GAO, Y., POST, M. J., and SIMONS, M. Prolineand arginine-rich peptides constitute a novel class of allosteric inhibitors of proteasome activity. Biochemistry 2003, 42, 8663–8670. 164 YAMANO, T. et al. Two distinct pathways mediated by PA28 and hsp90 in major histocompatibility complex class I antigen processing. J Exp Med 2002, 196, 185–196. 165 APCHER, G. S. et al. Human immunodeficiency virus-1 Tat protein interacts with distinct proteasomal alpha and beta subunits. FEBS Lett 2003, 553, 200–204. 166 HUANG, X. et al. The RTP site shared by the HIV-1 Tat protein and the 11S regulator subunit alpha is crucial for their effects on proteasome function including antigen processing. J Mol Biol 2002, 323, 771–782. 167 FAROUT, L. et al. Distribution of proteasomes and of the five proteolytic activities in rat tissues. Arch Biochem Biophys 2000, 374, 207–212. 168 BROOKS, P. et al. Subcellular localization of proteasomes and their regulatory

169

170

171

172

173

174

complexes in mammalian cells. Biochem. J. 2000, 346, 155–161. REITS, E. A., BENHAM, A. M., PLOUGASTE, B., NEEFJES, J., and TROWSDALE, J. Dynamics of proteasome distribution in living cells. EMBO J. 1997, 16, 6087–6094. ARABI, A., RUSTUM, C., HALLBERG, E., and WRIGHT, A. P. Accumulation of c-Myc and proteasomes at the nucleoli of cells containing elevated c-Myc protein levels. J Cell Sci 2003, 116, 1707–1717. KOPITO, R. R. Aggresomes, inclusion bodies and protein aggregation. Trends Cell Biol 2000, 10, 524–530. WOJCIK, C. and DEMARTINO, G. N. Intracellular localization of proteasomes. Int J Biochem Cell Biol 2003, 35, 579–589. HAMPTON, R. Y. ER-associated degradation in protein quality control and cellular regulation. Curr Opin Cell Biol 2002, 14, 476–482. MCCRACKEN, A. A. and BRODSKY, J. L. Evolving questions and paradigm shifts in endoplasmic-reticulum-associated degradation (ERAD). Bioessays 2003, 25, 868–877.

1

Molecular Machines for Protein Degradation Matthias Bochtler

Warszawa, Poland

Michael Groll Ludwig-Maximilians-Universit¨at M¨unchen, M¨unchen, Germany

Hans Brandstetter Max-Planck-Institute for Biochemistry, Martinsried, Germany

Tim Clausen Institute for Molecular Pathology, Vienna, Austria

Robert Huber Max-Planck-Institute for Biochemistry, Martinsried, Germany

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8

1 Introduction

The action of intracellular proteolytic enzymes is tightly controlled to avoid destruction of properly folded and functional proteins essential for cell viability and to restrict their activity towards sick molecules or/and those marked for destruction. The four (ﬁve) proteases discussed in this chapter display different regulatory mechanisms but show sequestration of their active sites inside molecular cages as a common structural principle, albeit being assembled from different building blocks in different shapes and with varying symmetries. The chapter focuses on structural, functional, and mutational studies from our laboratory. We are aware of other cage-forming proteases and their regulatory components that have been structurally characterized and which are mentioned in brief later in the context of our studies. The chapter is arranged in four main sections focused on the proteases HslVU, proteasome, tricorn (DPPIV), and DegP.

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Molecular Machines for Protein Degradation

2 The ATP-dependent Protease HslVU

ATP-dependent proteases are complex proteolytic machines. They are present in eubacteria, archaebacteria, in eukaryotic organelles and, as the 20S or 26S proteasome, in the eukaryotic cytosol and nucleoplasm. The activators of all known ATP-dependent proteases are related. They all contain an AAA(+) ATPase domain as a module [1] and are thought to assemble into hexameric particles or, in the case of 26S proteasomes, are present in six variants in the 19S activators [2]. Like the ATPases, the proteolytic components of the ATP-dependent proteases form higher order complexes, but unlike for the ATPases, the symmetry of the protease assemblies varies, and the folds of the subunits need not be related. ClpP is a serine protease, FtsH a metalloprotease, and HslV and the proteasomes from archaebacteria and eubacteria are threonine proteases. Although extensive biochemical data on both the bacterial and eukaryotic ATPdependent proteases are available, the characterization of these proteolytic machines at atomic resolution has proven difﬁcult, because of both the large size of these complexes and their lability to proteolysis and dissociation. No structural data at all are currently available for Lon and the mitochondrial ATP-dependent proteases. In the case of the cytosolic, membrane-integrated bacterial protease FtsH, atomic resolution data are available only for the ATPase domain [3, 4]. In contrast, the ATP-dependent activators of the ClpAP and ClpXP proteolytic machines have so far resisted crystallization. Atomic resolution data are available only for the proteolytic component ClpP [5], and separately for a ClpX monomer [6] and a ClpA monomer [7]. The bacterial protease HslVU is unique in two respects: at present, it is the only ATP-dependent protease to have atomic coordinates of the full complex determined; secondly, and in contrast to all other bacterial ATP-dependent proteases, it contains a proteolytic core that is related to the 20S proteolytic core of archaebacterial and eukaryotic proteasomes. The following sections summarize our understanding of HslVU biochemistry, crystallography, and enzymology and end with some speculation on the implications of these results for other ATP-dependent proteases. 2.1 HslVU Physiology and Biochemistry

In E. coli and most, but not all, other bacteria, the HslU (ATPase) and HslV (protease) genes are found in one operon under the control of a heat-shock promoter. The operon was ﬁrst found [8] and sequenced [9] in the course of a search for new heat-shock genes. It was later independently isolated again in screens for proteins that could down-regulate the heat-shock response [10] and for suppressors of the SOS-mediated inhibition of cell division in E. coli [11]. The observed biological responses in HslVU over-expression or deletion strains result from a decrease or increase in the steady levels of HslVU substrates [12]. HslVU, itself a heat-shock protein, affects the heat-shock response by degradation of the heat-shock factor σ 32

2 The ATP-dependent Protease HslVU

[10, 12] and the SOS response via the degradation of the cell-division inhibitor SulA [13, 14]. The physiological role of HslVU seems to be limited, probably because of overlapping substrate speciﬁcity with other ATP-dependent proteases in bacteria. The E. coli HslVU deletion strain has no phenotype at standard growth temperature, and it appears that HslVU is required for normal growth only at very high temperatures [12]. According to the protease database MEROPS, some bacterial species appear to lack an HslVU–type peptidase altogether [15]. Therefore, it came as a surprise that some HslV and HslU homologs were recently found in primordial eukaryotes where they appear to be simultaneously present with genuine 20S proteasomes [16]. Low expression levels and the lability of the HslVU complex make work with proteins from wild-type strains difﬁcult. Gratifyingly, the active protease can be reconstituted in vitro from over-expressed and puriﬁed components [17]. It requires ATP for the degradation of folded substrates and ATP or some of its analogs for the puriﬁcation of small chromogenic peptides. As expected, ATP-hydrolysis and proteolysis activities are mutually dependent [18]. In addition, the peptidase activity was found to depend in complex ways on the presence of various cations, especially K+ in the buffers [19]. 2.2 HslV Peptidase

On the sequence level, HslV shows sequence similarity with the β-subunits of arch-aebacterial and eukaryotic proteasomes, a fact that was immediately noticed when the E. coli gene was sequenced [9] and was later shown to extend to other related eubacterial sequences [20]. Electron microscopy (EM) of recombinant HslV subsequently suggested that the particle formed a dimer of hexamers that appeared to enclose only one central cavity without antechambers as in proteasomes [21]. The unexpected six-fold symmetry of HslV and the similarity in subunit fold with eukaryotic proteasomes were subsequently conﬁrmed by X-ray crystallography [8]. The crystallographic data also showed that the contracted ring compared to proteasomes resulted from small changes to the subunit–subunit interface, not from an entirely new mode of oligomerization [8] (see Figure 1). All 12 active sites of HslV are located on the inner walls of the hollow particle. In the E. coli particle, each active site has neighboring active sites 28 Å away on the same ring and 22 Å and 26 Å away on the opposite ring. The environment of the nucleophilic Thr1 looks similar to that in proteasomes, and the presence of a (putatively protonated) lysine residue near the active site probably helps to lower the pK a of the N-terminal α-amino group so that it is present in the unprotonated form, which can act as the general base to accept a proton from Thr1. Since the determination of the HslV crystal structure, two additional crystal structures from other species have been determined. The highest resolution structure available to date is the crystal structure of the Haemophilus inﬂuenzae enzyme [23]. Intriguingly, this structure showed the presence of cation-binding sites near the active centers [23], a ﬁnding that could subsequently be conﬁrmed for the

3

4 Molecular Machines for Protein Degradation

Fig. 1 E. coli HslV vs. T. acidophilum proteasome. Superposition of one hexameric ring of E. coli HslV (red) and of one heptameric ring of T. acidophilum proteosome $-subunits (green) in stereo representation.

The subunits at the “top” of the ring have been overlayed optimally. Note that the “tails” of the HslV subunits that point radially outwards are histidine tags and thus cloning artefacts.

Thermotoga maritima enzyme [24] and that explains, at least in qualitative terms, the dependence of HslVU activity on various cations in solution. Overall, the crystal structures of the enzymes from H. inﬂuenzae [23] and T. maritima [24] are very similar to the original structure of the E. coli enzyme. Therefore, it came as a surprise that the HslV homolog known as CodW from Bacillus subtilis behaves rather differently. Although the enzyme contains a threonine residue that aligns with the active site threonine of the E. coli, H. inﬂuenzae, and T. maritima enzymes, it does not contain the glycine at the C-terminus of the profragment that is believed to be required for efﬁcient autocatalytic processing, and indeed the polypeptide chain is processed ﬁve residues upstream of the conserved threonine that is the active-site nucleophile in other species to expose an N-terminal serine residue [25]. Whether this implies that the serine is in the spatial position normally ﬁlled by threonine, implying a discrepancy between the sequence-based and structure-based alignments, or whether it means that the accessory catalytic residues are either dispensable or anchored elsewhere on the sequence is currently not clear. 2.3 HslU ATPase

Based on the sequence, HslU can be easily classiﬁed as an ATPase by the presence of conventional Walker A (phosphate binding loop or P-loop) and Walker B (magnesium binding) motifs. Beyond this simple classiﬁcation, two competing models for HslU were proposed, classifying the enzyme either as a PDZ-domain

2 The ATP-dependent Protease HslVU

containing ATPase [26] or alternatively as a AAA(+)-type ATPase [1]. The crystal structure settled the issue in favour of the AAA(+) model [9]. AAA(+)-ATPases consist essentially of two structural domains that are connected through a short linker. A nucleotide binds at the interface of the two domains. As ﬁrst observed with HslU, the presence or absence of a nucleotide induces different relative orientations between the two domains [9]. With the availability of many different nucleotide states of HslU, the model was later reﬁned to include a dependence on the state of hydrolysis of the nucleotide [28]. The nucleotide is in a strategic position both at the interface of the N- and C-domains of one subunit and at the interface of adjacent subunits. A combination of mostly conserved residues from the two subunits around the nucleotide creates a highly polar environment [9]. Two arginine residues have attracted particular attention: R393 of E. coli HslU is thought to act as the “sensor” that transmits information on the presence or absence of nucleotide, and possibly on its identity, to the C-domain and thus controls the relative orientation of N- and C-domains in HslU. Another conserved arginine residue, R325, is anchored on the subunit that makes fewer contacts with the nucleotide and is the homolog of the proposed “arginine ﬁnger” in FtsH [29]. Although the term “arginine ﬁnger” (taken from small GTP-binding proteins Ras and Rho) implies a direct catalytic role for this residue, its distance from the nucleotide phosphates argues more for an indirect role. A similar conclusion has since been reached for ClpA [7]). Experimentally, mutation of either of the two arginine residues abolishes all ATP-dependent proteolysis activity [30]. Loss of subunit interactions plays a major role in the loss of function: The “arginine sensor” mutant R325E is fully and the “arginine ﬁnger” mutant R393 is partially dissociated in gel-ﬁltration experiments in the presence of salt [30]. A very complex picture has emerged from biochemical and crystallographic studies designed to characterize the substrate-binding sites in HslU. An essential role for the C-terminus of HslU was ﬁrst suggested on the basis of experimental studies that were designed based on the prediction of PDZ-like domains at the C-terminus of HslU [26]. Although the prediction of PDZ-like domains later turned out to be in error, the conclusion about the role of the C-terminus of HslU in substrate recognition was later corroborated with the deﬁnition of a biochemically deﬁned sensor and substrate discrimination domain (SSD) [31] that is also present in other AAA(+) ATPases and was suggested to act as the “hook” for substrates [32]. When the crystal structure of HslU became available, the SSD domain turned out to coincide with its C-domain. This ﬁnding is remarkable and not fully understood, since in all crystal structures of HslU the C-domain primarily mediates oligomerization contacts between HslU subunits. Its solvent-accessible regions are found far on the periphery of the HslU ring, far outside the cavity that is formed by the protruding I-domains (see Figure 2). From the crystal structure [9], it would appear likely that the protruding I-domains rather than the SSD domains act as the “hook”, although the ill-deﬁned tertiary structure of the I-domains makes speciﬁc interactions unlikely (see Figure 2). Consistent with this model, it was found experimentally that the I-domains are essential for the degradation of the folded substrate MBP-sulA [30]. A recent two-hybrid study is consistent with both points of view. It conﬁrms

5

6 Molecular Machines for Protein Degradation

Fig. 2 HslU surface colored according to domain. (A) View along the six-fold axis, seen from the side opposite to the I-domains. (B) View along the six-fold axis, seen from the side of the I-domains. Every second subunit of the ring is colored according to domain (N-domain yellow, I-domain blue, C-domain or SSD-domain red), the other subunits are colored in green. The diagram is based on the trig-

onal crystals of E. coli HslU that contain nucleotide in every other subunit. This asymmetry and crystallographic packing effects are responsible for the broken six-fold symmetry of the I-domains. Note that the I-domains of three subunits at the top of the ﬁgure have been cut away in (B) to allow a view on the globular N- and C-domains.

the essential role of the C-domain (SSD-domain) in oligomerization, but also supports a role for the I-domain and the SSD-domain in substrate degradation [33]. Presumably, if substrates are translocated through the central pore in HslU as EM data suggest for ClpXP [34], both the I-domains and the globular part of the ring would come into contact with substrate during substrate translocation, although the location of the C-domains (SSD-domains) on the periphery of the HslU ring then still needs to be reconciled with this model. The precise mode of recognition of substrates is even less clear for CodX, the HslU homolog from B. subtilis. In the absence of detergent, the I-domains of two hexameric CodX rings contact each other, leading to a head-to-head stacking of CodX rings and presumably the formation of a central cavity loosely surrounded by I-domains. As the dimer of CodX rings can associate with CodW protease on either side, repetitive, chain-type structures with alternating double rings of the peptidase CodW and the ATPase CodX can be formed [35]. The physiological signiﬁcance of these high molecular weight assemblies is currently not clear. 2.4 The HslVU–Protease Complex

Over the years, a key theme in ATP-dependent proteolysis has been the issue of “symmetry-matched” vs. “symmetry-mismatched” complexes. In the light of the

2 The ATP-dependent Protease HslVU

clearly established symmetry mismatch of the ClpAP [36] and ClpXP [37] complexes, the very clear EM data on the six-fold symmetry of HslV [21, 38] and reports about a predominant species of HslU with six-fold symmetry [21, 38] came as a surprise because they implied that a “ratcheting mechanism” of ATP-dependent proteolysis, if it existed at all, could not be operating in the HslVU system. This conclusion has since been conﬁrmed by all HslU and HslVU crystal structures [9, 39–41]. In all cases, HslU is hexameric and matches the oligomerization state of HslV. For the ﬁrst crystal structure of an HslU-HslV co-crystal, a controversial I-domainmediated contact between HslU and HslV was reported [10, 40, 43]. The contact was suspicious from the very beginning because of poor contact area, but seemed compatible with the known low afﬁnity between HslV and HslU and appeared to explain how the symmetry mismatched ClpXP and ClpAP complexes could be formed. Although a crystallographic reinterpretation of our original data that attributed this docking mode to overlooked twinning [40, 43] turned out to be itself in error [10], it is now clear from the combined results of cryoelectron microscopy [44], small-angle scattering [39] and several additional crystal structures of the complex [39, 40] that the physiological mode of interaction between HslU and HslV is with HslU I-domains distal to HslV (see Figure 3). 2.4.1 Allosteric Activation In the absence of a nucleotide, HslVU has residual activity at best, but the presence of several non-hydrolyzable ATP-analogs is sufﬁcient to stimulate HslVU–driven proteolysis activity against substrates that do not require unfolding, suggesting an allosteric effect of nucleotide on HslU and via HslU on HslV. This was further corroborated by the observation that a peptide vinyl sulfone formed a covalent complex with HslV only in the presence of HslU and a nucleotide [12]. The details of this allosteric mechanism emerged from the crystal structure of HslVU from H. inﬂuenzae. In this case, but not in other crystal structures of the HslVU complex, the normally buried C-termini of HslU distend and insert into active-site clefts in HslV to reach out almost to the HslV active centers [39] (see Figure 4). The crystal structure of HslVU in complex with a peptide vinyl sulfone inhibitor [46] and two independent biochemical studies [47, 48] that demonstrated the activatory properties of the C-terminal tails of HslU further corroborated this mechanism. In the light of these data, it is remarkable that wild-type HslU in the presence of ADP does not act as an activator for HslV, not even against unfolded or chromogenic substrates. A possible, but experimentally untested explanation could be that the C-termini of HslU are available for HslV binding only in the presence of activatory nucleotides. Whatever the details of the allosteric activation mechanism, it is already clear that HslU affects primarily the conformation of HslV active sites and not the accessibility of the HslV proteolytic chamber. Two independent lines of in vitro evidence support this conclusion. Firstly, an HslV mutant with a widened entrance channel does not show increased proteolytic activity in the absence of HslU, although it can still be activated like wild-type HslV by the presence of HslU [47]. Secondly, the crystal structure of the H. inﬂuenzae asymmetric HslVU protease in complex with an

7

8 Molecular Machines for Protein Degradation

Fig. 3 HslVU complex. Originally reported (A) and physiologically relevant (B) docking mode between HslV and HslU. In the physiologically relevant docking mode, the I-domains of HslU point away from the HslV.

inhibitory peptide vinyl sulfone has the inhibitor bound only in HslV subunits that are in contact with HslU [41], strongly arguing against accessibility of the proteolytic chamber as the rate-limiting factor at least under the experimental conditions. 2.5 A Comparison of HslVU with ClpXP and ClpAP

The protease core particles HslV and ClpP are assembled from subunits of entirely different fold and catalytic mechanism. ClpP is a serine protease that belongs to the crotonase superfamily of enzymes, a large class of enzymes that catalyze a variety of chemical reactions that all require the stabilization of an intermediate by an oxyanion hole [3]. HslV belongs to the family of Ntn-hydrolases that share the fold and use the N-terminal residue as the nucleophile [15]. Both ClpP [5] and HslV

2 The ATP-dependent Protease HslVU

Fig. 4 HslVU activation mechanism. (A) Stereo view (C" -trace) of the superposition of the C-domain of an HslU subunit (red) from the original E. coli HslVU complex onto that of the H. inﬂuenzae HslU subunit (green) from its complex. Two HslV subunits (pink and blue) from the H. inﬂuenzae complex are also shown to illustrate the binding of the C-terminal segment of H. inﬂuenzae HslU to the pocket between the HslV subunits (indicated also by a black

curved arrow). (B) Stereo view of the closeup of the C-terminal residues of an E. coli HslU subunit (red) from the complex. The carboxylate of the terminal leucine residue forms salt bridges with R394 of the same subunit and with R329 of an adjacent (yellow) HslU subunit that is not illustrated in (A) for clarity. (C) Stereo view of the closeup of the C-terminal residues of an HslU subunit from the H. inﬂuenzae HslVU complex.

9

10 Molecular Machines for Protein Degradation

subunits assemble into large oligomers that enclose a central proteolytic chamber, although the symmetry is different, since HslV is a dimer of hexamers and ClpP is a dimer of heptamers. In contrast to the protease components, which have varying symmetry, all known Clp ATPases are assembled from six identical subunits. These subunits contain either one (HslU and ClpX) or two (ClpA) copies of the AAA(+) module. In addition, the Clp ATPases contain additional domains that are unique for each ATPase. In HslU, a mostly helical I-domain is inserted into the AAA(+) module. ClpX contains an N-terminal domain that was shown to bind zinc [4] and act as a dimerization module [24]. Based on the latter result, a model of ClpX as a trimer of dimers was proposed, with N- and C-domains of ClpX forming a regular hexamer and the zinc-binding modules pairing into dimers [24]. In this context, it is remarkable that freshly isolated HslU behaves as a hexamer, but migrates with the apparent molecular weight of a dimer or trimer in gel ﬁltration after a freeze–thaw cycle [7]. Like ClpX, ClpA contains a non-AAA(+) domain at its N-terminus, and like the N-terminal domain in ClpX, this domain also has the capability to bind zinc [47]. However, unlike the N-domain of ClpX, the N-domain of ClpA is almost entirely helical and consists of two four-helix tandem motifs. In both ClpA and ClpX, the N-terminal non-AAA(+) motifs serve as “docking modules” for accessory proteolysis factors that modulate or change the activity of the proteolytic complex itself [25]. The N-domain of ClpX interacts speciﬁcally with the adapter protein SspB that stimulates the degradation of SsrA-tagged proteins [27]. The tag is jointly recognized by the SspB-ClpX complex, where SspB interacts with the N-terminal and central region of the SsrA tag and leaves the C-terminal region for interaction with ClpX [57, 58]. The C-terminal region of SspB shares considerable homology with the corresponding region in RssB, another ClpX adapter protein [27]. It appears that RssB promotes the degradation of a speciﬁc substrate, namely a subunit of RNA polymerase known as σ S [59]. It has been shown biochemically that in this case again the adapter protein and the ATPase recognize distinct sites in the substrate [60]. ClpA has its own adapter protein, ClpS. At least in vitro, ClpS switches ClpAP activity away from SsrA tagged towards heat-aggregated proteins [26]. The independent crystal structures of ClpS in complex with the N-domain of ClpA are available [47, 62]. They explain the speciﬁcity of ClpS for ClpA over other related Clp proteins, especially ClpB [62]. HslU lacks a domain upstream of the AAA(+) module. Consistent with this, no adapter proteins for HslU have been found so far, to the best of our knowledge. Mechanistically, there are important differences between the protease activation mechanisms of ClpXP/AP and HslVU. Most importantly, HslVU is a symmetrymatched proteolytic machine. In contrast, the deﬁnitive seven-fold symmetry of the ClpP protease and the well-established six-fold symmetry of ClpA and ClpX imply that ClpXP is a symmetry-mismatched system. It is hard to imagine how such an arrangement would be compatible with an HslVU–style activation mechanism with insertion of the C-termini of all activator subunits into clefts in protease. Moreover, the C-termini of ClpX particles from various species are very poorly conserved, arguing against their involvement in any allosteric activation mechanism [47].

2 The ATP-dependent Protease HslVU

Experimental evidence implicates an internal loop of ClpX in ClpP binding [63]. This loop is required for ClpXP proteolytic activity and may well be the functional equivalent of the C-terminus of HslU. If so, then the symmetry mismatch in ClpXP would suggest that only a subset of ClpX loops could insert into ClpP clefts at any given time. 2.6 HslVU Peptidase as a Model for the Eukaryotic 26S Proteasome?

On the sequence level, HslV shows sequence similarity with the β-subunits of arch-aebacterial and eukaryotic proteasomes. The crystal structure of E. coli HslV conﬁrmed that individual subunits share the Ntn-hydrolase fold with Thr1 at the N-terminus as the nucleophile, just as in proteasomes. Despite these similarities, there are substantial differences between bacterial HslVU and archaebacterial and eukaryotic 20S proteasomes. In contrast to HslVU, 20S proteasomes are assembled from four rings of seven subunits each, that build up a central proteolytic chamber and two ﬂanking antechambers. The essential role of the C-terminus of HslU has its direct counterpart in the essential role of the C-terminus of the ATP-dependent proteasome activator PA28 [64]. A complex of the yeast 20S proteasome with PA26, the Trypanosoma brucei homolog of PA28 has been crystallized and shows that the C-termini of PA28 insert into clefts in the 20S core particle [65], leading to an opening of the gates in the antechambers. So far, there is no evidence for allosteric activation in the 20S proteasome-PA26 complex or the 26S proteasome, where channel “gating” appears to be important as discussed below in Section 3 on the yeast 20S proteaseome. Currently, high-resolution EM image reconstructions for the 26S proteasome [66], but no atomic-resolution crystallographic data are available for any complex of 20S proteasomes with ATP-dependent activators. The expected assembly of PAN, the archaebacterial AAA(+) activator of proteasomes [67] into hexamers suggests a symmetry-mismatched complex in archaebacteria. The ATP-dependent proteasome activators of eukaryotic proteasomes known as PA700 and the 19S cap also contain six AAA(+) ATPases in a subcomplex of the 19S complex known as the “base” [2]. A priori, one would expect the six ATPases to form a ring with pseudo six-fold symmetry similar to the six-fold ring seen in bacterial AAA(+) activators, but two-hybrid experiments have suggested alternative models [68]. It is currently not clear whether the C-termini of the AAA(+) ATPases in the 19S cap play a similar role as in the ATP-independent PA28-20S proteasome complex. There is no consensus in the C-terminal sequences of different proteasomal AAA(+) ATPases of any particular species, but consensus sequences for the C-termini of any particular subunit from different species can be deﬁned. Unfortunately, the overall sequence similarity of homologous sequences from different species is too high to infer a functional role of the AAA(+) C-termini from sequence similarity. As discussed above, the eubacterial HslVU is distantly related in structure to the proteasome found in archaea and eukaryotes. Surprisingly, however, the structural

11

12 Molecular Machines for Protein Degradation

relationship is not reﬂected in the regulatory properties as will be described in Section 3, which focuses on structural studies of the yeast 20S proteasome and its activation, activity, and inhibition.

3 The Yeast 20S Proteasome

The most elaborate version of the proteasome core particle (CP) is found in eukaryotes as shown by the crystal structure of the yeast 20S proteasome [40] (see Figure 5B). Here, the α- and β-subunits have diverged into seven different subunits each as compared to the archaeal enzyme which mostly consists of two components [70]. The subunits are present in two copies and occupy precisely deﬁned positions within the 20S complex. As in the archaeal proteasome, the α-subunits are inactive and contribute to the antechambers of the particle, whereas the β-subunits set up the inner hydrolytic chamber. Remarkably, four of the seven different eukaryotic β-subunits lack residues that are essential for pro-peptide autolysis and are therefore proteolytically inactive. As in the archaeal CP, the remaining three subunits mature autoproteolytically to active threonine proteases but with a caspase-like (β1), trypsin-like (β2), and chymotrypsin-like (β5) activity, they exhibit different cleavage potentials [40]. The crystal structure of the bovine 20S proteasome [71] demonstrated that yeast and mammalian CPs are highly homologous in their structural architecture, quaternary assembly, and active-site geometry. However, in mammalian cells, three additional non-essential subunits, β1i, β2i and β5i, respectively, can replace their constitutive counterparts upon induction by the cytokine γ -interferon. The interchange of active subunits modiﬁes the CPs peptidase speciﬁcity and is important for the function of the immunoproteasome. The speciﬁcity of the S1 pockets of the induced subunits increases the yield of peptides favored for binding to MHC class I molecules and antigen presentation. The bovine proteasome structure provides an explanation of how the constitutive and inducible β-subunits can be mutually interchanged. In the yeast proteasome, these subunits are held in place by several speciﬁc interactions, which are absent in the mammalian homolog [71]. 3.1 The Proteasome, a Threonine Protease

As mentioned in Section 3, proteasomes are threonine proteases. Accordingly, a proton acceptor is required to activate the hydroxylic group of Thr1. Although there are several potential acidbase catalysts around Thr1, activation seems to proceed by its own terminal amino group [2, 41]. A positively charged side chain of Lys33 lowers the pK a of the Thr1 amino group (see Figure 6). The assignment of the N-terminus as the catalytic base in proteolysis is further supported by the fact that all Ntnhydrolases share a common fold, but generally do not display any kind of active-site consensus. A second essential factor for proteolysis is a catalytic water molecule that has been observed in the high-resolution structures of CPs [23, 40, 46, 71]. The

3 The Yeast 20S Proteasome

Fig. 5 Molecular surface of the archaeal (A), the eukaryotic 20S (B) and the HslV proteasome (C). The accessible surface is colored in blue, the clipped surface (along the cylinder axis) in white. To mark the position of the active sites, the complexes are shown with the bound inhibitor calpain (yellow). (A) The disorder of the ﬁrst Nterminal residues in the archaeal "-subunits generates a channel in the structure of the CP, (B) whereas the asymmetric but welldeﬁned arrangement of the " N-terminal tails seals the chamber in eukaryotic CPs. (C) The eubacterial “miniproteasome” has an open channel through which unfolded proteins and small peptides can access the proteolytic sites. (D) Ribbon plot of the free "-ring from A.fulgidus focusing on

the deﬁned N-termini (red). Tyr8 of each N-terminal part makes hydrogen bonds to Asp9 of the adjacent "-subunit (yellow), Arg10 (red) points toward the channel, generating a 13-Å entrance. The ﬁnal 2FO -FC electron density map, contoured at 1F is shown for the YDR-motif (E, F) Electron density maps of the yeast core particle from wild type and "3N mutant, respectively The individual N-terminal tails of the "subunits are drawn in different colours. Asp9 of subunit "3 plays a key role in stabilizing the closed state ofthe channel and is marked with a black arrow. In the "3N mutant, an open axial channel is visible, whose dimensions are comparable to those ofthe archaeal CP channel.

13

14 Molecular Machines for Protein Degradation

Fig. 6 Proteolytic site of the yeast $1 subunit. The protein backbone is drawn as a white coil with the active-site residues Thr1, Asp17, Lys33, Ser129, Asp166 and Ser169 shown in ball-and-stick mode. Owing to the salt-bridge with Asp17, the amino group of Lys33 should be positively charged and thus

be able to lower the pK a of Thr1O( electrostatically. The other active-site residues Ser129, Asp166 and Ser169 deﬁne the orientation of Thr1. A water molecule (green sphere) is located properly to shuttle protons between the terminal amino group and Thr1O( during proteolysis.

solvent molecule is ideally positioned to shuttle protons between the Thr1Oγ and the N-terminus. Furthermore it could act as the base for the cleavage of the acyl ester intermediate, thereby releasing Thr1Oγ for the next catalytic cycle [23]. In general, the cleavage products of the proteasome vary in length between 3 and 25 amino acids with an average length of 7 to 8 amino acids. Several models have been suggested for the “molecular ruler” that determines fragment length. On the basis of structural studies of mutants unable to autolyse, we suggest that it is determined by the substrate-binding clefts designed for peptides 7–9 amino acids long [41]. The likelihood of substrate cleavage depends on the mean residence time at the proteolytic sites, which is maximal if all binding sites are ﬁlled [22, 77]. The active subunits in eukaryotic CPs differ mainly in their binding pockets, yielding different cleavage speciﬁcities. However, it must be emphasized that the proteasome complex does not represent a simple collection of chymotrypsin-like, trypsin-like, and caspase-like enzymes. In fact it is the structural architecture of the proteolytic chamber that determines speciﬁcity. The local structure around each active site imposes a physical constraint on the peptide substrates, whereas the selectivity of the S1 pockets is less relevant. The mechanistic importance of the inner chamber can also be seen from the fact that substrate residues other than P1 inﬂuence degradation [13, 17, 45] and that neighboring subunits interfere with

3 The Yeast 20S Proteasome

the functions of the catalytic subunits [41, 49, 53]. However, inhibitor binding and mutational studies indicate that allosteric interactions between individual subunits are insigniﬁcant [41, 45, 49, 53, 83] contrasting the allosteric activation of HslV by HslU described in the previous section. 3.2 Inhibiting the Proteasome

Proteasome inhibitors have been instrumental in identifying numerous protein substrates and in elucidating the importance of the proteasome/ubiquitin pathway in many biological processes. Initially, non-speciﬁc cell-penetrating peptide aldehydes were used for this purpose. More recently, it became possible to synthesize compounds with increased potency and selectivity [1, 30]. Furthermore, based on the crystal structure of the yeast and bovine liver CP [40, 71], molecular modeling can now be used to engineer improved inhibitors. Besides the synthetic inhibitors, a variety of natural compounds is known to inhibit the CP. One of these natural inhibitors, lactacystin, was discovered by its ability to induce neurite outgrowth in a murine neuroblastoma cell line. Incubation of cells in the presence of radioactive lactacystin leads to the labelling of the β5 subunit [32] and to irreversible inhibition of the CP. As shown by X-ray analysis, the inhibitor is covalently attached to subunit β5 by an ester bond with the N-terminal Thr1Oγ [40] (see Figure ??A). The subunit selectivity of lactacystin can be attributed to its dimethyl group, which mimics a valine or a leucine side chain and closely interacts with Met45 in the hydrophobic S1 pocket of subunit β5. Epoxomicin, an α ,β -epoxyketone peptide, is a natural compound that potently and irreversibly inhibits the catalytic activity of the CP [83]. Unlike most other proteasome inhibitors, epoxomicin is highly speciﬁc for the proteasome and does not inhibit any other protease. The crystal structure of epoxomicin bound to the yeast CP explained the unique selectivity of the inhibitor (see Figure ??B). Adduct formation yields an unexpected morpholino ring, which is formed between the Thr1Oγ , the N-terminus, and the epoxy group of the inhibitor [43]. However it should be noted that proteasome inhibitors that covalently bind to the active βsubunits, usually cause apoptosis and cell death [66]. They are therefore cytotoxic and thus may not be pharmaceutically relevant. Recently, it was shown that certain natural products from Apiospora montagnei, TMC-95s, block the proteolytic activity of the CP selectively and reversibly in the low nanomolar range [67, 69]. The TMC-95s represent a novel class of proteasome inhibitors consisting of modiﬁed amino acids, which form a heterocyclic ring system. The crystal structure of the yeast CP in complex with TMC-95A shows the inhibitor non-covalently bound to all active sites [44] (see Figure ??C). TMC-95A was anchored by several speciﬁc hydrogen bonds, which are formed with main-chain atoms and strictly conserved residues of the β-subunits. The structures of TMC95s contain a crosslink between a tyrosine and an oxoindol side chain, resulting in a strained conformation that ﬁts ideally to the CP active site. Thus the entropic penalty of binding is lower than for more-ﬂexible ligands, which in turn explains

15

16 Molecular Machines for Protein Degradation

3 The Yeast 20S Proteasome

the speciﬁcity and high afﬁnity of the TMC-95s. Modeling studies indicate that it is possible to generate a TMC95 scaffold with a variety of functional groups attached to target the proteasomal S1 and S3 pockets and generate subunit speciﬁcity [54, 78]. All agents that speciﬁcally inhibit the proteasome are potentially of great pharmacological interest [80]. As the CP plays a dominant role in generating antigenic peptides, which are subsequently bound by MHC I molecules, compounds that block this activity might serve as a basis for the development of immunosuppressive drugs. Attempts are being made to design synthetic proteasome inhibitors using the discussed natural inhibitors as lead structures. In addition, proteasomal inhibitors represent powerful tools in molecular biology and can be utilized to identify novel cellular roles of the proteasome. 3.3 Access to the Proteolytic Chamber

In the Thermoplasma acidophilum and Archaeoglobus fulgidus CP, two narrow entry ports of 13-Å diameter exist at both ends of the cylinder, which prevent folded proteins entering [46, 70] (see Figure 5A). Many archaebacteria, such as Methanococcus jannaschii, contain a gene named PAN (proteasome-activating nucleotidase), which is highly homologous to the six ATPases in the 19S-component of the eukaryotic 26S proteasome [67]. It was shown that PAN selectively stimulates the degradation of unfolded proteins, whereas the digestion of small peptides is not enhanced. The threading of speciﬁc protein substrates into the lumen of the CP requires the action of an ATPase. The corresponding translocation process catalyzed by PAN follows ATP-dependent unfolding [6, 86]. However, it still remains to be clariﬁed whether complex formation is a prerequisite for the cooperation between PAN and CP or whether the two systems work independently. In contrast to the archaeal CPs, the hydrolytic chamber of the eukaryotic 20S proteasome is tightly sealed (see Figure 5B). The N-termini of the α-subunits project down and across the axial pore and block the entrances by several layers of interdigitating side chains, which form ← Fig. 7 Inhibitor binding to individual active sites of the yeast 20S proteasome. The inhibitors lactacystin (A), epoxomicin (B) and TMC95A (C) are colored green and are shown in stereo mode together with their unbiased electron densities. The active-site Thr1 is highlighted in black. (A) Covalent binding of the Streptomyces metabolite lactacystin to the active site of $5. The S1 pockets of the active subunits $1 and $2 differ from that of $5 and are not suitably constructed to bind the inhibitor. As discussed in the text, Met45 (black), which is located at the bottom of the $5-S1 pocket, makes the difference for inhibitor binding.

(B) Covalent binding of the proteasome inhibitor epoxomycin to $2. The electron density reveals the presence of a unique six-membered ring. The morpholino derivative results from adduct formation between epoxomycin and the proteasomal Thr1O( and amino terminus (pink sticks) and explains the speciﬁcity of the inhibitor towards Ntn-hydrolases. (C) Noncovalent binding of the speciﬁc proteasome inhibitor TMC-95A from Apiospora montagnei to $2. TMC-95A binds near the proteolytic centre in all active subunits in the extended substrate binding site.

17

18 Molecular Machines for Protein Degradation

a lattice-like structure [40] (see Figure 5E). Thus activation of the eukaryotic CPs requires substantial structural rearrangements of the N-terminal tails to open the molecular gate. This regulatory principle has been conﬁrmed by a yeast CP mutant, in which the ﬁrst nine amino acids of subunit α3 were deleted (α3N-mutant) [42]. The α3-N-terminal tail was chosen for deletion because it traverses the pore of the CP and contacts all other N-termini that are involved in the structural organization of the plug [40, 71]. In the crystal structure of the mutant, open axial pores were observed that were equivalent in size to those seen in the archaebacterial CP (see Figure 5F). Several points of evidence indicate that opening of the gate is indeed essential for catalytic activation, as all proteolytically active sites are simultaneously activated and no signiﬁcant structural changes can be seen between mutant and wild-type CP excluding allosteric effects. Furthermore, addition of the synthetic α3-N-terminal peptide to the α3N mutant restores wild-type behavior. An alanine scan of this peptide revealed that α3-Asp9 is essential for stabilizing the closed state of the channel. This aspartate residue closely interacts with Tyr8 and Arg10 of the neighboring subunit α4 (see Figure 5E). The strict conservation of this YDR motif and of other α-N-terminal residues suggests a universal mechanism for opening gates in eukaryotic CPs that has been conserved during evolution. We suggest that binding of regulatory proteins to the CP triggers the rearrangement of the α3-tail and thus opens the gate. This notion was further conﬁrmed by the crystal structure of a 20S/11S heterologous complex between the yeast 20S-proteasome and the Trypanosoma 11Sregulator [65]. This approach was justiﬁed by the ability of 11S regulators to activate 20S proteasomes from widely divergent species. The structure of the chimeric complex showed that one cylindrical regulator was bound at each end of the 20S barrel structure. Unlike the uncomplexed proteasome, all of the seven α-subunit Nterminal tails extend away from the CP in the complex and project towards the pore of the regulator. This rearrangement provides access to the proteolytic chamber and is basically achieved by two features: Firstly, the 11S-“activation loops” impose a more stringent seven-fold symmetry on the CP thereby straightening out the asymmetrically oriented α-tails and removing them from the entrance/exit gates. Secondly, the high-afﬁnity binding between CP and 11S is accomplished by the C-terminal sequences of the regulator, which insert into pockets formed between the 20S α-subunits. The major contact is observed between the C-terminal mainchain carboxylate of the 11S regulator and the entry of an internal helix of the CP α-subunit. The strength of this interaction is ampliﬁed by the heptameric assembly of the 20S/11S complex [65]. Activation of the CP by the 19S-complex is also regulated by controlling access to the proteolytic chamber, but the gating mechanism differs from that seen in the 11S/20S-complex. The 19S RP consists of two subcomplexes termed lid and base [35]. The base appears to form a ring like structure, including six conserved ATPase subunits (Rpt1–6), and Rpn1 and Rpn2, which are located proximal to the CP’s α-ring. Mutation studies indicated that the ATPase domain of Rpt2 plays a major role in regulating peptidase activity and that the 19S RP opens the gate to the protease in an ATP-dependent manner [68]. No detailed structural information

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 19

is as yet available that could provide further insight into how the 19S RP controls proteasomal activity. In some but not all archaea the proteasome is accompanied by another large cage-forming protease, the tricorn protease. Tricorn functionally interacts with the proteasome by cleaving the proteasomal peptide products into smaller peptides, which are further degraded into single amino acids by associated factors. Structural and functional aspects of tricorn are described in Section 4. The unexpected relationship between tricorn and the eukaryotic dipeptidyl peptidase IV revealed by these structural studies is also discussed in brief.

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV

Each living cell is a complex system and needs to continuously clear unnecessary or defective components. Within this context, the importance of the proteasome is well established (see Section 2). It predominantly carries out the degradation of cytosolic proteins and generates peptides varying in length between 3 and 25 amino acids. In order to be useful resources to the cell, these products need to be further degraded to eventually yield single amino acids. In the model organism Thermoplasma acidophilum a proteolytic system has been identiﬁed that does indeed perform this processing [101]. Based on the crystal structure of the tricorn protease [14], we provide evidence of how the tricorn protease accomplishes efﬁcient turnover of the proteasome-generated peptides. The structure of tricorn reveals a complex mosaic protein whereby ﬁve domains combine to form one of six subunits, which further assemble to form the D3 symmetric core protein. The structure shows how the individual domains coordinate the speciﬁc steps of substrate processing, including channeling of both the substrate to and the product from the catalytic site. Moreover, the structure shows how accessory protein components might additionally contribute to an even more complex protein machinery that efﬁciently collects the tricorn-released products. 4.1 Architecture of the Tricorn Protease

The hexameric D3-symmetric tricorn protein is assembled by two perfectly staggered and interdigitating trimeric rings with every subunit of one ring forming contacts almost exclusively with the two subunits of the other ring related by the molecular diads. The toroid structure has the shape of a distorted hexagon formed by a trimer of dimers (see Figure 8). The overall dimensions of the molecule are 160 Å within the plane normal to the three-fold axis and 88 Å parallel to it. The conically shaped central pore connects with additional cavities formed by the individual subunits like spokes of a wheel (see Figure 8). A single subunit is further divided into ﬁve sequential sub-domains, namely the N-terminal six-bladed

20 Molecular Machines for Protein Degradation

Fig. 8 Surface representation of the tricorn protease with the ribbon model of one subunit superimposed. The two orthogonal views are along the molecular two-fold and three-fold axis, respectively. The six solid spheres indicate the active-site positions.

β-propeller (β6) followed sequentially by a seven-bladed β-propeller (β7). Both β6 and β7 are topologically unclosed, an extremely rare feature observed only in the prolyl oligopeptidase (POP) [33] and DPIV protease [31, 96]. A PDZ-like domain (R761-D855) is interspersed between the two C-terminal mixed α-β domains. These C-terminal domains harbor the catalytic residues and exhibit the α-β hydrolase fold again underlining the relationship of tricorn with DPIV and POP. 4.2 Catalytic Residues and Mechanism

To elucidate the amino acids crucial for its catalytic activity, we have co-crystallized tricorn with a series of chloromethyl ketone-based inhibitors, including TLCK and TPCK for which we have conﬁrmed inhibitory efﬁcacy. For all of these inhibitors, we have observed continuous electron density connecting to the side chain of S965 which was unambiguously ﬁtted by the respective inhibitor. S965 is positioned at the entrance to helix H3 within sub-domain C2. The uncapped amino group of D966 forms, together with that of G918, the oxyanion hole, which is occupied by a water molecule in the uninhibited structure. H746 is ideally positioned to activate the catalytic S965 at a hydrogen-bonding distance of 2.7 Å. However, in none of the inhibitor complexes could we observe a covalent linkage between H746 and the inhibitor, as observed in the trypsin-like serine proteases [11]. Tricorn is related

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 21

to the cysteine proteinases in this respect [29]. We conﬁrmed that both residues are crucial for catalysis by constructing the single-site mutants S965A and H746A, both of which are amidolytically inactive. The H746 is correctly oriented by the Oγ of S745 which in turn is polarized by E1023. The arrangement of S965, H746, and the oxyanion hole suggests that the classical steps of peptide-bond hydrolysis follow the sequence of the trypsin-like serine proteases, namely the formation of the tetrahedral adduct, the acyl-enzyme complex, and hydrolysis. Tricorn has been shown to exhibit both tryptic and chymotryptic speciﬁcities [101]. The X-ray structure reveals that speciﬁcity for basic P1 residues is conferred by D936 which is provided by the diad-related subunit (see Figures 9 and 10). In this way, the previously described structural linkage (trimer of dimers) is translated into functional cooperativity within the dimers. Intriguingly, in the uninhibited high-resolution crystal structure, the acidic S1 speciﬁcity-determinant residue D936 was mobile. Consistent with this, the side chain of D936 in the TPCK complex structure adopts an alternative rotamer to allow the TPCK phenyl ring to freely access the hydrophobic niche formed by Y946, I969, V991, and F1013. D936 thus serves as a substrate-speciﬁcity switch accommodating both hydrophobic and basic P1 residues. The SO2 group of TPCK and TLCK interacts with the NH moiety of I994, thereby already suggesting the strand E993–P996 as the unprimedsubstrate docking site. These substrate-recognition sites are rather unrestricted in accord with tricorn’s broad substrate speciﬁcity [101, 108]. This situation is contrasted by the length restriction of the primed-substrate recognition site. A prominent cluster of basic residues (R131, R132) delineates the binding site of the substrate C-terminus. These basic residues, positioned on a ﬂexible loop as discussed in detail below, together with the primed-site topology, clearly mark tricorn as a carboxypeptidase. The geometric dimensions explain

Fig. 9 Stereo view of a 13-mer chloromethyl ketone bound to the active site. The electron density of the peptide directs to the $7 propeller.

22 Molecular Machines for Protein Degradation

Fig. 10 Detailed active-site view and substrate recognition as deduced from experimental complex structures. The substrate C-terminus is anchored by R131 and R132.

tricorn’s preferential di- and tri-carboxypeptidase activity, while the cleavage of longer peptides will require some conformational rearrangement and is energetically less favorable. By contrast, single amino acids cannot be cleaved off a substrate, because the P1 residue is unable to anchor its carboxylate-group on the basic backstop residues (see Figures 9 and 10). A negative charge was not tolerated at positions P3, P4, and P5 of a synthetic ﬂuorogenic AMC-substrate [108]. The crystal structure did not indicate any steric or electrostatic conﬂicts, if a canonical binding mode of these substrates was assumed. Owing to their lack of a free C-terminus, the charge polarity of N-terminally succinylated ﬂuorogenic substrates is inverted with respect to an unmodiﬁed peptide substrate and may lead to unproductive binding with inverted strand polarity. Each of the three C-terminal domains (C1, PDZ, C2) is remarkably similar to the respective domains (A, B, and C) found in the D1-processing protease (D1P) of photosystem II. The rms deviations between the Cα positions of these domains are 2.2, 2.3, and 2.7 Å with 84, 86, and 135 matching amino acids, respectively. A weak homology between these domains is recognizable in the primary sequences

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 23

(11, 19, and 20% identities). The relative arrangement of these domains, however, differs very much between tricorn and D1P. With the C2 domain aligned to the C domain of D1P, the orientation of the C1 domain differs from that of the D1P A-domain by 35◦ . Analogously, the required transformation to align the PDZ-like domains includes a 96◦ rotation. The rotation axes of these transformations are unrelated to each other. In addition, proper alignment of the PDZ domain requires a 30-Å translation. The catalytic serine residues (S965 and S372, respectively) are positioned on topologically equivalent positions at the helix entrance in the C2 (C) domain (D1P). Further, the amides forming the oxyanion hole (G918, D966, and G318, A373 in tricorn and D1P, respectively) superimpose to within 1 Å. As in other Tsp-like proteases, the residue serving as general base in D1P is a lysine (K372) residing within the C domain of D1P, while it is a histidine in tricorn (H746) which resides on tricorn’s C1 domain. The relative arrangement of the C1 and C2 domains in tricorn must, for that reason, remain very restricted to allow for proper catalysis. One role of the PDZ domain in substrate recognition has been shown for Tsp [5] and was analogously suggested for the tricorn protease [93]. While the GLGF substrate recognition element is structurally conserved (R764 IAC767 in tricorn), as pointed out earlier, it appears for a number of reasons unlikely that the tricorn PDZ will participate in substrate recognition in the same way as suggested for D1P [77]: (1) The putative substrate-binding site as deﬁned by the crystal structures of the C-terminal peptides complexed with PDZ domains [16, 28] is partly occupied by outer strands of blade 3 of β6 within the same subunit; (2) the generally conserved arginine (R247) involved in recognition of the carboxylate of the peptide C-terminus corresponds to a hydrophobic residue in tricorn (I851); (3) the orientation and position of tricorn PDZ differs so strongly from that seen in D1P that any analogy based on the sequential domain arrangement is invalidated on the basis of their respective three-dimensional domain arrangement. Instead, the PDZ domain mainly serves to scaffold the sub-domains as described earlier and, in addition, might be involved in recognition of associating component proteins.

4.3 Substrate Access and Product Egress Through β-propellers

The comparison with POP, including the open Velcro-topology [33], suggests an important role of the β-propellers for substrate access to and product exit from the active site [31] (see Figure 11). Both the β6 and β7 propeller axes are directed towards the active site of the protein, almost intersecting near S965. The arginine anchor (R131, R132) obstructs the otherwise direct connection from the active-site chamber to the exterior through the β6 propeller. Given these observations, we propose that the β6-propeller channel represents one, if not the major, rear exit from the catalytic chamber. This is consistent with the point mutation L184C, positioned within the β6 propeller. The introduced thiol group was modiﬁed with maleimide, partially blocking the β6 propeller. The activity of this mutant enzyme towards ﬂuorogenic

24 Molecular Machines for Protein Degradation

Fig. 11 Cartoon of the electrostatically driven processive substrate turnover.

substrates is signiﬁcantly reduced (<50%) compared with the wild-type protein [14, 64]. The substrate entrance and product exit paths are indicated in Figure 11. The chloromethyl ketone-based inhibitor-complex crystal structures suggested the strand E993–P996 as a recognition strand for the unprimed-substrate residues. This strand extrapolates towards the β7 channel (see Figure 9). The channel through the β7 propeller provides a signiﬁcantly shorter route from the catalytic chamber to the outside of the protein (60 Å) as compared to the alternative route through the central pore (83 Å). The latter path to the active site has multiple branchings and dead ends. Therefore, the β7 channel might be utilized by the enzyme for the preferred substrate passage to the active site. It is wide open but capped on its outside by four basic residues (R369, R414, R645, K646) which are only partially charge-compensated by one acidic residue (D456). This locally positive lid to the β7 propeller channel is encircled by acidic residues (D333, D335, D372, D456, D506, D508, E592, and E663). Except for E663, which is located on the hairpin connecting strand 3 and 4 of blade 7, all these charged amino acids are positioned between strands 1 and 2 of the respective β7 blades. The resulting charge distribution mimics an electrostatic lens, whereby peptides are pre-oriented with their C-termini towards the central basic propeller lid. Once the entrance to the β7 channel is opened by a concerted side-chain movement of R369, R414, R645, and K646, and possibly assisted by main-chain movements of A643–K646 (blade 7), a peptide is able to enter the channel in an extended conformation where it will ﬁnd multiple docking sites at the unsaturated inner strands of the β7 propeller blades. A similar substrate-gating ﬁlter mechanism through a seven-bladed β propeller has been suggested for the prolyl oligopeptidase [33, 34], and there is precedent for a β-hairpin binding into a seven-bladed propeller [52]. The preferred substrate entry through the β7 propeller channel is in line with the point mutation R414C, located in the β7 channel. Derivatization of this introduced thiol group with maleimide markedly reduced the ﬂuorogenic activity of this mutant to about 50% of the wild-type activity [14].

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 25

Tricorn cleaves substrates in a processive mode [64], indicating that only completely digested products will leave the inner protein chambers while larger products will be retained and processed as preferred substrates. The structure suggests several mechanisms to maintain “one way” processing. Basic lids (R414, R645, K646 and R131, R132) are present at the entrances to the β6 and β7 channels. The topology and size of the inner cavities favor an extended conformation of the substrate and the C-terminus of the substrate will be attracted to the basic β6 lid, thereby presenting the substrate’s scissile bond at the active site S965 for proteolyis. In one possible scenario, the primed product residues are released by the enzyme through the “rear exit” to the active site formed by the β6 propeller, which is gated by R131–R132. The arginine gate is located on a helical loop containing three glycines (G126, G130, G139) and not restrained to its position via any protein contacts. These glycines might function as hinge residues allowing the gate to move into a sufﬁciently voluminous cavity of mixed polarity (see Figure 10). The unprimed side of the substrate is held in place by a series of interactions with the protein. In addition to the observed ionic (D936) or hydrophobic S1 interaction site (Y946, I969, V991, F1013), the P1 main chain is held by its interaction with the oxyanion hole (G918, D966). P2–P4 residues will presumably utilize unsaturated main-chain hydrogen bonds at the strand I994–P996 (see Figure 10) and further interactions might occur in the β7 propeller channel as described in galactose oxidase [52]. The modeling studies and suggested substrate binding at the primed and unprimed sides are fully experimentally conﬁrmed by crystal-structural studies using C- and N-terminally extended covalently bound inhibitors [64]. Tricorn reportedly cooperates with three additional proteins, termed interacting factors F1, F2, and F3, to degrade oligopeptides sequentially to yield free amino acids [37] . F1 is a prolyl iminopeptidase with 14% sequence identity to the catalytic domain of prolyl oligopeptidase POP, which has an additional propeller domain [33, 37]. Guided by this structural scaffold of the latter structure, we speculate that F1 docks onto the six-bladed β-propeller of the tricorn core protein. As in POP, substrate would enter F1 through the propeller channel in this model. While a physical interaction of F1 with tricorn has been suggested [117], the exact mode of interaction of tricorn with F1, F2, and F3 has not been detailed so far. Similarly, there is evidence for functional but not physical interaction of tricorn with the proteasome [117]. A physical interaction between these molecules by aligning their respective central pores would imply a symmetry mismatch. While such a physical interaction would be consistent with the geometric dimensions of both molecules, its existence needs to be experimentally conﬁrmed and characterized. 4.4 Structural and Functional Relationship of Tricorn and DPIV

The situation in the tricorn protease is closely resembled by dipeptidyl peptidase IV (DPIV) where an eight-bladed topologically open β propeller and a side opening provide entrance to and exit from the active site (see Figure 12). Similar to tricorn, DPIV is a serine protease with low but signiﬁcant structural homology to the

26 Molecular Machines for Protein Degradation

Fig. 12 Ribbon representation of the tetrameric DPIV.

family of α/β-hydrolases. We superimposed the catalytic core elements, including the active-site serine and histidine, the strictly conserved helix following the active-site serine (Ser630-Ala642 and Ser965-Leu977, respectively), and tricorn’s ﬁve-stranded parallel β-sheet onto the equivalent strands of the eight-stranded DPIV-sheet. Both sheets have identical polarity. Signiﬁcantly, both tricorn propellers come to superimpose onto the two DPIV-openings, the tricorn β7 propeller onto the DPIV β8 propeller, and the tricorn β6 propeller onto the side exit, as schematically indicated in Figure 13. This similarity suggests that the β8 propeller provides substrate access to, and the side opening product release from, the DPIV active site. This tricorn-derived model is able to explain the high substrate selectivity critical for DPIV-function to activate or inactivate regulatory peptides. Passage through the β propeller tunnel requires the substrates to unfold thereby providing their “ﬁngerprint” to DPIV. Once the amino terminus of the peptide approaches the active site, it is still held in place by its C-terminus interacting with the β propeller which may contribute to conformationally activate the substrate for cleavage. After the nucleophilic attack the acyl-enzyme intermediate forms, while the primed product is directly released through the side exit. This explains why degradation of glucagon by DPIV is not processive, but occurs sequentially in two independent steps (glucagon 3–29, glucagon 5–29) [94]. Clearly, the ﬁnal determination of the functional roles of the DPIV openings awaits further experiments.

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 27

Fig. 13 Schematic representation of the active-site access and product egress in tricorn and DP IV.

4.5 The DegP Protease Chaperone: A Molecular Cage with Bouncers

In this section we describe DegP, a bacterial cage-forming protease, which has ho-mologs in all kingdoms of life. It is distinguished in essential ways from the previous systems by exhibiting extreme ﬂexibility and potential to change its overall shape and its internal structure. Structural ﬂexibility is translated into function and the unique property of DegP to act predominantly as a chaperone or as a protease dependent on temperature. DegP is a Janus-faced molecule appearing as helper or killer as cells need it. Cells have developed a sophisticated system of molecular chaperones and proteases to reduce the amount of unfolded or aggregated proteins [120]. Chaperones recognize hydrophobic stretches of polypeptides that become surface exposed as a consequence of misfolding or unfolding. If refolding attempts fail, irreversibly damaged polypeptides are removed by proteases. E. coli contains several intracellular proteases that recognize and degrade abnormally folded proteins. The biochemical and structural features of these ATPdependent proteases have been studied extensively (see Section 2). However, relatively little is known about proteases that are responsible for the degradation of non-native proteins in the periplasmic compartment of gram-negative bacteria. Such function has been attributed to the heat-shock protein DegP, also commonly referred to as HtrA or Protease Do. While most factors involved in protein quality control are ATP-dependent heat-shock proteins [38], DegP fulﬁlls this role without consuming chemical energy [79]. DegP homologs are found in bacteria, fungi, plants, and mammals. Some, but not all, are classical heat-shock proteins. They are localized in extracytoplasmic compartments and have a modular architecture composed of an N-terminal segment believed to have regulatory functions, a conserved trypsin-like protease domain and one or two PDZ domains at the C-terminus [20].

28 Molecular Machines for Protein Degradation

PDZ domains are protein modules that mediate speciﬁc protein–protein interactions and bind preferentially to the C-terminal 3–4 residues of the target protein [104]. Prokaryotic DegPs have been attributed to the tolerance against thermal, osmotic, oxidative, and pH stress as well as to pathogenicity [91]. A number of DegP substrates are known. These are either largely unstructured proteins such as casein, small proteins that tend to denature, hybrid proteins, or proteins that entered a non-productive folding pathway [70, 79, 113]. Stably folded proteins are normally not degraded. In addition to its protease activity, DegP has a general chaperone function. The dual functions switch in a temperature-dependent manner, the protease activity being most apparent at elevated temperatures [113]. The ability to switch between refolding and degradation activity and the large variety of known substrates make DegP a key factor controlling protein stability and turnover. 4.6 The DegP Protomer, a PDZ Protease

DegP from E. coli was crystallized at low temperatures in its chaperone conformation and analyzed [71]. The protomer can be divided into three functionally distinct domains, namely a protease and two PDZ domains, PDZ1 and PDZ2 (see Figure 14). Like other members of the trypsin family, the protease domain of DegP has two perpendicular β-barrel lobes with a C-terminal helix. The catalytic triad is located in the crevice between the two lobes. While the core of the protease domain is highly conserved, there are striking differences in the surface loops L1, L2, and L3 (for nomenclature see Perona and Craik [92], which are important for the adjustment of the catalytic triad (Asp105, His135, Ser210) and the speciﬁcity pocket S1. The enlarged loop LA protrudes into the active site of a molecular neighbor, where it intimately interacts with loops L1 and L2. The resulting loop triad LA*-L1-L2 completely blocks the substrate-binding cleft and results in a severe deformation of the proteolytic site abolishing formation of the catalytic triad, the oxyanion hole, and the S1 speciﬁcity pocket. Thus the protease domain of the DegP chaperone is present in an inactive state, in which substrate binding as well as catalysis is prevented [71]. The structure of the PDZ domains of DegP is similar to PDZ domains of bacterial origin [77]. Compared to the canonical 4+2 PDZ β-sandwich [16], the DegP PDZ domains show a circularly permuted secondary structure, in which the Nand C-terminal strands are exchanged. Furthermore, they contain a 20-residue insertion following the ﬁrst β-strand (including helix f) that is important for interand intramolecular contacts within the oligomer. In analogy to other PDZ domains, PDZ1 and/or PDZ2 should be involved in substrate binding. PDZ1 contains a deep binding cleft for substrate, which is mainly constructed by strand 14, its N-terminal loop (the so-called carboxylate-binding loop) and helix h. The carboxylate-binding loop is located in a highly positively charged region and is formed by an E-L-G-I motif, which is similar to the frequently observed G-L-G-F motif [16]. Binding speciﬁcity is mainly conferred by the speciﬁc conﬁguration of the 0, −2, and −3 binding pockets [109], where pocket 0 anchors the side chain of the

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 29

Fig. 14 Structure of DegP protomer. Ribbon presentation of the monomer, in which the individual domains are colored differently. Residues of the catalytic triad are shown in a ball-and-stick model. The nomenclature of secondary structure elements, the termini of the protein and regions that were not deﬁned by electron density are indicated.

C-terminal residue. In PDZ1, all pockets are built by mainly hydrophobic residues. The thermal motion factors point to the ﬂexibility of strand 14 and its associated carboxylate-binding loop, indicating the plasticity of the binding site. Thus PDZ1 seems to be well adapted to bind various stretches of hydrophobic peptide ligands. Different from PDZ1, the occluded binding site of PDZ2 is unlikely to be involved in substrate recognition. 4.7 The Two Forms of the DegP Hexamer

In the crystallographic asymmetric unit, two DegP molecules (A and B) were observed, which build up two distinct hexamers (see Figure 15). Both hexamers are formed by staggered association of two trimeric rings. Hexamer A is a largely open

30 Molecular Machines for Protein Degradation

Fig. 15 Structure of the DegP hexamer. Ribbon presentation of the monomer, in which the individual domains are colored differently. Residues of the catalytic triad are shown in a ball-and-stick model. The nomenclature of secondary structure elements, the termini of the protein and regions that were not deﬁned by electron density are indicated.

structure with a wide lateral passage penetrating the entire complex (see Figure 15A), whereas hexamer B corresponds to the closed form, in which a cylindrical 45-Å cavity containing the proteolytic sites is completely shielded from solvent (see Figure 15B). In both cases, the top and bottom of the DegP cage are constructed by the six protease domains, whereas the twelve PDZ domains generate the mobile sidewalls. The height of the cavity is determined by three molecular pillars, which

4 The Tricorn Protease and its Structural and Functional Relationship with Dipeptidyl Peptidase IV 31

are formed by the enlarged LA loops of the protease domain. The PDZ domains are able to adopt different conformations and represent side doors that may open. This en-bloc mobility enables the PDZ domains to function as tentacular arms capturing substrates and delivering them to the inner cavity. This structural organization is strikingly different from the other cage-forming proteases, where substrates enter the central cavity through narrow axial or lateral pores as described in Sections 2 to 4. 4.8 DegP, a Chaperone

E. coli DegP has the ability to stabilize and support the refolding of several non-native proteins in vivo and in vitro [84, 113]. Possible binding sites for misfolded proteins are located within the inner cavity (see Figure 16). The solvent-accessible height of this chamber is 15 Å at its center and increases to 18 Å near the outer entrance. Owing to these geometric constrictions, substrates must be partially unfolded to reach the active site (see Figure 15). As in other chaperones of known structure, the DegP cavity is lined by hydrophobic residues. Two major hydrophobic grooves can be distinguished, which are mainly constructed by residues of loop LA and L2. Notably, the hydrophobic binding sites of the PDZ1 domains are properly oriented to augment the number of potential binding patches. The alternating arrangement of polar and hydrophobic surfaces, both within one trimeric ring and between trimeric rings, should allow the binding of exposed hydrophobic side chains as well as of the peptide backbone of substrates. Taken together, the ceilings of the DegP cavity represent docking platforms for partially misfolded proteins. Both platforms are structurally ﬂexible and should thus allow binding of diverse polypeptides.

Fig. 16 Properties of the inner cavity. Half cut presentations of molecule A (left: side view, center and right: top views) with cut regions shown in dark gray. (Left) Surface representation of the internal tunnel illustrating its molecular-sieve character. Access is restricted to single secondary structure elements as shown by the modeled polyalanine helix, which is colored yellow. (Center)

Top view on the ceiling of the inner cavity with mapped thermal motion factors to show its plasticity. Flexible regions are colored red, rigid regions are blue. (Right) Formation of the hydrophobic binding patches within the cavity. Hydrophobic residues of the protease domain are shown in cyan, and the non-polar peptide-binding groove of PDZ1 in green.

32 Molecular Machines for Protein Degradation

4.9 The Protease Form

The protease conformation of DegP is still elusive as crystallization of a substratelike inhibitor complex has failed and maintenance of a stably folded protein precludes long-term experimentation at elevated temperatures where it displays protease activity. We propose a profound rearrangement of the LA* –L1–L2 loop triad into the canonical conformation of active serine proteases competent for substrate binding. This may be initiated by a collapse of the hydrophobic LA platforms and an enlargement of the hydrophobic contacts caused at high temperature. 4.10 Working Model for an ATP-independent Heat-shock Protein

Cage-forming proteases and chaperones can be ATP-dependent or -independent. In the former group, ATPase activity is important for recognition of target proteins, their dissociation and unfolding, their translocation within the complex, and for various gating mechanisms. The present crystal structure indicates why these functions are not relevant for DegP. DegP preferably degrades substrates, which are per se partially unfolded and which might accumulate under extreme conditions [79, 114, 116]. Alternatively, threading of substrate through the inner chamber could promote unfolding into an extended conformation. Removal of higher-order structural elements may reinitiate substrate folding after exit from DegP. By binding to the C-terminus or a β-hairpin loop of a protein, the PDZ domains could properly position the substrate for threading it into the central cavity. After accessing this chamber, the fate of the unfolded protein depends on the interplay and structural organization of loops LA, L1, and L2. Recruitment of PDZ domains for the gating mechanism should permit a direct coupling of substrate binding and translocation within the DegP particle. This two-step binding process is similar to that of other cageforming proteins such as the proteasome or the Clp proteases. Here, two binding sites (chambers) exist, the ﬁrst of which primarily determines substrate speciﬁcity.

Acknowledgment

M.B. thanks Dr. Ravishankar Ramachandran, Molecular and Structural Biology division, Central Drug Research Institute, Lucknow, India for a critical reading of his contribution to the manuscript.

References 1 NEUWALD, A. F., ARAVIND, L., SPOUGE, J. L., and KOONIN, E. V. AAA+: A class of chaperone-like ATPases associated with

the assembly, operation, and disassembly of protein complexes. Genome Res 1999, 9, 27–43.

References 2 GLICKMAN, M. H., RUBIN, D. M., FU, H., LARSEN, C. N., COUX, O., WEFES, I., PFEIFER, G., CJEKA, Z., VIERSTRA, R., BAUMEISTER, W., et al. Functional analysis of the proteasome regulatory particle. Mol. Biol. Rep. 1999, 26, 21–28. 3 KRZYWDA, S., BRZOZOWSKI, A. M., VERMA, C., KARATA, K., OGURA, T., and WILKINSON, A. J. The crystal structure of the AAA domain of the ATP-dependent protease FtsH of Escherichia coli at 1.5 A resolution. Structure (Camb) 2002, 10, 1073–1083. 4 NIWA, H., TSUCHIYA, D., MAKYIO, H., YOSHIDA, M., and MORIKAWA, K. Hexameric ring structure of the ATPase domain of the membrane-integrated metalloprotease FtsH from Thermus thermophilus HB8. Structure (Camb) 2002, 10, 1415–1423. 5 WANG, J., HARTLING, J. A., and FLANAGAN, J. M. The structure of ClpP at 2.3 A resolution suggests a model for ATP-dependent proteolysis. Cell 1997, 91, 447–456. 6 KIM, D. Y. and KIM, K. K. Crystal structure of ClpX molecular chaperone from Helicobacter pylori. J. Biol. Chem. 2003, 278, 50664–50670. 7 GUO, F., MAURIZI, M. R., ESSER, L., and XIA, D. Crystal structure of ClpA, an Hsp100 chaperone and regulator of ClpAP protease. J. Biol. Chem. 2002b, 277, 46743–46752. 8 CHUANG, S. E., and BLATTNER, F. R. Characterization of twenty-six new heat shock genes of Escherichia coli. J Bacteriol 1993, 175, 5242–5252. 9 CHUANG, S. E., BURLAND, V., PLUNKETT, G., DANIELS, D. L., and BLATTNER, F. R. Sequence analysis of four new heat shock genes constituting the hslu and hslv operons in Escherichia coli. Gene 1993, 134, 1–6. 10 MISSIAKIS, D., SCHWAGER, F., BETTON, J.-M., GEORGOPOULOS, C., and RAINA, S. Identiﬁcation and characterizaton of HslV HslU (ClpQ ClpY) proteins involved in overall proteolysis of misfolded proteins in Escherichia coli. EMBO J. 1996, 15, 6899–6909. 11 KHATTAR, M. M. Overexpression of the hslVU operon suppresses SOS-mediated

12

13

14

15

16

17

18

19

20

inhibition of cell division in Escherichia coli. FEBS Lett. 1997, 414, 402–404. KANEMORI, M., NISHIHARA, K., YANAGI, H., and YURA, T. Synergistic roles of HslVU and other ATP-dependent proteases in controlling in vivo turnover of σ 32 and abnormal proteins in Escherichia coli. J. Bact. 1997, 179, 7219–7225. KANEMORI, M., YANAGI, H., and YURA, T. The ATP-dependent HslVU/ClpQY protease participates in turnover of cell division inhibitor SulA in Escherichia coli. J Bacteriol 1999, 181, 3674–3680. SEONG, I. S., OH, J. Y., YOO, S. J., SEOL, J. H., and CHUNG, C. H. ATP-dependent degradation of SulA, a cell division inhibitor, by the HslVU protease in Escherichia coli. FEBS Lett. 1999, 456, 211–214. RAWLINGS, N. D., O’BRIEN, E., and BARRETT, A. J. MEROPS: the protease database. Nucleic Acids Res 2002, 30, 343–346. COUVREUR, B., WATTIEZ, R., BOLLEN, A., FALMAGNE, P., LE RAY, D., and DUJARDIN, J. C. Eubacterial HslV and HslU subunits homologs in primordial eukaryotes. Mol Biol Evol 2002, 19, 2110–2117. ROHRWILD, M., COUX, O., HUANG, H. C., MOERSCHELL, R. P., SOON, J. Y., SEOL, J. H., CHUNG, C. H., and GOLDBERG, A. L. HslV-HslU: A novel ATP-dependent protease complex in Escherichia coli related to the eukaryotic proteasome. Proc. Natl. Acad. Sci. USA 1996, 93, 5808–5813. SEOL, J. H., YOO, S. J., SHIN, D. H., SHIM, Y. K., KANG, M.-S., GOLDBERG, A. L., and CHUNG, C. H. The heat shock protein HslVU from Escherichia coli is a protein activated ATPase as well as an ATP-dependent proteinase. Eur. J. Biochem. 1997, 247, 1143–1150. HUANG, H.-C., and GOLDBERG, A. L. Proteolytic activity of the ATP-dependent protease HslVU can be uncoupled from ATP-hydrolysis. J. Biol. Chem. 1997, 272, 21364–21372. LUPAS, A., ZWICKL, P., and BAUMEISTER, W. Proteasome sequences in eubacteria. Trends Biochem. Sci. 1994, 19, 533–534.

33

34 Molecular Machines for Protein Degradation

21 ROHRWILD, M., PFEIFER, G., SANTARIUS, U., M¨ULLER, S. A., HUANG, H. C., ENGEL, A., BAUMEISTER, W., and GOLDBERG, A. L. The ATP-dependent HslVU protease from Escherichia coli is a four-ring structure resembling the proteasome. Nat. Struct. Biol 1997, 4, 133–139. 22 BOCHTLER, M., DITZEL, L., GROLL, M., and HUBER, R. Crystal structure of heat shock locus V (HslV) from Escherichia coli. Proc. Natl. Acad. Sci. USA 1997, 94, 6070–6074. 23 SOUSA, M. C. and MCKAY, D. B. Structure of Haemophilus inﬂuenzae HslV protein at 1.9 A resolution, revealing a cation-binding site near the catalytic site. Acta Crystallogr D Biol Crystallogr 2001, 57, 1950–1954. 24 SONG, H., BOCHTLER, M., AZIM, M., HARTMANN, C., HUBER, R., and RAMACHANDRAN, R. Isolation and characterization of the prokaryotic proteasome homolog HslVU (ClpQY) from Thermotoga maritima and the crystal structure of HslV. Biophys Chem. 2003, 100, 437–452. 25 KANG, M. S., LIM, B. K., SEONG, I. S., SEOL, J. H., TANAHASHI, N., TANAKA, K., and CHUNG, C. H. The ATP-dependent CodWX (HslVU) protease in Bacillus subtilis is an N-terminal serine protease. EMBO J. 2001, 20, 734–742. 26 LEVCHENKO, I., SMITH, C. K., WALSH, N. P., SAUER, R. T., and BAKER, T. A. PDZ-like domains mediate binding speciﬁcity in the Clp/Hsp100 family of chaperones and protease regulatory subunits. Cell 1997, 91, 939–947. 27 BOCHTLER, M., HARTMANN, C., SONG, H. K., BOURENKOV, G. P., BARTUNIK, H. D., and HUBER, R. The structures of HsIU and the ATP-dependent protease HsIU-HsIV. Nature 2000, 403, 800–805. 28 WANG, J., SONG, J. J., SEONG, I. S., FRANKLIN, M. C., KAMTEKAR, S., EOM, S. H., and CHUNG, C. H. Nucleotide-dependent conformational changes in a protease-associated ATPase HsIU. Structure (Camb) 2001b, 9, 1107–1116. 29 KARATA, K., INAGAWA, T., WILKINSON, A. J., TATSUTA, T., and OGURA, T. Dissecting the role of a conserved motif (the second

30

31

32

33

34

35

36

37

region of homology) in the AAA family of ATPases. Site-directed mutagenesis of the ATP-dependent protease FtsH. J. Biol. Chem. 1999, 274, 26225– 26232. SONG, H. K., HARTMANN, C., RAMACHANDRAN, R., BOCHTLER, M., BEHRENDT, R., MORODER, L., and HUBER, R. Mutational studies on HslU and its docking mode with HslV. Proc. Natl. Acad. Sci. USA 2000, 97, 14103–14108. SMITH, C. K., BAKER, T. A., and SAUER, R. T. Lon and Clp family proteases and chaperones share homologous substrate-recognition domains. Proc. Natl. Acad. Sci. USA 1999, 96, 6678–6682. WICKNER, S. and MAURIZI, M. R. Here’s the hook: similar substrate binding sites in the chaperone domains of Clp and Lon. Proc. Natl. Acad. Sci. USA 1999, 96, 8318–8320. LEE, Y. Y., CHANG, C. F., KUO, C. L., CHEN, M. C., YU, C. H., LIN, P. I., and WU, W. F. Subunit oligomerization and substrate recognition of the Escherichia coli ClpYQ (HslUV) protease implicated by in vivo protein-protein interactions in the yeast two-hybrid system. J. Bacteriol. 2003, 185, 2393–2401. ORTEGA, J., SINGH, S. K., ISHIKAWA, T., MAURIZI, M. R., and STEVEN, A. C. Visualization of substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 2000, 6, 1515–1521. KANG, M. S., KIM, S. R., KWACK, P., LIM, B. K., AHN, S. W., RHO, Y. M., SEONG, I. S., PARK, S. C., EOM, S. H., CHEONG, G. W., et al. Molecular architecture of the ATP-dependent CodWX protease having an N-terminal serine active site. EMBO J. 2003, 22, 2893–2902. KESSEL, M., MAURIZI, M. R., KIM, B., KOCSIS, E., TRUS, B. L., SINGH, S. K., and STEVEN, A. C. Homology in structural organization between E. coli ClpAP protease and the eukaryotic 26S proteasome. J. Mol. Biol 1995, 250, 587–594. GRIMAUD, R., KESSEL, M., BEURON, F., STEVEN, A. C., and MAURIZI, M. R. Enzymatic and structural similarities

References

38

39

40

41

42

43

44

45

between the Escherichia coli ATP-dependent proteases, ClpXP and ClpAP. J. Biol. Chem. 1998, 273, 12476–12481. KESSEL, M., WHU, W. F., GOTTESMAN, S., KOCSIS, E., STEVEN, A. C., and MAURIZI, M. R. Six-fold rotational symmetry of ClpQ, the E. coli homolog of the 20S proteasome, and its ATP-dependent activator, ClpY. FEBS Lett. 1996, 398, 274–278. SOUSA, M. C., TRAME, C. B., TSURUTA, H., WILBANKS, S. M., REDDY, V. S., and MCKAY, D. B. Crystal and solution structures of an HslUV protease-chaperone complex. Cell 2000, 103, 633–643. WANG, J., SONG, J. J., FRANKLIN, M. C., KAMTEKAR, S., IM, Y. J., RHO, S. H., SEONG, I. S., LEE, C. S., CHUNG, C. H., and EOM, S. H. Crystal structures of the HslVU peptidase-ATPase complex reveal an ATP- dependent proteolysis mechanism. Structure (Camb) 2001a, 9, 177–184. KWON, A. R., KESSLER, B. M., OVERKLEEFT, H. S., and MCKAY, D. B. Structure and reactivity of an asymmetric complex between HslV and I-domain deleted HslU, a prokaryotic homolog of the eukaryotic proteasome. J. Mol. Biol. 2003, 330, 185–195. BOCHTLER, M., SONG, H. K., HARTMANN, C., RAMACHANDRAN, R., and HUBER, R. The quaternary arrangement of HslU and HslV in a cocrystal: a response to Wang, Yale. J Struct Biol 2001, 135, 281–293. WANG, J. A second response in correcting the HslV-HslU quaternary structure. J Struct Biol 2003, 141, 7–8. ISHIKAWA, T., MAURIZI, M. R., BELNAP, D., and STEVEN, A. C. Docking of components in a bacterial complex. Nature 2000, 408, 667–668. BOGYO, M., MCMASTER, J. S., GACZINSKA, M., TORTORELLA, D., GOLDBERG, A. L., and PLOEGH, H. Covalent modiﬁcation of the active site threonine of proteasomal β-subunits and the Escherichia coli homolog HslV by a new class of inhibitors. Proc. Nat. Acad. Sci. USA 1997, 94, 6629–6634.

46 SOUSA, M. C., KESSLER, B. M., OVERKLEEFT, H. S., and MCKAY, D. B. Crystal structure of HslUV complexed with a vinyl sulfone inhibitor: corroboration of a proposed mechanism of allosteric activation of HslV by HslU. J. Mol. Biol. 2002, 318, 779–785. 47 RAMACHANDRAN, R., HARTMANN, C., SONG, H. K., HUBER, R., and BOCHTLER, M. Functional interactions of HslV (ClpQ) with the ATPase HslU (ClpY). Proc. Natl. Acad. Sci. USA 2002, 99, 7396–7401. 48 SEONG, I. S., KANG, M. S., CHOI, M. K., LEE, J. W., KOH, O. J., WANG, J., EOM, S. H., and CHUNG, C. H. The C-terminal tails of HslU ATPase act as a molecular switch for activation of HslV peptidase. J. Biol. Chem. 2002, 277, 25976–25982. 49 BABBITT, P. C. and GERLT, J. A. Understanding enzyme superfamilies. Chemistry as the fundamental determinant in the evolution of new catalytic activities. J. Biol. Chem. 1997, 272, 30591–30594. 50 BRANNIGAN, J. A., DODSON, G., DUGGLEBY, H. J., MOODY, P. C. E., SMITH, J. L., TOMCHICK, D. R., and MURZIN, A. G. A protein catalytic framework with an N-terminal nucleophile is capable of self-activation. Nature 1995, 378, 416–419. 51 BANECKI, B., WAWRZYNOW, A., PUZEWICZ, J., GEORGOPOULOS, C., and ZYLICZ, M. Structure-function analysis of the zinc-binding region of the Clpx molecular chaperone. J. Biol. Chem. 2001, 276, 18843–18848. 52 DONALDSON, L. W., WOJTYRA, U., and HOURY, W. A. Solution structure of the dimeric zinc binding domain of the chaperone ClpX. J. Biol. Chem. 2003, 278, 48991–48996. 53 BOCHTLER, M., Ph. D. Thesis, TU M¨unchen 1999. 54 GUO, F., ESSER, L., SINGH, S. K., MAURIZI, M. R., and XIA, D. Crystal structure of the heterodimeric complex of the adapter, ClpS, with the N-domain of the AAA+ chaperone, ClpA. J. Biol. Chem. 2002a, 277, 46753–46762. 55 DOUGAN, D. A., MOGK, A., ZETH, K., TURGAY, K., and BUKAU, B. AAA+

35

36 Molecular Machines for Protein Degradation

56

57

58

59

60

61

62

63

64

65

proteins and substrate recognition, it all depends on their partner in crime. FEBS Lett. 2002a, 529, 6–10. DOUGAN, D. A., WEBER-BAN, E., and BUKAU, B. Targeted delivery of an ssrA-tagged substrate by the adapter protein SspB to its cognate AAA+ protein ClpX. Mol Cell 2003, 12, 373–380. LEVCHENKO, I., GRANT, R. A., WAH, D. A., SAUER, R. T., and BAKER, T. A. Structure of a delivery protein for an AAA+ protease in complex with a peptide degradation tag. Mol Cell 2003, 12, 365–372. SONG, H. K. and ECK, M. J. Structural basis of degradation signal recognition by SspB, a speciﬁcity-enhancing factor for the ClpXP proteolytic machine. Mol Cell 2003, 12, 75–86. ZHOU, Y., GOTTESMAN, S., HOSKINS, J. R., MAURIZI, M. R., and WICKNER, S. The RssB response regulator directly targets sigma(S) for degradation by ClpXP. Genes Dev 2001, 15, 627–637. STUDEMANN, A., NOIRCLERC-SAVOYE, M., KLAUCK, E., BECKER, G., SCHNEIDER, D., and HENGGE, R. Sequential recognition of two distinct sites in sigma(S) by the proteolytic targeting factor RssB and ClpX. EMBO J. 2003, 22, 4111–4120. DOUGAN, D. A., REID, B. G., HORWICH, A. L., and BUKAU, B. ClpS, a substrate modulator of the ClpAP machine. Mol Cell 2002b, 9, 673–683. ZETH, K., RAVELLI, R. B., PAAL, K., CUSACK, S., BUKAU, B., and DOUGAN, D. A. Structural analysis of the adapter protein ClpS in complex with the N-terminal domain of ClpA. Nat. Struct. Biol. 2002, 9, 906–911. KIM, Y. I., LEVCHENKO, I., FRACZKOWSKA, K., WOODRUFF, R. V., SAUER, R. T., and BAKER, T. A. Molecular determinants of complex formation between Clp/Hsp100 ATPases and the ClpP peptidase. Nat. Struct. Biol. 2001, 8, 230–233. WILK, S. and CHEN, W.-E. Synthetic peptide based activators of the proteasome. Mol. Biol. Rep. 1997, 24, 119–124. WHITBY, F. G., MASTERS, E. I., KRAMER, L., KNOWLTON, J. R., YAO, Y., WANG, C. C., and HILL, C. P. Structural basis for

66

67

68

69

70

71

72

73

74

the activation of 20S proteasomes by 11S regulators. Nature 2000, 408, 115–120. WALZ, J., ERDMANN, A., KANIA, M., TYPKE, D., KOSTER, A. J., and BAUMEISTER, W. 26S proteasome structure revealed by three-dimensional electron microscopy. J Struct Biol 1998, 121, 19–29. ZWICKL, P., NG, D., WOO, K. M., KLENK, H. P., and GOLDBERG, A. L. An archaebacterial ATPase, homologous to ATPases in the eukaryotic 26 S proteasome, activates protein breakdown by 20 S proteasomes. J. Biol. Chem. 1999, 274, 26008–26014. RICHMOND, C., GORBEA, C., and RECHSTEINER, M. Speciﬁc interactions between the ATPase subunits of the 26S protease. J. Biol. Chem. 1997, 272, 13403–13411. GROLL, M., DITZEL, L., L¨OWE, J., STOCK, D., BOCHTLER, M., BARTUNIK, H. D., and HUBER, R. Structure of 20S proteasome from yeast at 2.4 Å resolution. Nature 1997, 386, 463–471. L¨OWE, J., STOCK, D., JAP, B., ZWICKL, P., BAUMEISTER, W., and HUBER, R. Crystal structure of the 20S proteasome from the archaeon T. acidophilum at 3.4 Å resolution. Science 1995, 268, 533– 539. UNNO, M., MIZUSHIMA, T., MORIMOTO, Y., TOMISUGI, Y., TANAKA, K., YASUOKA, N., and TSUKIHARA, T. The Structure of the Mammalian 20S Proteasome at 2.75 A Resolution. Structure 2002, 10 (5), 609–618. ARENDT, C. S. and HOCHSTRASSER, M. Eukaryotic 20S proteasome catalytic subunit propeptides prevent active site inactivation by N-terminal acetylation and promote particle assembly. EMBO J. 1999, 18, 3575–3585. GROLL, M., HEINEMEYER, W., JAGER, S., ULLRICH, T., BOCHTLER, M., WOLF, D. H., and HUBER, R. The catalytic sites of 20S proteasomes and their role in subunit maturation: a mutational and crystallographic study. Proc. Natl. Acad. Sci. USA 1999, 96, 10976–10983. GROLL, M., BRANDSTETTER, H., BARTUNIK, H., BOURENKOW, G., and HUBER, R. Investigations on the maturation and regulation of archaebacterial

References

75

76

77

78

79

80

81

82

proteasomes. J Mol Biol. 2003, 327, 75–83. DITZEL, L., HUBER, R., MANN, K., HEINEMEYER, W., WOLF, D. H., and GROLL, M. Conformational constraints for protein self-cleavage in the proteasome. J. Mol. Biol. 1998, 279, 1187–1191. DICK, T. P., NUSSBAUM, A. K., DEEG, M., HEINEMEYER, W., GROLL, M., SCHIRLE, M., KEILHOLZ, W., STEVANOVIC, S., WOLF, D. H., HUBER, R., et al. Contribution of proteasomal beta-subunits to the cleavage of peptide substrates analyzed with yeast mutants. J. Biol. Chem. 1998, 273, 25637–25646. NUSSBAUM, A. K., DICK, T. P., KEILHOLZ, W., SCHIRLE, M., STEVANOVIC, S., DIETZ, K., HEINEMEYER, W., GROLL, M., WOLF, D. H., HUBER, R., et al. Cleavage motifs of the yeast 20S proteasome beta subunits deduced from digests of enolase 1. Proc. Natl. Acad. Sci. USA 1998, 95, 12504–12509. CARDOZO, C., VINITSKY, A., MICHAUD, C., and ORLOWSKI, M. Evidence that the nature of amino acid residues in the P3 position directs substrates to distinct catalytic sites of the pituitary multicatalytic proteinase complex (proteasome). Biochemistry 1994, 33, 6483–6489. BOGYO, M., SHIN, S., MCMASTER, J. S., and PLOEGH, H. L. Substrate binding and sequence preference of the proteasome revealed by active-site-directed afﬁnity probes. Chem Biol 1998, 5, 307–320. GROLL, M., NAZIF, T., HUBER, R., and BOGYO, M. Probing structural determinants distal to the site of hydrolysis that control substrate speciﬁcity of the 20S proteasome. Chem. Biol. 2002, 9, 1–20. HEINEMEYER, W., FISCHER, M., KRIMMER, T., STACHON, U., and WOLF, D. H. The active sites of the eukaryotic 20S proteasome and their involvement in subunit precursor processing. J. Biol. Chem. 1997, 272, 25200–25209. J¨AGER, S., GROLL, M., HUBER, R., WOLF, D. H., and HEINEMEYER, W. Proteasome beta-type subunits: unequal roles of propeptides in core particle maturation

83

84

85

86

87

88

89

90

91

and a hierarchy of active site function. J. Mol. Biol. 1999, 291, 997–1013. WENZEL, T., ECKERSKORN, C., LOTTSPEICH, F., and BAUMEISTER, W. Existence of a molecular ruler in proteasomes suggested by analysis of degradation products. FEBS Lett. 1994, 349, 205–209. ADAMS, J., BEHNKE, M., CHEN, S., CRUICKSHANK, A. A., DICK, L. R., GRENIER, L., KLUNDER, J. M., MA, Y.-T., PLAMONDON, L., and STEIN, R. L. Potent and selective inhibitors of the proteasome: dipeptidyl boronic acids. Bioorg. Med. Chem. Lett. 1998, 8, 333–338. ELOFSSON, M., SPLITTGERBER, U., MYUNG, J., MOHAN, R., and CREWS, C. M. Towards subunit-speciﬁc proteasome inhibitors: synthesis and evaluation of peptide alpha , beta -epoxyketones. Chem Biol 1999, 6, 811–822. FENTEANY, G., STANDAERT, R. F., LANE, W. S., CHOI, S., COREY, E. J., and SCHREIBER, S. L. Inhibition of proteasome activities and subunit-speciﬁc amino-terminal threonine modiﬁcation by lactacystin. Science 1995, 268, 726–731. MENG, L., MOHAN, R., KWOK, B. H., ELOFSSON, M., SIN, N., and CREWS, C. M. Epoxomicin, a potent and selective proteasome inhibitor, exhibits in vivo antiinﬂammatory activity. Proc. Natl. Acad. Sci. USA 1999, 96, 10403–10408. GROLL, M., KIM, K. B., KAIRIES, N., HUBER, R., and CREWS, C. M. Crystal structure of epoxomicin: 20S proteasome reveals a molecular basis for selectivity of alpha , beta -epoxyketone proteasome inhibitors. J. Am. Chem. Soc. 2000b, 122, 1237–1238. KLOETZEL, P. Antigen processing by the proteasome. Nat Rev Mol Cell Biol. 2001, 2, 179–187. KOGUCHI, Y., KOHNO, J., NISHIO, M., TAKAHASHI, K., OKUDA, T., OHNUKI, T., and KOMATSUBARA, S. TMC-95A, B, C, and D, novel proteasome inhibitors produced by Apiospora montagnei Sacc. TC 1093. Taxonomy, production, isolation, and biological activities. J Antibiot (Tokyo) 2000, 53, 105–109. KOHNO, J., KOGUCHI, Y., NISHIO, M., NAKAO, K., KURODA, M., SHIMIZU, R.,

37

38 Molecular Machines for Protein Degradation

92

93

94

95

96

97

98

99

OHNUKI, T., and KOMATSUBARA, S. Structures of TMC-95A-D: novel proteasome inhibitors from Apiospora montagnei sacc. TC 1093. J Org Chem 2000, 65, 990–995. GROLL, M., KOGUCHI, Y., HUBER, R., and KOHNO, J. Crystal structure of the 20 S proteasome: TMC-95A complex: a non-covalent proteasome inhibitor. J. Mol. Biol. 2001, 311, 543– 548. KAISER, M., GROLL, M., RENNER, C., HUBER, R., and MORODER, L. The core structure of TMC-95A is a promising lead for reversible proteasome inhibition. Angew. Chem. Int. Ed. 2002, 41, 780–783. LIN, S. and DANISHEFSKY, S. The total synthesis of proteasome inhibitors TMC-95A and TMC-95B: discovery of a new method to generate cis-propenyl amides. Angew Chem Int Ed Engl. 2002, 41, 512–515. LOIDL, G., GROLL, M., MUSIOL, H. J., HUBER, R., and MORODER, L. Bivalency as a principle for proteasome inhibition. Proc. Natl. Acad. Sci. USA 1999, 96, 5418–5422. NAVON, A., and GOLDBERG, A. L. Proteins are unfolded on the surface of the ATPase ring before transport into the proteasome. Mol Cell 2001, 8 (6), 1339–1349. BENAROUDJ, N., ZWICKL, P., SEEMULLER, E., BAUMEISTER, W., and GOLDBERG, A. L. ATP hydrolysis by the proteasome regulatory complex PAN serves multiple functions in protein degradation. Mol Cell 2003, 11, 69–78. GROLL, M., BAJOREK, M., K¨OHLER, A., MORODER, L., RUBIN, D. M., HUBER, R., GLICKMAN, M. H., and FINLEY, D. A gated channel into the proteasome core particle. Nat. Struct. Biol. 2000a, 7, 1062–1067. GLICKMAN, M. H., RUBIN, D. M., COUX, O., WEFES, I., PFEIFER, G., CJEKA, Z., BAUMEISTER, W., FRIED, V. A., and FINLEY, D. A subcomplex of the proteasome regulatory particle required for ubiquitin-conjugate degradation and related to the COP9-signalosome and eIF3. Cell 1998, 94, 615–623.

100 K¨OHLER, A., CASCIO, P., LEGGETT, D. S., WOO, K. M., GOLDBERG, A. L., and FINLEY, D. The axial channel of the proteasome core particle is gated by the Rpt2 ATPase and controls both substrate entry and product release. Molecular Cell 2001, 7, 1143–1152. 101 TAMURA, T., TAMURA, N., CEJKA, Z., HEGERL, R., LOTTSPEICH, F., and BAUMEISTER, W. Tricorn Protease – The Core of a Modular Proteolytic System. Science 1996a, 274, 1385–1389. 102 BRANDSTETTER, H., KIM, J.-S., GROLL, M., and HUBER, R. The crystal structure of the tricorn protease reveals a protein disassembly line. Nature 2001, 414, 466–469. 103 F¨ULO¨ P, V., B¨OCSKEI, Z., and POLGA´ R, L. Prolyl Oligopeptidase: An Unusual β-Propeller Domain Regulates Proteolysis. Cell 1998, 94, 161–170. 104 ENGEL, M., HOFFMANN, T., WAGNER, L., WERMANN, M., HEISER, U., KIEFERSAUER, R., HUBER, R., BODE, W., DEMUTH, H.-U., and BRANDSTETTER, H. The crystal structure of dipeptidyl peptidase IV (CD26) reveals its functional regulation and enzymatic mechanism. Proc. Natl. Acad. Sci. USA 2003, 100, 5063–5068. 105 RASMUSSEN, H. B., BRANNER, S., WIBERG, F. C., and WAGTMANN, N. Crystal structure of human dipeptidyl peptidase IV/CD26 in complex with a substrate analog. Nat. Struct. Biol. 2003, 9. 106 BODE, W., MAYR, I., BAUMANN, U., HUBER, R., STONE, S. R., and HOFSTEENGE, J. The reﬁned 1.9 A crystal structure of human alpha-thrombin: interaction with D-Phe-Pro-Arg chloromethylketone and signiﬁcance of the Tyr-Pro-Pro-Trp insertion segment. EMBO J. 1989, 8, 3467–3475. 107 EICHINGER, A., BEISEL, H. G., JACOB, U., HUBER, R., MEDRANO, F. J., BANBULA, A., POTEMPA, J., TRAVIS, J., and BODE, W. Crystal structure of gingipain R: an Arg-speciﬁc bacterial cysteine proteinase with a caspase-like fold. EMBO J. 1999, 18, 5453–5462. 108 TAMURA, T., TAMURA, N., LOTTSPEICH, F., and BAUMEISTER, W. Tricorn protease (TRI) interacting factor 1 from Thermoplasma acidophilum is a proline

References

109

110

111

112

113

114

115

116

117

iminopeptidase. FEBS Lett. 1996b, 398, 101–105. BEEBE, K. D., SHIN, J., PENG, J., CHAUDHURY, C., KHERA, J., and PEI, D. Substrate Recognition through a PDZ Domain in Tail-Speciﬁc Protease. Biochemistry 2000, 39, 3149–3155. PONTING, C. P. and PALLEN, M. J. β-Propeller repeats and a PDZ domain in the tricorn protease: predicted self-compartmentalisation and C-terminal polypeptide-binding strategies of substrate selection. FEMS Microbiol. Lett. 1999, 179, 447–451. LIAO, D. I., QIAN, J., CHISHOLM, D. A., JORDAN, D. B., and DINER, B. A. Crystal structures of the photosystem II D1 C-terminal processing protease. Nat. Struct. Biol. 2000, 7, 749–753. CABRAL, J. H. M., PETOSA, C., SUTCLIFFE, M. J., RAZA, S., BYRON, O., POY, F., MARFATIA, S. M., CHISHTI, A. H., and LIDDINGTON, R. C. Crystal structure of a PDZ domain. Nature 1996, 382, 649–652. DOYLE, D. A., LEE, A., LEWIS, J., KIM, E., SHENG, M., and MACKINNON, R. Crystal structures of a complexed and peptide-free membrane protein-binding domain: Molecular basis of peptide recognition by PDZ. Cell 1996, 85, 1067–1076. KIM, J.-S., GROLL, M., MUSIOL, H. J., BEHRENDT, R., KAISER, M., MORODER, L., HUBER, R., and BRANDSTETTER, H. Navigation inside a protease: substrate selection and product exit in the tricorn protease from Thermoplasma acidophilum. J. Mol. Biol. 2002, 324, 1041–1050. F¨ULO¨ P, V., SZELTNER, Z., and POLGAR, L. Catalysis of serine oligopeptidases is controlled by a gating ﬁlter mechanism. EMBO Reports 2000, 1, 277–281. ITO, N., PHILLIPS, S. E. V., STEVENS, C., OGEL, Z. B., MCPHERSON, M. J., KEEN, J. N., YADAV, K. D. S., and KNOWLES, P. F. Novel thioether bond revealed by a 1.7 A crystal structure of galactose oxidase. Nature 1991, 35, 87–90. GOETTIG, P., GROLL, M., KIM, J.-S., HUBER, R., and BRANDSTETTER, H. Crystal structures of the tricorn interacting aminopeptidase F1 with various ligands

118

119

120

121

122

123

124

125

126

127

reveal its catalytic mechanism. EMBO J. 2002, 21. TAMURA, N., LOTTSPEICH, F., BAUMEISTER, W., and TAMURA, T. The role of tricorn protease and its aminopeptidaseinteracting factors in cellular protein degradation. Cell 1998, 95, 637–648. POSPISILIK, J. A., HINKE, S. A., PEDERSON, R. A., HOFFMANN, T., ROSCHE, F., SCHLENZIG, D., GLUND, K., HEISER, U., MCINTOSH, C. H., and DEMUTH, H. Metabolism of glucagon by dipeptidyl peptidase IV (CD26). Regul Pept 2001, 96, 133–141. WICKNER, S., MAURIZI, R., and GOTTESMAN, S. Posttranslational Quality Control: Folding, Refolding, and Degrading Proteins. Science 1999, 286, 1888–1893. GOTTESMAN, S., WICKNER, S., and MAURIZI, M. R. Protein quality control: Triage by chaperones and proteases. Genes & Development 1997, 11, 815– 823. LIPINSKA, B., ZYLICZ, M., and GEORGOPOULOS, C. The Htra (Degp) Protein, Essential For Escherichia-Coli Survival At High-Temperatures, Is an Endopeptidase. J. Bacteriol. 1990, 172, 1791–1797. CLAUSEN, T., SOUTHAN, C., and EHRMANN, M. The HtrA family of proteases: implications for protein composition and cell fate. Mol Cell 2002, 10, 443–455. SHENG, M. and SALA, C. PDZ domains and the organization of supramolecular complexes. Annu. Rev. Neurosci. 2001, 24, 1–29. PALLEN, M. J. and WREN, B. W. The HtrA family of serine proteases. Molec. Microbiol. 1997, 26, 209–221. KOLMAR, H., WALLER, P. R. H., and SAUER, R. T. The DegP and DegQ periplasmic endoproteases of Escherichia coli: Speciﬁcity for cleavage sites and substrate conformation. J. Bacteriol. 1996, 178, 5925–5929. SPIESS, C., BEIL, A., and EHRMANN, M. A temperature-dependent switch from chaperone to protease in a widely conserved heat shock protein. Cell 1999, 97, 339–347.

39

40 Molecular Machines for Protein Degradation

128 KROJER, T., GARRIDO-FRANCO, M., HUBER, R., EHRMANN, M., and CLAUSEN, T. Crystal structure of DegP (HtrA) reveals a new protease-chaperone machine. Nature 2002, 416, 455–459. 129 PERONA, J. J. and CRAIK, C. S. Structural basis of substrate speciﬁcity in the serine proteases. Protein Sci 1995, 4, 337– 360. 130 SONGYANG, Z., FANNING, A. S., FU, C., XU, J., MARFATIA, S. M., CHISHTI, A. H., CROMPTON, A., CHAN, A. C., ANDERSON, J. M., and CANTLEY, L. C. Recognition of unique carboxyl-terminal motifs by distinct PDZ domains. Science 1997, 275, 73–77. 131 MISRA, R., CASTILLOKELLER, M., and DENG, M. Overexpression of protease-deﬁcient DegP(S210A) rescues the lethal phenotype of Escherichia coli

OmpF assembly mutants in a degP background. J. Bacteriol. 2000, 182, 4882–4888. 132 SWAMY, K. H. S., CHIN, H. C., and GOLDBERG, A. L. Isolation and Characterization of Protease Do From Escherichia-Coli, a Large Serine Protease Containing Multiple Subunits. Arch. Biochem. Biophys. 1983, 224, 543–554. 133 STRAUCH, K. L., JOHNSON, K., and BECKWITH, J. Characterization of Degp, a Gene Required For Proteolysis in the Cell-Envelope and Essential For Growth of Escherichia-Coli At High-Temperature. J. Bacteriol. 1989, 171, 2689–2696. 134 WANG, J. A corrected quaternary arrangement of the peptidase HslV and atpase HslU in a cocrystal structure. J Struct Biol 2001, 134, 15–24.

1

Protein Sorting to the Lysosome

John McCullough, Michale J. Clague, and Sylvie Urb´e University of Liverpool, Liverpool, United Kingdom

Originally published in: Protein Degradation, Volume 3. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31435-0

1 Introduction

The eukaryotic cell contains two major proteolytic systems that mediate protein turnover. The 26S proteasome governs the degradation of most intracellular proteins, whereas the lysosome is responsible for downregulation of cell surface receptors and for the degradation of exogenous proteins that have been internalized. The discovery of ubiquitin in the 1970s led to the identiﬁcation of the 26S proteasome, which supplanted the notion of the lysosome as the principal player in intracellular protein degradation [1]. There is therefore a nice irony that, as the molecular mechanisms of lysosomal sorting have been unravelled, covalent attachment of ubiquitin has been shown to play a key role in routing proteins through the endomembrane system for lysosomal degradation. Appendage of K48-linked polyubiquitin chains targets proteins for proteasomal destruction. However, polyubiquitin chains linked through any of six other internal lysine residues are also represented within eukaryotic cells and their functions are largely unknown [2]. Emerging roles for ubiquitin have been accompanied by an appreciation of ubiquitination as a dynamic modiﬁcation, which can be used to govern the assembly and disassembly of macromolecular complexes, much like phosphorylation, through interaction with speciﬁc ubiquitin-binding domains. Accordingly, additional roles can be proposed for ubiquitin modiﬁcation at endosomal membranes, such as cell signalling. A receptor normally enters the endosomal system through incorporation into Clathrin-coated vesicles (CCVs) and delivery to a tubulo-vesicular compartment known as the early or sorting endosome (Figure 1) [3]. From here receptors may recycle to the plasma membrane, or be selected for lysosomal sorting, by incorporation into small vesicles that bud away from the cytosol into the vacuolar lumen Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Protein Sorting to the Lysosome

Fig. 1 Protein sorting in the endocytic pathway. Upon ligand activation (not shown), growth factor receptors (e.g., EGF receptor) are internalized via Clathrin-coated vesicles (CCV) and delivered to the tubulo-vesicular early endosome. Here, ubiquitinated receptors are sequestered into the lumenal

vesicles of the multi-vesicular body (MVB), which delivers its content to the lysosome. Other plasma membrane proteins that are not ubiquitinated (e.g., Transferrin receptor) enter the pathway via the same route but are recycled back to the plasma membrane.

to generate multi-vesicular bodies (MVBs). MVBs may then fuse directly with lysosomes or deliver material to late endosomes, both of which contain acid-dependent proteases. Classic electron microsocopy studies established this paradigm, largely by following the itinerary of epidermal growth factor receptors (EGFR), subsequent to ligand-induced internalization [4–6]. This endocytic pathway has been conserved in yeast, with the yeast vacuole being functionally equivalent to the lysosome. Thus, much of our recent knowledge has been driven by yeast genetic studies of mutants defective in vacuolar transport (vps mutants) [7], which has led to the identiﬁcation of the MVB sorting machinery. In this article we shall focus ﬁrst on the work that led to an appreciation of a role for ubiquitin on the endocytic pathway, before discussing the complex machinery that engages with ubiquitin for sorting towards the lysosome. We shall then consider the various E3-ligases and de-ubiquitinating enzymes (DUBs), which have

2 Identiﬁcation of Ubiquitin as an Endosomal Sorting Signal

been mooted to regulate the ubiquitin status of endosomal proteins, before ﬁnally considering the role of different polyubiquitin chains at the endosome.

2 Identiﬁcation of Ubiquitin as an Endosomal Sorting Signal

The ﬁrst hints that ubiquitin might play a role in endocytosis emerged in the mid-1980s, when it was found that a number of plasma membrane receptors, including PDGF receptor and GH receptor, were modiﬁed with ubiquitin [8, 10]; reviewed in [151]. In the early 1990s, studies revealed that ubiquitination of cell surface receptors that signal through tyrosine kinase activation occurs in response to binding of ligands. The ﬁrst example of such was the T-cell receptor (TCR). Multiple TCR subunits were found to be ubiquitinated on cytosolic lysine residues in response to receptor occupancy [9]. It required yeast studies to clarify a function for conjugation of ubiquitin to membrane proteins. A link between ubiquitin and endocytosis was established using three different approaches. Firstly, Kolling and Hollenberg analysed the downregulation of Ste6p, the ATP-binding cassette (ABC) transporter for secretion of the pheromone α-factor [11]. They observed the accumulation of ubiquitinated forms of Ste6p in plasma membrane fractions prepared from a mutant Saccharomyces cerevisiae strain with impaired endocytosis. Second, by analysing yeast strains lacking ubiquitin-conjugating enzymes, Hicke and Riezman showed that ubiquitination of the yeast G protein-coupled α-factor receptor Ste2p marked this plasma membrane protein for proteolysis [12]. Using mutant strains, defective for either vacuolar hydrolases or proteasome function, they were able to establish that ubiquitination of Ste2p was required for the vacuolar degradation pathway. A third line of evidence for a link between ubiquitin and endocytosis in yeast was obtained by genetic analysis of the ammonium-induced downregulation of amino acid permeases. Npi1 was isolated as a nitrogen permease inactivator gene in 1983 [13]; Hein et al., subsequently showed that the gene encodes an E3-ligase, Rsp5p [14]. In contrast to proteasomal degradation pathways, appendage of polyubiquitin chains to substrate proteins is not required for ubiquitin-dependent endocytosis in yeast. Ste2p is efﬁciently internalized under conditions where polyubiquitin chain formation is suppressed by the use of ubiquitin mutants lacking internal lysines [15]. Fusion of a single ubiquitin in-frame to the stable plasma membrane protein Pma1p stimulates endocytosis of this protein [16]. For other proteins, speciﬁcally yeast permeases, such as uracil permease and the general amino acid permease, Gap1p, maximal internalization rates require the formation of diubiquitin chains linked through Lys63 of ubiquitin [17, 18]. In mammalian cells, the smeary appearance of ubiquitinated receptors on Western blots was initially interpreted as reﬂecting polyubiquitination. However, it was not clear how polyubiquitinated receptors would escape proteasomal targeting, and instead be degraded in the lysosome. This issue has since been resolved by using a combination of monoclonal antibodies, which allow discrimination between

3

4 Protein Sorting to the Lysosome

mono- and polyubiquitinated proteins. EGFR, PDGFR and Met tyrosine kinase receptors are all in fact monoubiquitinated at multiple lysine residues within the receptors [19–21]. Furthermore, Haglund et al., showed that fusion of ubiquitin to the C-terminus of EGFR results in constitutive endocytosis, which cannot be further enhanced by EGF stimulation. However, data from the Sorkin laboratory has shown that the dopamine transporter can be polyubiquitinated and contains a mixture of K11, K48 and K63-linked ubiquitin [22]. In addition to promoting the internalization of receptors from the plasma membrane, ubiquitination also promotes sorting of receptors towards late endosomes and the vacuole. Mutation of lysine 6 in the cytoplasmic tail of the yeast vacuolar protein Phm5p inhibits its sorting to lumenal vesicles, but this can be restored by the biosynthetic addition of a single ubiquitin to create a Ub-Phm5p fusion protein [23]. In mammalian cells, transferrin receptor normally recycles back to the plasma membrane after internalization to sorting endosomes [24]. Fusion of ubiquitin to the C-terminus of this receptor prevents its recycling, through interaction with the endosomal protein Hrs (Section 3.2; [25]).

3 Ubiquitin-mediated Sorting at the Endosome: The MVB Sorting Machinery

Our appreciation of the complexity of the molecular machinery responsible for MVB formation and receptor sorting to the lysosome, owes everything to the comprehensive characterization of vps mutant strains in yeast [26]. These vps mutants can be subdivided into classes based on their characteristic phenotype, and it is Class E mutants that present with an enlarged, swollen prevacuolar compartment, indicative of a defect at the stage of MVB formation and inward vesiculation. The sorting events at the prevacuolar or early endosome that lead to the translocation of receptors into internal vesicles are thought to be mediated by a succession of at least four multiprotein complexes, which each have the ability to recognize ubiquitinated cargo through ubiquitin-interacting domains: the Hrs/STAM complex and the ESCRT complexes I, II and III. A ﬁfth component that is essential for this process is an ATPase of the AAA family called Vps4 or SKD1. 3.1 Endosome-associated Ubiquitin Interacting Domains: Structure and Function

Several of the class E vps genes contain domains that are predicted to interact with ubiquitin. A feature common to all ubiquitin-binding motifs is their low afﬁnity (100–500 µM, see Table 1). This makes sense in the face of the high concentration of free ubiquitin in the cytosol which has been estimated at 10 µM [31]; too high an afﬁnity would mean that these modules would be permanently plugged with ubiquitin and unable to dynamically interact with ubiquitinated cargo. The small size of these domains has made them amenable to structural analysis.

3 Ubiquitin-mediated Sorting at the Endosome: The MVB Sorting Machinery Table 1 Ubiquitin-binding domains in endocytic proteins

Name

Length

Examples

K d (l M)

Structure

UIM

∼20 ∼40 ∼45–50 ∼60 ∼150 var.

NZF

∼25–30

Vps36

Hrs:∼300 Vps27:∼100–300 STAM:∼200 n.d. Vps9: 20 µM Tsg101:∼500 n.d. yGGA: 100–400 Vps36:∼200

α-helix

UBA CUE UEV VHS GAT

Hrs/Vps27, STAM/Hsel, Eps15, Epsin Cbl Tollip Tsg101/Vps23 STAM GGA3

Triple α-helix Triple α-helix α-helix/β-sheets Eight α-helices Triple α-helix Zn ﬁnger

A large variety of ubiquitin-binding motifs are found in proteins involved in endocytic membrane trafﬁc. The average length of the motif is given in amino acid residues. Kd values correspond to measured afﬁnities for free ubiquitin. n.d.: not determined. Refer to text for references and abbreviations.

The ubiquitin-interacting motif (UIM) was ﬁrst identiﬁed in an unbiased bioinformatic screen based on homology to the ubiquitin-interacting region in the regulatory subunit S5a (Rpn10 in yeast) of the 26S proteasome [27]. It became immediately clear that proteins involved in endocytic trafﬁcking were highly represented amongst the emerging list of UIM proteins. UIM domains are found in Hrs, STAM and Epsin as well as in their yeast counterparts Vps27, HseI and Ent1/2. The UIM is characterized by a short 20 amino acid motif with a highly conserved stretch xxAxxxSxxAc, where denotes a large hydrophobic, and Ac an acidic, residue, and which is preceded by a block of four mostly acidic residues [27]. Solution structures of the UIMs of Hrs and Vps27, as well as a crystal structure of the second UIM of Vps27, are now available and indicate that the motif folds as a short amphipathic helix and interacts with the Leu8-Ile44-Val70 hydrophobic patch in ubiquitin [28–30]. The structures indicate that UIM binding to a monoubiquitinated protein would occlude Lys48 of ubiquitin, rendering it unavailable for ubiquitin chain extension through this residue. This may provide one mechanism that protects ubiquitinated receptors from proteasomal degradation. Ubiquitin-associated (UBA) domains [32, 33] and CUE-domains (named after the founding member Cue1: coupling of ubiquitin conjugation to ER degradation) span 40 to 50 amino acids and share a three-helix bundle structure [34–36]. Two highresolution crystal structures of the CUE domain show that binding to ubiquitin, as in the case of the UIM, is through hydrophobic surfaces, and that Lys48 of ubiquitin is likewise occluded by the interaction [34, 36]. Ubiquitin E2 variant (UEV) domains have homology to the E2 conjugating enzymes that ligate ubiquitin to substrates, but they lack the catalytic cysteine. In contrast to UIM, UBA and CUE domains, the crystal structure of the ESCRT I component Tsg101 UEV domain complexed with ubiquitin indicates that both Lys 48 and Lys 63 of ubiquitin remain fully accessible [37, 38].

5

6 Protein Sorting to the Lysosome

Many UIM, UBA and CUE domains promote self-ubiquitination of their host protein. This can promote a network of proteins held together by ubiquitin interactions, or may regulate the availability of the ubiquitin-binding domain in the host protein [39, 40]. 3.2 The Hrs–STAM Complex and the Endosomal Clathrin Coat

Initial engagement between the sorting machinery and ubiquitinated cargo is mediated through interaction with UIM domains in hepatocyte growth factor tyrosine regulated substrate (Hrs) and signal transducing adapter molecule (STAM), also called Hrs-binding protein (Hbp). In yeast these proteins correspond to Vps27 and Hse1, respectively. Mutations in the UIM of either of these proteins result in speciﬁc defects in the sorting of ubiquitinated proteins into the vacuole lumen [41, 42]. Association of Hrs with endosomal membranes is mediated through binding of its FYVE domain to the inositol lipid PtdIns3P [43, 44]. Hrs fulﬁls an adapter function through direct interaction with ubiquitinated cargo and with the terminal domain (TD) of clathrin heavy chain [45, 46]. Both Hrs and claxsthrin are components of an unusual coat structure that assembles on the vacuolar surface of sorting endosomes [47]. Typically, the coat presents as an extended ﬂat surface, which gives the impression of opposing the natural curvature of the membrane. It comprises two relatively electron-dense layers separated by a thin electron-lucent layer. This particular clathrin coat does not form clathrin-coated vesicles, but instead may provide a matrix capable of trapping and concentrating ubiquitinated receptors such as EGFR [45]. On the other hand, the recycling transferrin receptor, whilst free to diffuse into this region, is not retained within it [47]. A central role of Hrs/Vps27 in both receptor sorting and MVB formation has been proposed. Vps27 is a class E vps mutant defective in lumenal vesicle formation and Hrs knock-out in Drosophila provides a similar phenotype and inhibits EGFR downregulation [48]. siRNA knock-down of Hrs in mammalian cells partially inhibits Met receptor and EGFR downregulation [49, 50]. A role for Hrs in both receptor sorting and lumenal vesicle formation suggests that the two processes may be tightly coupled, with consequent advantages with respect to the loading efﬁciency. Both receptor sorting and lumenal vesicle formation can also be inhibited by overexpression of Hrs [51]. This inhibition of internal vesicle formation is contingent on an intact Hrs UIM domain, suggesting that it may play both positive (receptor sorting) and negative (vesicle formation) roles in the pathway leading to lumenal vesicle budding. Yet another UIM-domain-containing protein, Eps15, has been reported as an additional subunit of the Hrs–STAM complex [49]. Eps15 has previously been shown to play a role in the CCV-mediated internalization of activated EGFR [52, 53]. The fact that Eps15 clearly functions early on in endocytosis has made it difﬁcult to address whether it is an essential component at a later stage of endosomal sorting. All three proteins, Hrs, STAM and Eps15, are tyrosine phosphorylated in response to growth factors, which could conceivably regulate further associations

3 Ubiquitin-mediated Sorting at the Endosome: The MVB Sorting Machinery

with components of the sorting machinery [44, 54, 55] or with other signalling pathways [56]. 3.3 GGA and Tom1: Alternative Sorting Adapters?

Hrs and STAM are founder members of the family of proteins with a VHS (Vps27p/Hrs/STAM) domain. Striking similarities have now been found with other members, which share the ability to link ubiquitin and Clathrin: GGA (Golgiassociated g-adaptin homologues, Arf-binding) and Tom1/Tom1L1 (target of Myb1) (Figure 2) [57–60]. GGA proteins have previously been described as monomeric adapters that are involved in clathrin-coated vesicular transport between the Golgi and the endosome as well as the endosome and the vacuole/lysosome [150]. Ubiquitin binding is conferred by the GGA and Tom (GAT) domain and by the VHS domain, whereas clathrin is recruited by the Hinge-region [57, 62–64]. The GAT domain is also responsible for binding the small GTPase Arf [65]. In the case of Tom1 and Tom1L1, the GAT domain binds ubiquitin and a CUE-domain protein called Tollip in a mutually exclusive way [60, 66]. Tom1, but not Tom1L1 also binds to a FYVE-domain protein, Endoﬁn, strengthening the analogy with the HrsSTAM complex [67]. Endoﬁn or Tollip co-expression are required for recruitment of Tom1 to endosomes [66, 67]. Finally, all three VHS-domain sorting complexes, Hrs-STAM, GGA and Tom1L1, can independently recruit the internal vesicle formation machinery through a P(S/T)AP motif which interacts with the ESCRTI component tumour suppressor gene 101 (Tsg101, see below) [58, 61, 68]. A yeast two-hybrid assay has also suggested that the VHS domains of GGA1 and 3, as well as of Tom1 are capable of binding to Hrs [58]. It is unclear if the existence of multiple sorting complexes is an indication of redundancy in lysosomal sorting or whether the multitude of adapters provides a higher level of cargo selectivity to the sorting process. An analogy may be made with the clathrin-coated vesicle internalization pathway where a variety of adapters operate to provide both redundant and speciﬁc sorting mechanisms [69]. 3.4 The ESCRT Machinery

Yeast studies have deﬁned three distinct multimeric Vps protein complexes, named endosomal sorting complex required for transport (ESCRT) I, II and III, which are proposed to be sequentially recruited and activated at the endosome (Figure 3) [26]. These proteins constitute the core of the MVB formation machinery downstream of the Hrs–STAM complex. At least two of these complexes bind ubiquitin, and the ubiquitinated receptors may be passed along from one complex to another. The ﬁrst complex, ESCRTI (∼350 kDa) is composed of Vps23, Vps28 and Vps37. The ubiquitin-binding site of this complex is found in Vps23, which has a UEV domain. The mammalian homologue of Vps23, Tsg101 (tumour suppressor gene 101) was originally identiﬁed in a tissue culture screen for genes whose disruption

7

8 Protein Sorting to the Lysosome

Fig. 2 Ubiquitin-binding adapter complexes implicated in MVB sorting. Three endosomal adapter complexes have been implicated in the sorting of proteins from the endosome to the lysosome. Each of these complexes interacts with clathrin, which decorates the endosomal membrane in discrete patches. Ubiquitin binding is conferred by UIM (Hrs, STAM), VHS (STAM, GGA), or GAT domains (GGA, Tom1) and might

provide a means to concentrate ubiquitinated receptors in the endosomal clathrincoated microdomains. The PS/TAP motif in the Hrs, GGA and Tom1 subunits may then recruit the ESCTI complex via direct interaction with Tsg101. It is as yet unclear whether these sorting complexes act independently or in concert with each other. Black rectangle: receptor cargo; PI3P: PtdIns3P; Ub: ubiquitin.

3 Ubiquitin-mediated Sorting at the Endosome: The MVB Sorting Machinery

causes cell transformation [70]. Tsg101 as well as hVps28 disruption by RNAi and antibody injection, respectively, clearly interfere with EGFR downregulation in human cells and cause a marked accumulation of ubiquitin on endosomes [71, 72]. The mammalian homologue of Vps37 has only recently been identiﬁed and is the least conserved member of ESCRTI [73, 74]. ESCRTI is recruited to endosomal membranes through binding of the UEV domain of TSG101/Vps23 to a conserved PT/(S)AP motif within Hrs–Vps27 [68, 75, 76]. Hrs and TSG101 are present in both cytosolic and membrane fractions, but only associate at the membrane [75]. How can this ordered ESCRT complex assembly be attained speciﬁcally at the membrane? Hrs is localized in part at the membrane through interaction of its FYVE domain with the inositide lipid PtdIns3P that is concentrated at early endosomes [43, 44], where it can then bind to ubiquitinated receptors. It is possible that PtdIns3P or ubiquitin binding could induce a conformational change in Hrs that unmasks a TSG101 binding site. TSG101 itself has a PTAP motif, which may interact with its own UEV domain – competitive binding by Hrs may then release TSG101 into a relaxed conformation permissive for ESCRT II recruitment [68, 76, 77]. ESCRTII was described as a cytosolic complex (∼155 kDa) composed of the class E vps proteins Vps22/Eap30, Vps25/Eap25 and Vps36/Eap45 that transiently associates with endosomal membranes in an ESCRTI-dependent manner (Figure 3). Two 3.6-Å resolution crystal structures of an ESCRTII complex containing one molecule of Vps22, the carboxy-terminal domain of Vps36 and two molecules of Vps25 suggest that the complex has the shape of a capital letter ‘Y’, of which the sub-complex Vps22 and Vps36 form one branch. A ﬂexible linker extending from the tip of this branch would lead to two consecutive NZF motifs, the second of which is believed to bind ubiquitinated cargo [78–80]. However, this domain was not solved owing to proteolysis during crystallization. The authors suggest that this structure could provide a “long swinging arm” for transfer of cargo over substantial distances. The NZF motif is lacking in Eap45, the mammalian orthologue of Vps36, but this is compensated for by the inclusion of a GLUE domain, which has been found to have ubiquitin-binding properties [81]. The ESCRTII complex is required for the membrane recruitment of the Snf7Vps20 ESCRTIII sub-complex via an interaction between Vps25 and Vps20 [80, 82], and this in turn is a prerequisite for the recruitment of the other two ESCRTIII proteins Vps24 and Vps2 to the membranes [83, 84]. These last four class E Vps proteins belong to a structurally related “family” of small, highly charged, coiled-coil proteins that in mammalian cells are referred to as CHMPs (charged multivesicular proteins) [85]. Membrane association of this complex may also be conferred by myristoylation of Vps20–CHMP6 and the ability of Vps24–CHMP3 to bind to the phosphatidyl-inositol lipid PtdIns(3,5)P2 [86]. The mammalian homologues of Vps2 and Snf7 are called CHMP2(a and b) and CHMP4(a, b and c), respectively. One highly connected protein is AIP1 or ALIX (ALG2-interacting protein) previously implicated in apoptosis. AIP1 shows interaction with both ESCRTI (Tsg101) and ESCRTIII complexes (Snf7–CHMP4(a, b and c)) and may therefore act as a bridge between these two [87]. The yeast homologue of AIP1, Bro1, plays a role

9

10 Protein Sorting to the Lysosome

Fig. 3 Interactions of the MVB sorting machinery. The ﬁrst point of engagement of ubiquitinated proteins with the MVB sorting machinery is the Hrs–STAM complex. Hrs (Vps27 in yeast) associates with the endosomal membrane through interaction of its FYVE-domain with PtdIns3P. Both Hrs and STAM (HseI in yeast) bind ubiquitin via their UIM domains. Hrs–Vps27 recruits the ESCRTI complex (composed of Vps23–Tsg101, Vps28 and Vps37), through interaction of its PS/TAP motif with Tsg101–Vps23, which in turn binds ESCRTII (Vps22, Vps25 and Vps36; Eap30, Eap25 and Eap45 in mammalian cells). Both ESCRTI and ESCRTII complexes are able to bind ubiquitin via the UEV domain in Tsg101, and the GLUE and NZF domains in Eap45 and Vps36, respectively. ESCRTII recruits and activates the last

complex in this cascade, ESCRTIII. An additional connection between ESCRTI and ESCRTIII complexes is mediated through AIP1, which in yeast (Bro1) has also been implicated in recruiting the DUB Doa4 to the prevacuolar compartment. ESCRTIII is composed of two subcomplexes, Snf7Vps20 and Vps2–Vps24. These proteins belong to a family of highly charged proteins called CHMPs in mammalian cells. Membrane association of the ESCRTIII complex may be mediated by binding of Vps24 to PtdIns(3,5)P2 and a myristoyl moiety attached to Vps20. The ESCRTIII complex interacts with the AAA-ATPase Vps4 (SKD1 in mouse), which is thought to dissociate the components of the sorting machinery. Black bar: receptor, e.g., EGFR; PI3P: PtdIns3P; PI3,5P2 : PtdIns(3,5)P2 ; Ub: ubiquitin.

4 Ubiquitin Ligases and Endosomal Sorting

in recruiting the deubiquitinating enzyme Doa4 [88, 89], possibly through stabilization of Snf7 at the endosome, with which the enzyme interacts directly [90]. Doa4 is not essential for MVB formation or sorting [23, 91], but acts to recycle ubiquitin from cargo that has progressed beyond ubiquitin-dependent steps in the MVB sorting pathway [92]. 3.5 Vps4–SKD1

The ESCRT-III component Snf7, recruits a Class E Vps protein belonging to the AAA-ATPase family, called Vps4 in yeast, and SKD1 in mouse cells [26, 93–95]. The last resolved step in the MVB formation cascade is the dissociation of the ESCRTmachinery powered by Vps4–SKD1 hydrolysis of ATP. This is inferred from the dramatic phenotype of ATPase-defective Vps4–SKD1 mutants in yeast and mammalian cells, respectively. Vps4 yeast deletion strains show a typical Class E swollen prevacuolar compartment on which the entire upstream sorting machinery accumulates [71, 89, 92, 96]. Overexpression of a catalytically inactive Vps4 mutant in mammalian cells recapitulates this phenotype by promoting the formation of enlarged endosomes on which an “Hrs-clathrin coat”, as well as the ESCRTI machinery accumulates [72, 95, 97, 98]. At the ultrastructural level, cells expressing mutant Vps4 also show a depletion of internal vesicles, and membrane proteins destined for the lysosome accumulate at the peripheral membrane of this abnormal compartment [98]. This suggests that dissociation of the sorting machinery and the formation of internal vesicles are tightly coupled to allow efﬁcient recycling of ESCRT proteins.

4 Ubiquitin Ligases and Endosomal Sorting

The speciﬁcity of ubiquitination is largely due to cognate interactions with E3 ligases, of which there are probably in excess of 600 in mammals. By recruiting speciﬁc E2 enzymes they may also determine the topology of the ubiquitin extension. E3s are generally split into two major classes: the RING (really interesting new gene) ligases have a catalytic domain based on a double zinc-ﬁnger whilst the HECT (homologous to E6-AP carboxyl terminus) ligases contain a 350 amino acid C-terminal domain within which lies a conserved catalytic cysteine. HECT ligases recognize their substrate via WW domains that interact with various proline-rich sequences. They form a thiolester intermediate with ubiquitin, whereas RING ligases promote the direct transfer of ubiquitin from the E2 to the substrate [99]. RING ligases can be further subdivided into those in which the substrate binding site and the RING domain are encoded within a single polypeptide (e.g. c-Cbl) and those in which they are contained within different proteins of a larger complex (e.g. SCF and APC/Cyclosome). We will brieﬂy review the major E3 ligases associated with endocytic trafﬁcking.

11

12 Protein Sorting to the Lysosome

4.1 Nedd4 Family

HECT domain proteins of the Nedd4 family regulate the trafﬁcking of a variety of biosynthetic and endosomal cargo. They have a common architecture consisting of several WW domains and a C-terminal HECT domain. Most members also have an amino terminal C2 domain, which commonly bind to phosphoinositides [100]. The sole member of the Nedd4 family in S. cerevisiae is Rsp5, which seems to be the only ligase required for ubiquitination of cell surface proteins. Early studies demonstrated its involvement in constitutive ubiquitination of the uracil permease Fur4p [101] and the general amino acid permease Gap1p [102]. It is located and functions at multiple sites on the endocytic pathway [103]. Accumulation at the prevacuolar compartment in vps4 cells suggests a function in the MVB sorting pathway. Mutation of the C2 domain disrupts this localization and inhibits sorting of the biosynthetic pathway-derived cargo carboxypeptidase S, but not the endocytosed receptor Ste2, for which ubiquitin can be appended at the plasma membrane [104, 105]. Some transporters such as Fur4p are downregulated both by endocytosis from the cell surface and by diversion from the biosynthetic pathway at the Golgi to the MVB pathway, without reaching the surface. Both pathways require Rsp5 function at the plasma membrane and prevacuole respectively [106]. Drosophila Nedd4 regulates endocytosis of Notch and suppresses its ligandindependent activation [107]. In mammalian cells several family members, including Nedd4 and AIP4/Itch, have been implicated in endocytic trafﬁcking. Nedd4 directly mediates ubiquitination of the epithelial Na+ transporter ENaC, targeting it for downregulation [108]. AIP4/Itch is disrupted in nonagouti lethal or itchy mice, which are characterized by abnormal immune responses and constant itching [109]. It interacts with and ubiquitinates mammalian Notch and the G-protein-coupled chemokine receptor CXCR4 [110, 111]. Salient substrates also include components of the endocytic machinery. Eps15 and Hrs ubiquitination are mediated by Nedd4 or by AIP4 [110, 112–114]. Ubiquitination of these proteins depends on their UIM and it is thought that the HECT ligase is partially recruited through an interaction between its covalently attached ubiquitin and the UIM domain. Once the protein is ubiquitinated, its UIM may be occupied by its own ubiquitin and no longer able to recruit another HECT ligase. In this way, polyubiquitination and subsequent targeting of the endocytic machinery to the proteasome may be prevented [39]. 4.2 c-Cbl

In mammalian cells, the major E3 ligase implicated in endocytic trafﬁcking of RTKs is the cellular proto-oncogene Cbl [115]. It is recruited via its SH2 domain to phosphorylated RTKs. The viral oncogene v-cbl lacks the RING ﬁnger motif, and displaces endogenous Cbl, to allow growth factor receptors to escape from downregulation. Similarly, loss of Cbl binding ability through mutation is a recurring theme in oncogenic deregulation of RTKs [116]. The ubiquitination of the Met

5 Endosomal DUBs

receptor is a well-characterized example. Cbl is recruited to activated Met via Grb2, then subsequent binding of its TKB domain to the juxtamembrane autophosphorylated Tyr1003 is proposed to elicit a conformational change within Cbl necessary for activation. A Y1003F mutation inhibits ligand-dependent ubiquitination of Met and leads to cell transformation [117]. Overexpression of c-Cbl, but not mutants lacking ligase activity, dramatically stimulate the degradation of EGFR, apparently without affecting the rate of EGFR internalization [118, 119]. This observation was interpreted to reﬂect ubiquitindependent lysosomal sorting [118]. Further evidence that Cbl might be required for a later step than internalization has come from work done in Cbl-/- mouse embryonic ﬁbroblasts. This work indicated that EGFR is internalized to endosomes at a normal rate in the absence of Cbl, but its degradation was inhibited [120]. However, the exact site of Cbl-action is still much debated and various reports have since proposed a positive role for Cbl-dependent ubiquitination on the EGFR internalization step [121, 122]. Cbl also acts as a multivalent adapter for at least 40 proteins and some of its roles correspond to this adapter function rather than to its ligase activity. For example, a Cbl–Cin85–endophilin complex positively regulates both Met receptor and EGFR endocytosis independently of E3 ligase activity [123, 124].

5 Endosomal DUBs

Deubiquitinating enzymes (DUBs) regulate the ubiquitin status of endosomal proteins in opposition to E3 ligases. Deubiquitination is not considered to be an obligate step on the MVB sorting pathway as biosynthetic production of chimeric proteins incorporating ubiquitin results in efﬁcient targeting to MVBs [23]. DUB activity at the endosome may be necessary to maintain the pool of free ubiquitin required for endosomal sorting. On the other hand it can negatively regulate lysosomal protein degradation if it acts on ubiquitinated receptors prior to their commitment to the lysosomal pathway (Figures 4 and 5). DUBs implicated in endosomal sorting include the yeast proteins Ubp1, Ubp2 and Doa4, and the mammalian proteins UBPY (USP8) and AMSH. 5.1 Ubp1 and Ubp2

Overexpression of a soluble form of Ubp1 is able to stabilize the ABC-transporter Ste6 and the α-factor receptor Ste2, which are transported to the vacuole for degradation [125]. The stabilization effect of Ste6 was shown not to be due to deubiquitination of Ste6 itself, suggesting that the target of Ubp1 may be a component of the protein-transport machinery. Ubp2 shows speciﬁcity for K63 over K48-linked polyubiquitin chains and can antagonize Rsp5 E3 ligase activity [126]. Interestingly

13

14 Protein Sorting to the Lysosome

Fig. 4 Sorting of ubiquitinated receptors at the endosome. Ubiquitination of growth factor receptors (e.g., EGF receptor (EGFR)), by an E3 ligase (e.g. Cbl) promotes their interaction with Hrs in a Clathrin-coated microdomain. DUBs (e.g., AMSH and UBPY) may counteract Cbl activity and oppose MVB sorting and favour recycling. Hrs recruits the ESCRTI complex, which activates and assembles ESCRTII and III. This in-

duces the translocation of the ubiquitinated receptor into internal vesicles of the MVB. Ubiquitin itself is recycled by a DUB (e.g., Doa4) just before, or in parallel with, the disassembly of the ESCRT-machinery by the AAA-ATPase Vps4, which irreversibly seals the sorting process. Note that this illustration combines elements from yeast and mammalian cells.

Ubp2 co-puriﬁes with Rsp5 and is physically linked to Rsp5 through an adapter protein Rup1. 5.2 Doa4

The yeast protein Doa4 was originally shown to interact with the 26S proteasome and proposed to promote proteolysis through removal of ubiquitin from proteolytic intermediates on the proteasome [127, 128]. It has also been proposed to maintain free ubiquitin levels by recycling ubiquitin from cargo molecules that have been committed to the lysosomal sorting pathway, prior to their sequestration away from the cytosol [92, 129]. In common with ESCRT complex components, Doa4 accumulates on the endosome following inactivation of the ATPase Vps4 [92]. Deletion of ESCRT III-complex components blocks this localization [88, 92] and

5 Endosomal DUBs

Fig. 5 The endocytic pathway in yeast and mammalian cells. Endocytosed proteins (e.g., Ste2 and EGFR) are sorted at the early endosome by multiple components of the multivesicular body (MVB) sorting machinery (Hrs–STAM, Vps27–HseI, ESCRTI–III). From here, cargo can be either recycled

back to the plasma membrane or sorted to the MVB or prevacuolar compartments. Sorting to the lysosome and vacuole respectively is enhanced by the activity of E3-ligases (Rsp5, Cbl, Nedd4, AIP4), which may be opposed by DUBs (Ubp1, AMSH, UBPY).

a direct interaction with the ESCRTIII component Snf7 has been shown through two-hybrid screens [90]. 5.3 UBPY

UBPY (USP8), a member of the ubiquitin-speciﬁc processing protease (UBP) family, displays the highest similarity to Doa4 amongst mammalian DUBs. It accumulates upon growth stimulation of starved human ﬁbroblasts and downregulates in response to growth arrest induced by cell–cell contact [130]. A link to endosomal protein sorting was ﬁrst suggested when UBPY was identiﬁed in a far-Western screen for Hbp–STAM binding partners [131]. Mutagenic analysis identiﬁed a consensus sequence PX(V/I)(D/N)RXXKP as a binding module for interaction with the Hbp-STAM-SH3 domain-binding motif [131, 132]. This represents a novel SH3 binding motif lacking the canonical PXXP sequence. UBPY can hydrolyse both K48and K63-linked chains as well as monoubiqitinated EGFR. Overexpression of UBPY retards EGFR degradation whilst conﬂicting data have been reported for the effect

15

16 Protein Sorting to the Lysosome

of siRNA-mediated UBPY knock-down, which was found to either enhance [133] or inhibit EGFR downregulation [134]; Row, P.E., McCullough, J., Clague, M.J. and Urb´e, S., unpublished observation). UBPY can exist as a trimolecular complex with Otubain 1, a member of the OTU family of DUBs, and GRAIL, an E3 ligase crucial for the induction of CD4 T cell anergy [135]. UBPY was shown to deubiquitinate GRAIL in vitro and Otubain 1 opposed this action by limiting the DUB activity of UBPY. UBPY-dependent deubiquitination has also been suggested to prevent degradation of the E3 ligase Nrdp1, which plays a role in regulating steady-state levels of ErbB3 and ErbB4 [136] Gnesutta et al. [137] demonstrated an in vitro interaction between mouse UBPY and the N-terminal half of CDC25Mm, a Ras nucleotide exchange factor. This interaction may be functional in cells, since ubiquitination of CDC25Mm in HEK293 cells is diminished following co-expression of mUBPY. 5.4 AMSH

AMSH was originally isolated as a novel adapter molecule from human T cells that interacts with the SH3 domain of STAM (associated molecule with the SH3 domain) [138]. Endogenous STAM was pulled down when AMSH was immunoprecipitated from IL-2 cell lysates in the presence of cross-linkers and both proteins were shown to interact when co-transfected into cells. Intriguingly, AMSH shares the PX(V/I)(D/N)RXXKP STAM/Hbp-SH3 domain-binding motif of UBPY [131]. Thus, AMSH presumably competes with UBPY for binding to the SH3 domain of STAM. AMSH belongs to the JAMM (JAB1/MPN/Mov34) metallo-enzyme family of deubiquitinating enzymes, of which the Rpn11-POH1 subunit of the 19S proteasome lid was the ﬁrst described representative [139, 140]. No DUB activity was observed when Rpn11 was isolated from its multisubunit complex. In contrast, puriﬁed AMSH cleaves ubiquitin chains in vitro, and is hence the ﬁrst JAMM-domain DUB to exhibit activity in isolation [141]. In common with Ubp2, AMSH displays speciﬁcity for K63-over K48-linked polyubiquitin chains [141]. AMSH localizes to endosomes and an inactivating mutation in the JAMM domain of AMSH was shown to promote the accumulation of ubiquitin on endosomes [141]. Concomitantly, this mutant stabilizes an ubiquitinated form of STAM, which is contingent on an intact UIM within STAM. This led us to suggest that ubiquitin, which is appended to STAM in a UIM-dependent fashion and which would normally be removed by either AMSH or UBPY, may provide an additional binding site for enzymatically inactive AMSH. Hence, the inactive mutant of AMSH could act as a “substrate trap” mutant. Ubiquitinated EGFR provides a substrate for AMSH in vitro and siRNA-mediated knock-down of AMSH enhances the degradation rate of EGFR [141]. We have proposed a model for the role of AMSH on endosomes in which AMSH can counteract the E3 ligase activity of c-Cbl on EGFR, before the ubiquitinated receptor has been committed to the lysosomal sorting pathway (Figure 4). AMSH activity

6 Polyubiquitin Linkages and Endocytosis 17

will therefore favour recycling of the receptor. Note that existing data suggest that either AMSH or UBPY can fulﬁl this role (Figure 5). AMSH also associates with a number of signalling molecules. Signalling defects in cells derived from AMSH-deﬁcient mice were not obvious, although the mice die at about three weeks of age [142]. It was originally hypothesized that AMSH may play a role in cytokine-mediated signalling through its interaction with STAM [138]. A novel Grb2 family member, Gads/Grf40, also associates with AMSH [143]. Gads has been shown to be involved in T-cell receptor (TCR) signalling; Gads knock-out or Gads SH2 transgenic mice show impairment in pre-T-cell development. In addition, AMSH also interacts with inhibitory Smads (I-Smads), and this association negatively regulates their function, and thereby promotes bone morphogenetic protein (BMP)-mediated signalling [144]. Li and Seth [145] have demonstrated that AMSH itself is ubiquitinated by the E3 ligase Smurf2. This is contingent on both proteins binding to the adaptor molecule RFN11 and leads to a reduction in steady-state levels of AMSH by proteasomal degradation. We can now see that association of E3 ligase activity with DUB activity is a recurring theme common to UBP2, UBPY and AMSH.

6 Polyubiquitin Linkages and Endocytosis

Although monoubiquitination may represent a minimal requirement for endosomal sorting and RTKs do not seem to incorporate polyubiquitin chains, there is a large body of work suggesting polyUb involvement in some sorting events. 6.1 Proteasome Involvement in Endocytic Sorting

The downregulation of a sub-set of RTKs and other receptors requires proteasomal activity. In most cases studied so far this requirement does not reﬂect proteasomal degradation of the receptor per se. Rather, this activity is permissive for receptor sorting towards the lysosomal degradation pathway. Thus receptor downregulation can be sensitive to both proteasome inhibitors (e.g. lactacystin) and inhibition of lysosomal acidiﬁcation (e.g. concanamycin). K48-linked polyubiquitin chains specify proteasomal degradation and are therefore indirectly implicated in the lysosomal sorting process. Well-characterized examples include interleukin-2 receptors [146], Growth Hormone Receptor (GHR) [147] and the RTK Met [50, 148]. In each case, inhibition of the proteasome promotes recycling of internalized receptors at the expense of sorting to lysosomes. Interestingly, ubiquitination of GHR itself appears to be dispensable for downregulation, which has led to a model in which polyubiquitination and proteasomal degradation of an unidentiﬁed accessory factor is required [147]. An example of such a scenario may be found in neurons, where the endocytosis of AMPA-type glutamate receptors also requires ubiquitination and proteasomal degradation of the scaffolding protein PSD-95 [149].

18 Protein Sorting to the Lysosome

Proteasome inhibition could reduce free ubiquitin levels leading to a block in endocytosis. In the case of Met receptor the inhibitory effect of lactacystin can be overcome by overexpression of ubiquitin, but not by a form of ubiquitin unable to form K48 linkages (K48R) [21]. It is bafﬂing that a K48-linkage dependence is observed in the presence of a proteasomal inhibitor. Does this represent an entirely novel function for K48-linked ubiquitin, which has previously been uniquely associated with a proteasomal targeting signal? One scenario may be that the critical step for lysosomal sorting consists of the sequestration of a polyubiquitinated accessory factor away from Met, which is followed by incidental proteasomal degradation. In this model, proteasome activity is only required to generate free ubiquitin (possibly locally), which can contribute to K48-chain formation.

6.2 K63-linked Ubiquitin

Pioneering studies in yeast have provided evidence for a role of K63-linked polyubiquitin in vacuolar sorting. Gap1p and Fur4p have both been shown to be modiﬁed with short (2–3) K63-linked polyubiquitin chains [17, 18, 106]. In doa4 cells, which show low levels of endocytosis due to limited availabilty of ubiquitin, the endocytosis rate can be restored by overexpression of wild type or K48R ubiquitin but not by K63R ubiquitin, which can participate in monoubiquitination events but cannot form K63-linked chains. K63-linked chains are well represented in the pool of total cellular polyubiquitin [2]. It may well be that in the excitement over monoubiquitination, we have underestimated the role of K63-linked chains in lysosomal sorting. After all, these can provide higher afﬁnity interactions with ubiquitin-binding domains, whilst still avoiding proteasomal degradation.

7 Conclusions

Now that most of the core components of the ubiquitin-dependent MVB sorting pathway have been identiﬁed, the challenge lies in elucidating the choreography underlying this complex process. How is cargo passed between ESCRT complexes or is the whole process more co-operative? The signiﬁcance of the ubiquitin-chain topology is likely to receive more attention and more ubiquitin-binding domains with distinct speciﬁcities remain to be identiﬁed. Allied to this one may consider that ubiquitin may not simply be a tag for sorting at the endosome but may coordinate spatially segregated signalling events through recruitment of adapter proteins and enzymes with ubiquitin-binding domains. The endosome may thus provide a “hot spot” for ubiquitin-dependent signalling.

References

Acknowledgements

We thank the Wellcome Trust and North West Cancer Research Fund for funding our research in this area. SU is a Wellcome Career Development Fellow and JM is the recipient of a Wellcome Trust prize studentship.

References 1 CIECHANOVER, A. (2005). Proteolysis: from the lysosome to ubiquitin and the proteasome. Nat. Rev. Mol. Cell Biol. 6, 79–86. 2 PENG, J., SCHWARTZ, D., ELIAS, J. E., THOREEN, C. C., CHENG, D., MARSISCHKY, G., ROELOFS, J., FINLEY, D., and GYGI, S. P. (2003). A proteomics approach to understanding protein ubiquitination. Nat. Biotechnol. 21, 921–926. 3 CLAGUE, M. J. (1998). Molecular aspects of the endocytic pathway. Biochem. J. 336, 271–282. 4 FELDER, S., MILLER, K., MOEHREN, G., ULLRICH, A., SCHLESSINGER, J., and HOPKINS, C. R. (1990). Kinase activity controls the sorting of the epidermal growth factor receptor within the multivesicular body. Cell 61, 623–634. 5 GORDEN, P., CARPENTIER, J. L., COHEN, S., and ORCI, L. (1978). Epidermal growth factor: morphological demonstration of binding, internalization, and lysosomal association in human ﬁbroblasts. Proc. Natl. Acad. Sci. USA 75, 5025–5029. 6 HAIGLER, H. T., MCKANNA, J. A., and COHEN, S. (1979). Direct visualization of the binding and internalization of a ferritin conjugate of epidermal growth factor in human carcinoma cells A-431. J. Cell Biol. 81, 382–395. 7 BANKAITIS, V. A., JOHNSON, L. M., and EMR, S. D. (1986). Isolation of yeast mutants defective in protein targeting to the vacuole. Proc. Natl. Acad. Sci. USA 83, 9075–9079. 8 LEUNG, D. W., SPENCER, S. A., CACHIANES, G., HAMMONDS, R. G., COLLINS, C., HENZEL, W. J., BARNARD, R., WATERS, M. J., and WOOD, W. I. (1987). Growth hormone receptor and serum binding protein: puriﬁcation, cloning and expression. Nature 330, 537–543.

9 YARDEN, Y., ESCOBEDO, J. A., KUANG, W. J., YANG-FENG, T. L., DANIEL, T. O., TREMBLE, P. M., CHEN, E. Y., ANDO, M. E., HARKINS, R. N., FRANCKE, U. et al. (1986). Structure of the receptor for platelet-derived growth factor helps deﬁne a family of closely related growth factor receptors. Nature 323, 226–232. 10 BONIFACINO, J. S., and WEISSMAN, A. (1998). Ubiquitin and the control of protein fate in the secretory and endocytic pathways. Annu. Rev. Cell. Dev. Biol. 14, 19–57. 11 CENCIARELLI, C., HOU, D., HSU, K. C., RELLAHAN, B. L., WIEST, D. L., SMITH, H. T., FRIED, V. A., and WEISSMAN, A. M. (1992). Activation-induced ubiquitination of the T cell antigen receptor. Science 257, 795–797. 12 KOLLING, R. and HOLLENBERG, C. P. (1994). The ABC-transporter Ste6 accumulates in the plasma membrane in a ubiquitinated form in endocytosis mutants. Embo J. 13, 3261–3271. 13 HICKE, L. and RIEZMAN, H. (1996). Ubiquitination of a yeast plasma membrane receptor signals its ligand-stimulated endocytosis. Cell 84, 277–287. 14 GRENSON, M. (1983). Inactivation-reactivation process and repression of permease formation regulate several ammonia-sensitive permeases in the yeast Saccharomyces cerevisiae. Eur. J. Biochem. 133, 135–139. 15 HEIN, C., SPRINGAEL, J. Y., VOLLAND, C., HAGUENAUER-TSAPIS, R., and ANDRE, B. (1995). NPl1, an essential yeast gene involved in induced degradation of Gap1 and Fur4 permeases, encodes the Rsp5 ubiquitin-protein ligase. Mol. Microbiol. 18, 77–87.

19

20 Protein Sorting to the Lysosome

16 TERREL, J., SHIH, S., DUNN, R., and HICKE, L. (1998). Mol. Cell 1, 193–202. 17 SHIH, S. C., SLOPER-MOULD, K. E., and HICKE, L. (2000). Monoubiquitin carries a novel internalization signal that is appended to activated receptors. Embo J. 19, 187–198. 18 GALAN, J. M. and HAGUENAUER-TSAPIS, R. (1997). Ubiquitin lys63 is involved in ubiquitination of a yeast plasma membrane protein. Embo J. 16, 5847–5854. 19 SPRINGAEL, J. Y., GALAN, J. M., HAGUENAUER-TSAPIS, R., and ANDRE, B. (1999). NH4ρ-induced down-regulation of the Saccharomyces cerevisiae Gap1p permease involves its ubiquitination with lysine-63-linked chains. J. Cell Sci. 112 (Pt 9), 1375–1383. 20 HAGLUND, K., SIGISMUND, S., POLO, S., SZYMKIEWICZ, I., DI FIORE, P. P., and DIKIC, I. (2003). Multiple monoubiquitination of RTKs is sufﬁcient for their endocytosis and degradation. Nat. Cell Biol. 5, 461–466. 21 MOSESSON, Y., SHTIEGMAN, K., KATZ, M., ZWANG, Y., VEREB, G., SZOLLOSI, J., and YARDEN, Y. (2003). Endocytosis of receptor tyrosine kinases is driven by monoubiquitylation, not polyubiquitylation. J. Biol. Chem. 278, 21323–21326. 22 CARTER, S., URBE´ , S., and CLAGUE, M. J. (2004). The met receptor degradation pathway: Requirement for K48-linked polyubiquitin independent of proteasome activity. J. Biol. Chem. 281, 5094–5105. 23 MIRANDA, M., WU, C. C., SORKINA, T., KORSTJENS, D., and SORKIN, A. (2005). Enhanced ubiquitylation and accelerated degradation of the dopamine transporter mediated by protein kinase C. J. Biol. Chem. 280, 35617–35624. 24 REGGIORI, F. and PELHAM, H. R. (2001). Sorting of proteins into multivesicular bodies: ubiquitin-dependent and -independent targeting. EMBO J. 20, 5176–5186. 25 HOPKINS, C. R. (1983). Intracellular routing of transferrin and transferrin receptors in epidermoid carcinoma A431 cells. Cell 35, 321–330.

26 RAIBORG, C., BACHE, K. G., GILLOOLY, D. J., MADSHUS, I. H., STANG, E., and STENMARK, H. (2002). Hrs sorts ubiquitinated proteins into clathrin-coated microdomains of early endosomes. Nature Cell Biol. 4, 394–398. 27 KATZMANN, D. J., ODORIZZI, G., and EMR, S. D. (2002). Receptor downregulation and multivesicular-body sorting. Nat. Rev. Mol. Cell Biol. 3, 893–905. 28 HAAS, A. L. and BRIGHT, P. M. (1987). The dynamics of ubiquitin pools within cultured human lung ﬁbroblasts. J. Biol. Chem. 262, 345–351. 29 HOFMANN, K., and FALQUET, L. (2001). A ubiquitin-interacting motif conserved in components of the proteasomal and lysosomal protein degradation systems. TIBS. 26, 347–350. 30 FISHER, R. D., WANG, B., ALAM, S. L., HIGGINSON, D. S., ROBINSON, H., SUNDQUIST, W. I., and HILL, C. P. (2003). Structure and ubiquitin binding of the ubiquitin-interacting motif. J. Biol. Chem. 278, 28976–28984. 31 SHEKHTMAN, A. and COWBURN, D. (2002). A ubiquitin-interacting motif from Hrs binds to and occludes the ubiquitin surface necessary for polyubiquitination in monoubiquitinated proteins. Biochem. Biophys. Res. Commun. 296, 1222–1227. 32 SWANSON, K. A., KANG, R. S., STAMENOVA, S. D., HICKE, L., and RADHAKRISHNAN, I. (2003). Solution structure of Vps27 UIM-ubiquitin complex important for endosomal sorting and receptor downregulation. Embo J. 22, 4597–4606. 33 HOFMANN, K. and BUCHER, P. (1996). The UBA domain: a sequence motif present in multiple enzyme classes of the ubiquitination pathway. Trends Biochem. Sci. 21, 172–173. 34 VADLAMUDI, R. K., JOUNG, I., STROMINGER, J. L., and SHIN, J. (1996). p62, a phosphotyrosine-independent ligand of the SH2 domain of p56lck, belongs to a new class of ubiquitin-binding proteins. J. Biol. Chem. 271, 20235–20237. 35 KANG, R. S., DANIELS, C. M., FRANCIS, S. A., SHIH, S. C., SALERNO, W. J., HICKE, L., and RADHAKRISHNAN, I. (2003). Solution structure of a CUE-ubiquitin complex

References

36

37

38

39

40

41

42

43

44

reveals a conserved mode of ubiquitin binding. Cell 113, 621–630. MUELLER, T. D. and FEIGON, J. (2002). Solution structures of UBA domains reveal a conserved hydrophobic surface for protein-protein interactions. J. Mol. Biol. 319, 1243–1255. PRAG, G., MISRA, S., JONES, E. A., GHIRLANDO, R., DAVIES, B. A., HORAZDOVSKY, B. F., and HURLEY, J. H. (2003). Mechanism of ubiquitin recognition by the CUE domain of Vps9p. Cell 113, 609–620. PORNILLOS, O., ALAM, S. L., RICH, R. L., MYSZKA, D. G., DAVIS, D. R., and SUNDQUIST, W. I. (2002). Structure and functional interactions of the Tsg101 UEV domain. Embo J. 21, 2397–2406. SUNDQUIST, W. I., SCHUBERT, H. L., KELLY, B. N., HILL, G. C., HOLTON, J. M., and HILL, C. P. (2004). Ubiquitin recognition by the human TSG101 protein. Mol. Cell 13, 783–789. DI FIORE, P. P., POLO, S., and HOFMANN, K. (2003). When ubiquitin meets ubiquitin receptors: a signalling connection. Nat. Rev. Mol. Cell Biol. 4, 491–497. HICKE, L. and DUNN, R. (2003). Regulation of membrane protein transport by ubiquitin and ubiquitin-binding proteins. Annu. Rev. Cell Dev. Biol. 19, 141–172. BILODEAU, P. S., URBANOWSKI, J. L., WINISTORFER, S. C., and PIPER, R. C. (2002). The Vps27p Hse1p complex binds ubiquitin and mediates endosomal protein sorting. Nat. Cell Biol. 4, 534–539. SHIH, S. C., KATZMANN, D. J., SCHNELL, J. D., SUTANTO, M., EMR, S. D., and HICKE, L. (2002). Epsins and Vs27p/Hrs contain ubiquitin-binding domains that function in receptor endocytosis. Nature Cell Biol. 4, 389–393. GILLOOLY, D. J., MORROW, I. C., LINDSAY, M., GOULD, R., BRYANT, N. J., GAULLIER, J. M., PARTON, R. G., and STENMARK, H. (2000). Localization of phosphatidylinositol 3-phosphate in yeast and mammalian cells. EMBO J. 19, 4577–4588.

45 URBE´ , S., MILLS, I. G., STENMARK, H., KITAMURA, N., and CLAGUE, M. J. (2000). Endosomal localization and receptor dynamics determine tyrosine phosphorylation of hepatocyte growth factor-regulated tyrosine kinase substrate. Mol. Cell Biol. 20, 7685– 7692. 46 CLAGUE, M. J. (2002). Membrane transport: a coat for ubiquitin. Curr. Biol. 12, R529–R531. 47 RAIBORG, C., GRONVOLD BACHE, K., MEHLUM, A., STANG, E., and STENMARK, H. (2001). Hrs recruits clathrin to early endosomes. EMBO J. 20, 5008–5021. 48 SACHSE, M., URBE´ , S., OORSCHOT, V., STROUS, G. J., and KLUMPERMAN, J. (2002). Bilayered clathrin coats on endosomal vacuoles are involved in protein sorting toward lysosomes. Mol. Biol. Cell 13, 1313–1328. 49 LLOYD, T. E., ATKINSON, R., WU, M. N., ZHOU, Y., PENNETTA, G., and BELLEN, H. J. (2002). Hrs regulates endosome membrane invagination and tyrosine kinase receptor signaling in Drosophila. Cell 108, 261–269. 50 BACHE, K. G., RAIBORG, C., MEHLUM, A., and STENMARK, H. (2003). STAM and Hrs are subunits of a multivalent ubiquitin-binding complex on early endosomes. J. Biol. Chem. 278, 12513–12521. 51 HAMMOND, D. E., CARTER, S., MCCULLOUGH, J., URBE´ , S., VANDE WOUDE, G., and CLAGUE, M. J. (2003). Endosomal dynamics of met determine signaling output. Mol. Biol. Cell 14, 1346–1354. 52 URBE´ , S., SACHSE, M., ROW, P. E., PREISINGER, C., BARR, F. A., STROUS, G., KLUMPERMAN, J., and CLAGUE, M. J. (2003). The UIM domain of Hrs couples receptor sorting to vesicle formation. J. Cell Sci. 116, 4169–4179. 53 CONFALONIERI, S., SALCINI, A. E., PURI, C., TACCHETTI, C., and Di FIORE, P. P. (2000). Tyrosine phosphorylation of Eps15 is required for ligand-regulated, but not constitutive, endocytosis. J. Cell Biol. 150, 905–912. 54 TORRISI, M. R., LOTTI, L. V., BELLEUDI, F., GRADINI, R., SALCINI, A. E.,

21

22 Protein Sorting to the Lysosome

55

56

57

58

59

60

61

62

63

CONFALONIERI, S., PELICCI, P. G., and Di FIORE, P. P. (1999). Eps15 is recruited to the plasma membrane upon epidermal growth factor receptor activation and localizes to components of the endocytic pathway during receptor internalization. Mol. Biol. Cell 10, 417–434. CLAGUE, M. J. and URBE´ , S. (2001). The interface of receptor trafﬁcking and signalling. J. Cell Sci. 114, 3075– 3081. FAZIOLI, F., MINICHIELLO, L., MATOSKOVA, W. T., WONG, W. T., and Di FIORE, P. P. (1993). Eps15, a novel tyrosine kinase substrate exhibits transforming activity. Mol. Cell Biol. 13, 5814–5828. ROW, P. E., CLAGUE, M. J., and URBE, S. (2005). Growth factors induce differential phosphorylation proﬁles of the Hrs-Stam complex: a common node in signaling networks with signal speciﬁc properties. Biochem. J. 389, 629–636. BILODEAU, P. S., WINISTORFER, S. C., ALLAMAN, M. M., SURENDHRAN, K., KEARNEY, W. R., ROBERTSON, A. D., and PIPER, R. C. (2004). The GAT domains of clathrin-associated GGA proteins have two ubiquitin-binding motifs. J. Biol. Chem. 279, 54808–54816. PUERTOLLANO, R. (2005). Interactions of TOM1L1 with the multivesicular body sorting machinery. J. Biol. Chem. 280, 9258–9264. BONIFACINO, J. S. (2004). The GGA proteins: adaptors on the move. Nat. Rev. Mol. Cell Biol. 5, 23–32. SCOTT, P. M., BILODEAU, P. S., ZHDANKINA, O., WINISTORFER, S. C., HAUGLUND, M. J., ALLAMAN, M. M., KEARNEY, W. R., ROBERTSON, A. D., BOMAN, A. L., and PIPER, R. C. (2004). GGA proteins bind ubiquitin to facilitate sorting at the trans-Golgi network. Nat. Cell Biol. 6, 252–259. YAMAKAMI, M., YOSHIMORI, T., and YOKOSAWA, H. (2003). Tom1, a VHS domain-containing protein, interacts with tollip, ubiquitin, and clathrin. J. Biol. Chem. 278, 52865–52872. PUERTOLLANO, R., RANDAZZO, P. A., PRESLEY, J. F., HARTNELL, L. M., and BONIFACINO, J. S. (2001). The GGAs

64

65

66

67

68

69

70

71

72

promote ARF-dependent recruitment of clathrin to the TGN. Cell 105, 93–102. SHIBA, Y., KATOH, Y., SHIBA, T., YOSHINO, K., TAKATSU, H., KOBAYASHI, H., SHIN, H. W., WAKATSUKI, S., and NAKAYAMA, K. (2004). GAT (GGA and Tom1) domain responsible for ubiquitin binding and ubiquitination. J. Biol. Chem. 279, 7105–7111. ZHU, Y., DORAY, B., POUSSU, A., LEHTO, V. P., and KORNFELD, S. (2001). Binding of GGA2 to the lysosomal enzyme sorting motif of the mannose 6-phosphate receptor. Science 292, 1716–1718. DELL’ANGELICA, E. C., PUERTOLLANO, R., MULLINS, C., AGUILAR, R. C., VARGAS, J. D., HARTNELL, L. M., and BONIFACINO, J. S. (2000). GGAs: a family of ADP ribosylation factor-binding proteins related to adaptors and associated with the Golgi complex. J. Cell Biol. 149, 81–94. KATOH, Y., SHIBA, Y., MITSUHASHI, H., YANAGIDA, Y., TAKATSU, H., and NAKAYAMA, K. (2004). Tollip and Tom1 form a complex and recruit ubiquitin-conjugated proteins onto early endosomes. J. Biol. Chem. 279, 24435–24443. SEET, L. F., LIU, N., HANSON, B. J., and HONG, W. (2004). Endoﬁn recruits TOM1 to endosomes. J. Biol. Chem. 279, 4670–4679. PORNILLOS, O., HIGGINSON, D. S., STRAY, K. M., FISHER, R. D., GARRUS, J. E., PAYNE, M., HE, G. P., WANG, H. E., MORHAM, S. G., and SUNDQUIST, W. I. (2003). HIV Gag mimics the Tsg101-recruiting activity of 4 Ubiquitin and Protein Sorting to the Lysosome the human Hrs protein. J. Cell Biol. 162, 425–434. PUERTOLLANO, R. and BONIFACINO, J. S. (2004). Interactions of GGA3 with the ubiquitin sorting machinery. Nat. Cell Biol. 6, 244–251. TRAUB, L. M. (2003). Sorting it out: AP-2 and alternate clathrin adaptors in endocytic cargo selection. J. Cell Biol. 163, 203–208. LI, L. and COHEN, S. N. (1996). Tsg101: a novel tumor susceptibility gene isolated

References

73

74

75

76

77

78

79

80

81

by controlled homozygous functional knockout of allelic loci in mammalian cells. Cell 85, 319–329. BABST, M., ODORIZZI, G., ESTEPA, E. J., and EMR, S. D. (2000). Mammalian tumour susceptibility gene 101 (TSG101) and the yeast homologue, Vps23p, both function in late endosomal trafﬁcking. Trafﬁc 1, 248–258. BISHOP, N., HORMAN, A., and WOODMAN, P. (2002). Mammalian class E vps proteins recognize ubiquitin and act in the removal of endosomal protein-ubiquitin conjugates. J. Cell Biol. 157, 91–101. BACHE, K. G., SLAGSVOLD, T., CABEZAS, A., ROSENDAL, K. R., RAIBORG, C., and STENMARK, H. (2004). The growth-regulatory protein HCRP1/hVps37A is a subunit of mammalian ESCRT-I and mediates receptor down-regulation. Mol. Biol. Cell 15, 4337–4346. STUCHELL, M. D., GARRUS, J. E., MULLER, B., STRAY, K. M., GHAFFARIAN, S., MCKINNON, R., KRAUSSLICH, H. G., MORHAM, S. G., and SUNDQUIST, W. I. (2004). The human endosomal sorting complex required for transport (ESCRT-I) and its role in HIV-1 budding. J. Biol. Chem. 279, 36059–36071. BACHE, K. G., BRECH, A., MEHLUM, A., and STENMARK, H. (2003). Hrs regulates multivesicular body formation via ESCRT recruitment to endosomes. J. Cell Biol. 162, 435–442. LU, Q., HOPE, L. W., BRASCH, M., REINHARD, C., and COHEN, S. N. (2003). TSG101 interaction with HRS mediates endosomal trafﬁcking and receptor down-regulation. Proc. Natl. Acad. Sci. USA 100, 7626–7631. CLAGUE, M. J. and URBE´ , S. (2003). Hrs function: viruses provide the clue. Trends Cell Biol. 13, 603–606. ALAM, S. L., SUN, J., PAYNE, M., WELCH, B. D., BLAKE, B. K., DAVIS, D. R., MEYER, H. H., EMR, S. D., and SUNDQUIST, W. I. (2004). Ubiquitin interactions of NZF zinc ﬁngers. Embo J. 23, 1411–1421. HIERRO, A., SUN, J., RUSNAK, A. S., KIM, J., PRAG, G., EMR, S. D., and HURLEY, J. H. (2004). Structure of the ESCRT-II

82

83

84

85

86

87

88

89

endosomal trafﬁcking complex. Nature 431, 221–225. TEO, H., PERISIC, O., GONZALEZ, B., and WILLIAMS, R. L. (2004). ESCRT-II, an endosome-associated complex required for protein sorting: crystal structure and interactions with ESCRT-III and membranes. Dev. Cell 7, 559–569. SLAGSVOLD, T., AASLAND, R., HIRANO, S., BACHE, K. G., RAIBORG, C., TRAMBAIANO, D., WAKATSUKI, S., and STENMARK, H. (2005). Eap45 in mammalian ESCRT-II binds ubiquitin via a phosphoinositide-interacting GLUE domain. J. Biol. Chem. 280, 19600–19606. VON SCHWEDLER, U. K., STUCHELL, M., MULLER, B., WARD, D. M., CHUNG, H. Y., MORITA, E., WANG, H. E., DAVIS, T., HE, G. P., CIMBORA, D. M., SCOTT, A., KRANSSLICH, H. G., KAPLAN, J., MORHAM, S. G., and SUNDQUIST, W. J. (2003). The protein network of HIV budding. Cell 114, 701–713. BABST, M., KATZMANN, D. J., ESTEPA-SABAL, E. J., MEERLOO, T., and EMR, S. D. (2002). Escrt-III: an endosome-associated heterooligomeric protein complex required for mvb sorting. Dev. Cell 3, 271–282. BABST, M., KATZMANN, D. J., SNYDER, W. B., WENDLAND, B., and EMR, S. D. (2002). Endosome-associated complex, ESCRT-II, recruits transport machinery for protein sorting at the multivesicular body. Dev. Cell 3, 283–289. HOWARD, T. L., STAUFFER, D. R., DEGNIN, C. R., and HOLLENBERG, S. M. (2001). CHMP1 functions as a member of a newly deﬁned family of vesicle trafﬁcking proteins. J. Cell Sci. 114, 2395–2404. WHITLEY, P., REAVES, B. J., HASHIMOTO, M., RILEY, A. M., POTTER, B. V., and HOLMAN, G. D. (2003). Identiﬁcation of mammalian Vps24p as an effector of phosphatidylinositol 3,5-bisphosphate-dependent endosome compartmentalization. J. Biol. Chem. 278, 38786–38795. KATOH, K., SHIBATA, H., SUZUKI, H., NARA, A., ISHIDOH, K., KOMINAMI, E., YOSHIMORI, T., and MAKI, M. (2003). The

23

24 Protein Sorting to the Lysosome

90

91

92

93

94

95

96

97

ALG-2-interacting protein Alix associates with CHMP4b, a human homologue of yeast Snf7 that is involved in multivesicular body sorting. J. Biol. Chem. 278, 39104–39113. LUHTALA, N. and ODORIZZI, G. (2004). Bro1 coordinates deubiquitination in the multivesicular body pathway by recruiting Doa4 to endosomes. J. Cell Biol. 166, 717–729. ODORIZZI, G., KATZMANN, D. J., BABST, M., AUDHYA, A., and EMR, S. D. (2003). Bro1 is an endosome-associated protein that functions in the MVB pathway in Saccharomyces cerevisiae. J. Cell Sci. 116, 1893–1903. BOWERS, K., LOTTRIDGE, J., HELLIWELL, S. B., GOLDTHWAITE, L. M., LUZIO, J. P., and STEVENS, T. H. (2004). Protein-protein interactions of ESCRT complexes in the yeast Saccharomyces cerevisiae. Trafﬁc 5, 194–210. URBANOWSKI, J. L. and PIPER, R. C. (2001). Ubiquitin sorts proteins into the intralumenal degradative compartment of the late endosome/vacuole. Trafﬁc 2, 622–630. AMERIK, A. Y., NOWAK, J., SWAMINATHAN, S., and HOCHSTRASSER, M. (2000). The DoA4 deubiquitinating enzyme is functionally linked to the vacuolar protein-sorting and endocytic pathways. Mol. Biol. Cell 11, 3365–3380. LIN, Y., KIMPLER, L. A., NAISMITH, T. V., LAUER, J. M., and HANSON, P. I. (2005). Interaction of the mammalian ESCRT-III protein hSnf7 1 with itself, membranes, and the AAAρ ATPase SKD1. J. Biol. Chem. 280, 12799–12809. PERIER, F., COULTER, K. L., LIANG, H., RADEKE, C. M., GABER, R. F., and VANDENBERG, C. A. (1994). Identiﬁcation of a novel mammalian member of the NSF/CDC48p/Pas1p/TBP-1 family through heterologous expression in yeast. FEBS Lett. 351, 286–290. YOSHIMORI, T., YAMAGATA, F., YAMAMOTO, A., MIZUSHIMA, N., KABEYA, Y., NARA, A., MIWAKO, I., OHASHI, M., OHSUMI, M., and OHSUMI, Y. (2000). The mouse SKD1, a homologue of yeast Vps4p, is required for normal endosomal trafﬁcking and morphology

98

99

100

101

102

103

104

105

106

in mammalian cells. Mol. Biol. Cell 11, 747–763. BABST, M., WENDLAND, B., ESTEPA, E. J., and EMR, S. D. (1998). The Vps4p AAA ATPase regulates membrane association of a Vps protein complex required for normal endosome function. Embo J. 17, 2982–2993. FUJITA, H., YAMANAKA, M., IMAMURA, K., TANAKA, Y., NARA, A., YOSHIMORI, T., YOKOTA, S., and HIMENO, M. (2003). A dominant negative form of the AAA ATPase SKD1/VPS4 impairs membrane trafﬁcking out of endosomal/lysosomal compartments: class E vps phenotype in mammalian cells. J. Cell Sci. 116, 401–414. SACHSE, M., STROUS, G. J., and KLUMPERMAN, J. (2004). ATPase-deﬁcient hVPS4 impairs formation of internal endosomal vesicles and stabilizes bilayered clathrin coats on endosomal vacuoles. J. Cell Sci. 117, 1699–1708. DUPRE, S., URBAN-GRIMAL, D., and HAGUENAUER-TSAPIS, R. (2004). Ubiquitin and endocytic internalization in yeast and animal cells. Biochim. Biophys. Acta 1695, 89–111. INGHAM, R. J., GISH, G., and PAWSON, T. (2004). The Nedd4 family of E3 ubiquitin ligases: functional diversity within a common modular architecture. Oncogene 23, 1972–1984. GALAN, J. M., MOREAU, V., ANDRE, B., VOLLAND, C., and HAGUENAUER-TSAPIS, R. (1996). Ubiquitination mediated by the Npi1p/Rsp5p ubiquitin-protein ligase is required for endocytosis of the yeast uracil permease. J. Biol. Chem. 271, 10946–10952. SPRINGAEL, J. Y. and ANDRE, B. (1998). Nitrogen-regulated ubiquitination of the Gap1 permease of Saccharomyces cerevisiae. Mol. Biol. Cell 9, 1253–1263. WANG, G., MCCAFFERY, J. M., WENDLAND, B., DUPRE, S., HAGUENAUER-TSAPIS, R., and HUIBREGTSE, J. M. (2001). Localization of the Rsp5p ubiquitin-protein ligase at multiple sites within the endocytic pathway. Mol. Cell Biol. 21, 3564–3575. DUNN, R., KLOS, D. A., ADLER, A. S., and HICKE, L. (2004). The C2 domain of the

References

107

108

109

110

111

112

113

114

Rsp5 ubiquitin ligase binds membrane phosphoinositides and directs ubiquitination of endosomal cargo. J. Cell Biol. 165, 135–144. MORVAN, J., FROISSARD, M., HAGUENAUER-TSAPIS, R., and URBAN-GRIMAL, D. (2004). The ubiquitin ligase Rsp5p is required for modiﬁcation and sorting of membrane proteins into multivesicular bodies. Trafﬁc 5, 383– 392. BLONDEL, M. O., MORVAN, J., DUPRE, S., URBAN-GRIMAL, D., HAGUENAUER-TSAPIS, R., and VOLLAND, C. (2004). Direct sorting of the yeast uracil permease to the endosomal system is controlled by uracil binding and Rsp5p-dependent ubiquitylation. Mol. Biol. Cell 15, 883–895. SAKATA, T., SAKAGUCHI, H., TSUDA, L., HIGASHITANI, A., AIGAKI, T., MATSUNO, K., and HAYASHI, S. (2004). Drosophila Nedd4 regulates endocytosis of notch and suppresses its ligand-independent activation. Curr. Biol. 14, 2228–2236. ROTIN, D., KANELIS, V., and SCHILD, L. (2001). Trafﬁcking and cell surface stability of ENaC. Am. J. Physiol. Renal Physiol. 281, F391–F399. PERRY, W. L., HUSTAD, C. M., SWING, D. A., O’SULLIVAN, T. N., JENKINS, N. A., and COPELAND, N. G. (1998). The itchy locus encodes a novel ubiquitin protein ligase that is disrupted in a18H mice. Nat. Genet. 18, 143–146. MARCHESE, A., RAIBORG, C., SANTINI, F., KEEN, J. H., STENMARK, H., and BENOVIC, J. L. (2003). The E3 ubiquitin ligase AIP4 mediates ubiquitination and sorting of the G protein-coupled receptor CXCR4. Dev. Cell 5, 709–722. QIU, L., JOAZEIRO, C., FANG, N., WANG, H. Y., ELLY, C., ALTMAN, Y., FANG, D., HUNTER, T., and LIU, Y. C. (2000). Recognition and ubiquitination of Notch by Itch, a hect-type E3 ubiquitin ligase. J. Biol. Chem. 275, 35734–35737. ANGERS, A., RAMJAUN, A. R., and MCPHERSON, P. S. (2004). The HECT domain ligase itch ubiquitinates endophilin and localizes to the trans-Golgi network and endosomal system. J. Biol. Chem. 279, 11471–11479.

115 KATZ, M., SHTIEGMAN, K., TAL-OR, P., YAKIR, L., MOSESSON, Y., HARARI, D., MACHLUF, Y., ASAO, H., JOVIN, T., SUGAMURA, K., and YARDEN, Y. (2002). Ligand-independent degradation of epidermal growth factor receptor involves receptor ubiquitylation and Hgs, an adaptor whose ubiquitin-interacting motif targets ubiquitylation by Nedd4. Trafﬁc 3, 740–751. 116 POLO, S., SIGISMUND, S., FARETTA, M., GUIDI, M., CAPUA, M. R., BOSSI, G., CHEN, H., De CAMILLI, P., and DI FIORE, P. P. (2002). A single motif responsible for ubiquitin recognition and monoubiquitination in endocytic proteins. Nature 416, 451–455. 117 THIEN, C. B. F. and LANGDON, W. Y. (2001). Cbl: many adaptions to regulate protein tyrosine kinases. Nature Rev. 2, 294–305. 118 PESCHARD, P. and PARK, M. (2003). Escape from Cbl-mediated downregulation: a recurrent theme for oncogenic deregulation of receptor tyrosine kinases. Cancer Cell 3, 519–523. 119 PESCHARD, P., FOURNIER, T. M., LAMORTE, L., NAUJOKAS, M. A., BAND, H., LANGDON, W. Y., and PARK, M. (2001). Mutation of the c-Cbl TKB domain binding site on the Met receptor tyrosine kinase converts it into a transforming protein. Mol. Cell 8, 995–1004. 120 LEVKOWITZ, G., WATERMAN, H., ZAMIR, E., KAM, Z., OVED, S., LANGDON, W. Y., BEGUINOT, L., GEIGER, B., and YARDEN, Y. (1998). c-Cbl/Sli-1 regulates endocytic sorting and ubiquitination of the epidermal growth factor receptor. Genes Dev. 12, 3663–3674. 121 THIEN, C. B. F., WALKER, F., and LANGDON, W. Y. (2001). Ring ﬁnger mutations that abolish c-Cbl directed polyubiquitination and down regulation of the EGF receptor are insufﬁcient for cell transformation. Mol. Cell 7, 355–365. 122 DUAN, L., MIURA, Y., DIMRI, M., MAJUMDER, B., DODGE, I. L., REDDI, A. L., GHOSH, A., FERNANDES, N., ZHOU, P., MULLANE-ROBINSON, K., RAO, N., DONOGHUE, S., ROGERS, R. A., BOWTELL, D., NARAMURA, M., GU, H., BAND, V.,

25

26 Protein Sorting to the Lysosome

123

124

125

126

127

128

129

130

and BAND, H. (2003). Cbl-mediated ubiquitinylation is required for lysosomal sorting of epidermal growth factor receptor but is dispensable for endocytosis. J. Biol. Chem. 278, 28950–28960. de MELKER, A. A., van der HORST, G., and BORST, J. (2004). Ubiquitin ligase activity of c-Cbl guides the epidermal growth factor receptor into clathrin-coated pits by two distinct modes of Eps15 recruitment. J. Biol. Chem. 279, 55465–55473. HUANG, F. and SORKIN, A. (2005). Grb2-mediated Recruitment of the RING Domain of Cbl to the EGF Receptor Is Essential and Sufﬁcient to Support Receptor Endocytosis. Mol. Biol. Cell. 16, 1268–1281. PETRELLI, A., GILESTRO, G. F., LANZARDO, S., COMOGLIO, P. M., MIGONE, N., and GIORDANO, S. (2002). The endophilin-CIN85-Cbl complex mediates ligand-dependent downregulation of c-Met. Nature 416, 187–190. SOUBEYRAN, P., KOWANETZ, K., SZYMKIEWICZ, I., LANGDON, W. Y., and DIKIC, I. (2002). Cbl-CIN85-endophilin complex mediates ligand-induced downregulation of EGF receptors. Nature 416, 183–187. SCHMITZ, C., KINNER, A., and KOLLING, R. (2005). The deubiquitinating enzyme Ubp1 affects sorting of the ABC-transporter Ste6 in the endocytic pathway. Mol. Biol. Cell. 16, 1319– 1329. KEE, Y., LYON, N., and HUIBREGTSE, J. M. (2005). The Rsp5 ubiquitin ligase is coupled to and antagonized by the Ubp2 deubiquitinating enzyme. Embo J. 24, 2414–2424. PAPA, F. R., AMERIK, A. Y., and HOCHSTRASSER, M. (1999). Interaction of the Doa4 deubiquitinating enzyme with the yeast 26S proteasome. Mol. Biol. Cell 10, 741–756. PAPA, F. R. and HOCHSTRASSER, M. (1993). The yeast DOA4 gene encodes a deubiquitinating enzyme related to a product of the human tre-2 oncogene. Nature 366, 313–319.

131 SWAMINATHAN, S., AMERIK, A. Y., and HOCHSTRASSER, M. (1999). The Doa4 deubiquitinating enzyme is required for ubiquitin homeostasis in yeast. Mol. Biol. Cell 10, 2583–2594. 132 NAVIGLIO, S., MATTECUCCI, C., MATOSKOVA, B., NAGASE, T., NOMURA, N., DI FIORE, P. P., and DRAETTA, G. F. (1998). UBPY: a growth-regulated human ubiquitin isopeptidase. Embo J. 17, 3241–3250. 133 KATO, M., MIYAZAWA, K., and KITAMURA, N. (2000). A de-ubiquitinating enzyme UBPY interacts with the SH3 domain of Hrs binding protein via a novel binding motif Px(V/I)(D/N)RxxKP. J. Biol. Chem. 275, 37481–37487. 134 KANEKO, T., KUMASAKA, T., GANBE, T., SATO, T., MIYAZAWA, K., KITAMURA, N., and TANAKA, N. (2003). Structural insight into modest binding of a non-PXXP ligand to the signal transducing adaptor molecule-2 Src homology 3 domain. J. Biol. Chem. 278, 48162–48168. 135 MIZUNO, E., IURA, T., MUKAI, A., YOSHIMORI, T., KITAMURA, N., and KOMADA, M. (2005). Regulation of EGF receptor down-regulation by UBPY-mediated deubiquitination at endosomes. Mol. Biol. Cell 16, 5163–5174. 136 BOWERS, K., PIPER, S. C., EDELING, M. A., GRAY, S. R., OWEN, D. J., LEHNER, P. J., and LUZIO, J. P. (2005). Degradation of endocytosed epidermal growth factor and virally-ubiquitinated MHC class I is independent of mammalian ESCRTII. J. Biol. Chem. 281, 5094–5105. 137 SOARES, L., SEROOGY, C., SKRENTA, H., ANANDASABAPATHY, N., LOVELACE, P., CHUNG, C. D., ENGLEMAN, E., and FATHMAN, C. G. (2004). Two isoforms of otubain 1 regulate T cell energy via GRAIL. Nat. Immunol. 5, 45–54. 138 WU, X., YEN, L., IRWIN, L., SWEENEY, C., and CARRAWAY, K. L., 3rd. (2004). Stabilization of the E3 ubiquitin ligase Nrdp1 by the deubiquitinating enzyme USP8. Mol. Cell Biol. 24, 7748–7757. 139 GNESUTTA, N., CERIANI, M., INNOCENTI, M., MAURI, I., ZIPPEL, R., STURANI, E., BORGONOVO, B., BERRUTI, G., and MARTEGANI, E. (2001). Cloning and

References

140

141

142

143

144

145

characterization of mouse UBPy, a deubiquitinating enzyme that interacts with the ras guanine nucleotide exchange factor CDC25(Mm)/Ras-GRF1. J. Biol. Chem. 276, 39448–39454. TANAKA, N., KANEKO, K., ASAO, H., KASAI, H., ENDO, Y., FUJITA, T., TAKESHITA, T., and SUGAMURA, K. (1999). Possible involvement of a novel STAM-associated molecule “AMSH” in intracellular signal transduction mediated by cytokines. J. Biol. Chem. 274, 19129–19135. MAYTAL-KIVITY, V., REIS, N., HOFMANN, K., and GLICKMAN, M. H. (2002). MPN+, a putative catalytic motif found in a subset of MPN domain proteins from eukaryotes and prokaryotes, is critical for Rpn11 function. BMC Biochem. 3, 28. VERMA, R., ARAVIND, L., OANIA, R., MCDONALD, W. H., YATES, J. R., 3rd, KOONIN, E. V., and DESHAIES, R. J. (2002). Role of Rpn11 metalloprotease in deubiquitination and degradation by the 26S proteasome. Science 298, 611–615. MCCULLOUGH, J., CLAGUE, M. J., and URBE´ , S. (2004). AMSH is an endosome-associated ubiquitin isopeptidase. J. Cell Biol. 166, 487–492. ISHII, N., OWADA, Y., YAMADA, M., MIURA, S., MURATA, K., ASAO, H., KONDO, H., and SUGAMURA, K. (2001). Loss of neurons in the hippocampus and cerebral cortex of AMSH-deﬁcient mice. Mol. Cell Biol. 21, 8626–8637. ASADA, H., ISHII, N., SASAKI, Y., ENDO, K., KASAI, H., TANAKA, N., TAKESHITA, T.,

146

147

148

149

150

151

TSUCHIYA, S., KONNO, T., and SUGAMURA, K. (1999). Grf40, A novel Grb2 family member, is involved in T cell signaling through interaction with SLP-76 and LAT. J. Exp. Med. 189, 1383–1390. ITOH, F., ASAO, H., SUGAMURA, K., HELDIN, C. H., ten DIJKE, P., and ITOH, S. (2001). Promoting bone morphogenetic protein signaling through negative regulation of inhibitory Smads. Embo J. 20, 4132–4142. LI, H. and SETH, A. (2004). An RNF11: Smurf2 complex mediates ubiquitination of the AMSH protein. Oncogene 23, 1801–1808. ROCCA, A., LAMAZE, C., SUBTIL, A., and DAUTRY-VARSAT, A. (2001). Involvement of the ubiquitin/proteasome system in sorting of the Interleukin 2 receptor b chain to late endocytic compartments. Mol. Biol. Cell 12, 1293–1301. van KERKHOF, P., GOVERS, R., ALVES DOS SANTOS, C. M., and STROUS, G. J. (2000). Endocytosis and degradation of the growth hormone receptor are proteasome-dependent. J. Biol. Chem. 275, 1575–1580. HAMMOND, D. E., URBE´ , S., VANDE WOUDE, G. F., and CLAGUE, M. J. (2001). Downregulation of MET, the receptor for hepatocyte growth factor. Oncogene 20, 2761–2770. COLLEDGE, M., SNYDER, E. M., CROZIER, R. A., SODERLING, J. A., JIN, Y., LANGEBERG, L. K., LU, H., BEAR, M. F., and SCOTT, J. D. (2003). Ubiquitination regulates PSD-95 degradation and AMPA receptor surface expression. Neuron 40, 595–607.

27

1

Regulation of Proteolysis at the Proteasome Portal Monika Bajorek, and Michael H. Glickman

Technion — Israel Institute of Technology, Haifa, Israel

Originally published in: Protein Degradation, Volume 2. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31130-0 Abstract

The proteolytic active sites of the 26S proteasome are sequestered within the central chamber of its 20S catalytic core particle. Access to this chamber is through a narrow channel deﬁned by the outer alpha subunits. An intricate lattice of interactions anchors the N-termini of these alpha subunits, blocking access to the channel in free 20S core particles of eukaryotes. Entry of substrates can be enhanced by attachment of activators or regulatory particles to the proteolytic 20S core. Regulatory particles rearrange the blocking residues to form an open pore and promote substrate entry into the proteolytic chamber. Channel gating is apparently partially rate limiting for proteasome activity, as facilitating substrate entry in the open channel state leads to enhanced overall proteolysis rates. Interestingly, some substrates, particularly hydrophobic ones, can activate gate opening themselves, thus facilitating their own destruction. Properties of channel gating and the interactions required to maintain stable closed and open conformations and their consequences for proteasome function are discussed.

1 Background

In eukaryotes, the 26S proteasome hydrolyzes most nuclear, cytoplasmic, and endoreticulum (ER) proteins into peptides of varying lengths. Normally, substrates destined for elimination are ﬁrst covalently attached to multiple molecules of ubiquitin (Ub) – a process that is executed by a cascade of ubiquitinating enzymes speciﬁc for each class of substrate – and then recognized by the 26S proteasome, unfolded, translocated into the proteolytic chamber, and irreversibly degraded [1–3]. The somewhat simpler 20S proteasomes found in archaea and some bacteria (actinomycetes) can degrade and remove non-ubiquitinated proteins in a very similar Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Regulation of Proteolysis at the Proteasome Portal

manner, though recognition of substrates and anchoring them to the proteasome while they are prepared for degradation probably differ due to lack of the ubiquitin tag in these organisms [4, 5]. In either case, substrates are threaded through a narrow channel leading into the isolated internal chamber where the proteolytic active sites are located. A gated porthole at the entrance to this channel may play a deﬁned role in controlling both the nature of preferred substrates and the rate at which they enter the proteasome lumen where they are irreversibly hydrolyzed (Figure 1). This chapter will focus on this gate and mechanisms of maintaining opened or closed states. The 26S proteasome is composed of two sub-complexes: the 20S core particle of the proteasome (CP) where proteolysis takes place and a 19S regulatory particle (RP) that prepares substrates for entry into the CP. A detailed description of proteasome structure and associated activities can be found in other chapters in this volume, as well as in many detailed reviews [6–15]. Pertinent to understanding regulation of substrate entry, the 20S CP is a cylindrical structure composed of four stacked heptameric rings engendering a sequestered proteolytic chamber. Each of the two outer rings is composed of seven α subunits, and each of the two identical inner rings is formed from seven β subunits. The β rings contain the proteolytic active sites, while the outer α rings deﬁne the channel leading into the internal proteolytic chamber [16–19]. Overall, the 20S CP forms a α 7 β 7 β 7 α 7 barrel structure [6]. The α and β rings of archaeal 20S proteasomes are made up of seven copies each of a single α or β subunit, giving the structure as a whole a sevenfold symmetry along the central axis. In contrast, there are seven distinct subunits in each ring of the 20S CP from eukaryotes. When visualized from outside, these subunits are labeled counterclockwise α1 through α7 and β1 through β7, respectively. Although the seven α or β subunits within the 20S CP of each species are structurally almost superimposable, their sequence identities are usually in the 20–40% range (see Ref. [7] and references therein). These differences are probably of signiﬁcance as they are well maintained; sequences of orthologous subunits from different species are more than 55% identical [20, 21]. The puriﬁed 20S CP in its so-called latent form can slowly hydrolyze short or unstructured polypeptides as well as some proteins with hydrophobic or misfolded patches [22–25]. There is increasing evidence that degradation of unstructured nonubiquitinated proteins by the 20S CP may play biological roles in vivo as well [22, 26– 31]. To degrade ubiquitinated substrates, attachment of the ATPase-containing 19S RP to the surface of the α ring of the 20S CP is required [25, 32]. Attachment of 19S RP enhances the basal peptidase activity of proteasomes as well [33, 34]. In archaea and select prokaryotes, ATPase rings such as the proteasome-activating nucleotidase (PAN) complex or the AAA ATPase ring complex (ARC) serve as rudimentary regulatory particles that enhance proteolysis by 20S proteasomes but probably do not affect peptidase rates [35–38]. One manner by which such ATPase-containing regulatory particles activate proteolysis is by unfolding substrates and translocating them into the proteolytic chamber [36, 39–45]. However, other regulatory particles can also inﬂuence the proteolytic activity of the proteasome in an ATP-independent manner. For example, a number of non-ATPase activators – such as PA28, PA26,

1 Background

3

4 Regulation of Proteolysis at the Proteasome Portal

and 11S Reg complexes – attach to the 20S CP and enhance peptidase, but not proteolysis, rates [46–53]. Other conditions such as exposure to low ionic strength, sodium dodecyl sulfate (SDS), or small hydrophobic peptides can also activate hydrolysis rates and give rise to an activated 20S CP [18, 22, 24, 34, 54, 55]. These results (together with other observations) suggest that the latent 20S CP is found in a self-imposed repressed state and that it is possible to activate the intrinsic peptidase activity in an ATP-independent manner. Structural analysis revealed the nature of this auto-inhibition by identifying a gated channel leading substrates into the proteolytic chamber [17–19]. Alleviation of inhibition by opening the substrate channel for trafﬁc is a basic feature of proteasomes, a key property necessary to understand their function.

2 The Importance of Channel Gating

In order to protect cells from mistaken degradation of random proteins, entry into the proteasome lumen must be strictly regulated. Indeed, access to the channel in ←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− Fig. 1 A porthole into the proteasome. Top view presentations showing the surface structure of the " ring of free 20S CPs from various preparations: (A) mammalian, (B) mammalian in theoretical open state, (C) yeast (S. cerevisiae), (D) yeast in theoretical fully open state, (E) the "3N mutant from yeast, (F) yeast in open conformation imposed by attachment of the PA26S activator, (G) archaea (T. acidophilum), (H) bacteria (Rhodococcus erythropolis). Structures depicted in A, C, E, F, G, and H are the actual 2D determinations extracted from the crystal structure deposited in the Protein Data Bank (PDB) and visualized as a surface view with the Viewerlite program. Acidic residues are colored in red, alkali in blue, and hydrophobic in white. Note the dominance of acidic residues in the pore region of the closed state in yeast and mammalian 20S CPs. These structures indicate that the gross surface structure of eukaryotic proteasomes is remarkably similar, with slight structural divergence apparent in the archaeal and bacterial versions. The N-terminus upstream of threonine 13 in the archaeal complex (the gray region shown in Figure 2) is disordered and thus does not show up in the electron density map, giving the appearance of an open confor-

mation (G). Structures shown in B and D are models in which the electron density of the corresponding tail residues in each " subunit was deleted to mimic an open conformation. Complete removal of these tail segments unveils a pore in the open state. The actual crystal structure determination of the "3N 20S CP indicates that "3 is a pivotal subunit in controlling channel gating. Deletion of the "3 tail alone (E) causes disordering in neighboring subunits (up to the red arrow in Figure 2); however, some obstruction remains when compared with the theoretical fully open state (D). Deletion of the two opposing tails from the "3 and "7 subunits generates a fully activated complex, apparently due to removal of this residual obstruction [22]. Interestingly, attachment of PA26 to the "-ring surface rearranges the tail regions of each subunit, coupled with signiﬁcant disordering (up to the blue arrow in Figure 2), thus imposing an open conformation (F). By deleting the electron density of the PA26 chains and focusing only on the structure of the 20S subunits, we depict in panel F the actual situation that occurs upon PA26 binding to 20S from yeast. Additional details can be found in the original publications [16–19, 55, 59].

2 The Importance of Channel Gating 5

the latent 20S CP of eukaryotes is restricted by the N-termini of the seven α subunits [17–19]. Each tail assumes a unique conformation while pointing inwards to the center of the ring [7, 9, 10]. Figures 1A and 1C describe the surface of the latent 20S CP as it may appear to an approaching substrate; the interlaced residues in latent proteasomes form a sealed surface obstructing passage of proteins and even short peptides. This property is a common feature of eukaryotic proteasomes and is well conserved between yeast and mammalian complexes. The blocking residues at the entrance to the proteolytic channel probably account for the repressed proteolytic activity of the latent 20S CP of eukaryotes [18, 56]. The behavior of archaeal proteasomes is somewhat different from that of the proteasomes of eukaryotes. In contrast to the sealed chamber in the eukaryotic complex, crystal structures of 20S proteasomes obtained from archaeal organisms distinctly ﬁnd the proteasome in an open state. Disorder in the conformation of the ﬁrst 12 amino acids of each α subunit creates a pore in the center of the α ring contiguous with the channel leading into the lumen (Figure 1G). This open state accounts for enhanced peptidase rates measured for this complex [16, 36, 57]. Despite the appearance of an open pore in the archaeal 20S CP (Figure 1), dynamic conformations of the α-subunit tails may partially restrict passage of intact proteins through the pore, necessitating mechanisms for activation of protease activity. For instance, by locking these residues in a stable, open conformation, regulatory complexes found in these organisms (such as PAN) are able to accelerate proteolysis of full-sized proteins [36, 43, 55, 57, 58]. Certain conditions might promote switching of the N-termini from an unstructured into a structured, open conformation without requiring a regulatory complex [57]. Apparently, a similar situation occurs for 20S proteasomes present in bacteria [59]. The N-termini encompassing the ﬁrst eight amino acids of the subunits in the α ring of the Rhodococcus proteasome are disordered (Figure 2), creating the appearance of an open pore (Figure 1H). The ARC regulatory complex found in these organisms probably stimulates proteolytic activity by anchoring these tails into a static, open conformation, similarly to the role of PAN in archaeal organisms. Eukaryotic 20S CPs can also be found in an open conformation. Repeated freezethawing of puriﬁed 20S CPs; mild chemical treatments such as exposure to low ionic strength, to low levels of sodium dodecyl sulfate (SDS), or to short hydrophobic peptides; and attachment of regulatory complexes and even mutations in residues near the central pore all activate hydrolysis rates [18, 22, 24, 34, 54, 55]. Presumably, all these treatments lead to disordering in pore residues involved in gating the channel, thus opening up a porthole into the 20S CP (Figure 1B,D). Rearrangement of channel-blocking residues results in facilitated substrate access into the proteolytic chamber and activation of proteasomes. Activated 20S CPs can easily hydrolyze unfolded or hydrophobic proteins, in some instances more rapidly even than intact 26S holoenzymes [22, 23]. It should be noted that free 20S CPs can spontaneously switch between the latent “closed” and activated “open” conformations [60]; however, under physiological conditions it appears that the majority of free 20S CPs from eukaryotes are found in a closed and latent state.

6 Regulation of Proteolysis at the Proteasome Portal

Fig. 2 Sequence alignment of " subunit Ntermini. N-termini of the seven different " subunits from S. cerevisiae compared to archaeal and bacterial homologues. The seven a subunits ("1-"7) of yeast (Sc) are shown along with the single " subunit found in bacteria (Re) and archaea (Ta). In normal text, to the left of the bold region lie the tail sequences, which differ from one " subunit to the other. Residue numbers for the yeast and bacteria " subunits are assigned based on the alignment to archaea. Residues in the N-terminus to the arrow are disordered in "3N mutant (red arrow) and upon attachment of PA26S (blue arrow). The conserved residues forming the so-called canonical cluster (YDR-P-Y) documented

to play a role in stabilizing the closed or open conformations are underlined. The Nterminal sequence up to the ﬁrst " helical structure of each CP " subunit (as determined by the crystal structure of the yeast CP [17, 18]) is shown in light gray. This region is homologous to the disordered segments in the N-terminal regions of the " subunit from T. acidophilum [16, 57]. The YD(R) motif in each tail is underlined. Note that in all subunits from yeast a tyrosine residue is present in the same location. In six of the seven subunits, an aspartate follows, and in three, an arginine completes the YDR motif. The remainder of the tail region is divergent between subunits.

Similar to the situation described above for archaeal proteasomes, a distinction can be made between the appearance of an open channel due to disordering in residues lining the pore region and one in which the blocking tails are secured in an open structure. Structural determination of a 20S CP–PA26 complex depicts the α subunit N-termini pointing away from the center of the ring, opening up an unobstructed porthole into the channel [55] (see also Figure 1). The conformation differences between the closed and open states of N-termini could explain how regulatory particles activate proteolytic activity by rearranging the blocking residues to facilitate substrate entry [43, 54, 55]. Regulatory complexes such as the 19S RP participate in channel gating and facilitate substrate entry. This activates proteolysis and allows the resulting proteasome holoenzyme to fulﬁll its role in regulated protein degradation [7]. Indeed, under standard growth conditions, the majority of proteasomes in yeast cells are found as 26S holoenzymes. That said, 20S CPs are abundant in certain cases and are found

3 A Porthole into the Proteasome 7

regularly in mammalian tissue. Given that 20S core particles are slower than 26S holoenzymes at hydrolyzing most test substrates and are unable to proteolyze polyubiquitinated proteins, it is unclear what the biological function of the 20S CP is, or whether it is an important player in cellular protein breakdown. Abating bulk protein degradation upon proteasome disassembly may be a requirement for survival under certain stress conditions. For instance, prolonged starvation, oxidative stress, and severe heat-induced damage result in dissociation of proteasome holoenzymes and elevated levels of 20S CPs [22, 28, 29, 61]. There is some evidence that the catalytically repressed 20S CP can serve as a reservoir of proteasome components for reassembly when resumption of proteolysis is needed. Nevertheless, free 20S CPs may play limited roles in degradation of unstructured or non-ubiquitinated proteins under both normal growth and stress conditions [22, 26, 28–31].

3 A Porthole into the Proteasome 3.1 The Closed State

The ﬁrst 12 residues in the archaeal complex, right up to the ﬁrst α helix in the protein (Figure 2), are naturally disordered, giving the impression of an open channel (Figure 1G). Similar disordering is found in the equivalent sections of α subunits in the bacterial 20S proteasome puriﬁed from Rhodococcus (Figure 1H). In comparison, the corresponding N-termini of the seven α subunits in eukaryotic complexes point towards the center of the ring, sealing the entrance to the proteolytic channel (Figure 1A,C). In eukaryotes, the paralogous subunits within the α ring show structural and sequence similarities over the bulk of the protein, yet diverge at their amino-terminal region in both sequence and relative length (Figure 2). Precisely at the center of the ring, each tail accepts a unique conformation, stabilizing a well-deﬁned and non-symmetric closed conﬁguration [7] (see also Figure 3). The tail of α3 is somewhat distinct from the others in that it points directly across the surface of the α ring towards the center, maintaining close contact to every other α subunit. The importance of these tail regions is highlighted by their extreme conservation across eukaryotes; while each tail is highly conserved in different species, the corresponding regions are divergent from one subunit to another [7]. These properties suggest that the N-termini play a critical structural role that has been maintained in core particles in all eukaryotes, and it is precisely their differences that are integral to their function. Low-energy bonds formed between speciﬁc tail residues are critical for stabilizing the open or closed conformations. For example, in latent 20S CPs, aspartate at position 9 in the N-terminus of α3 contacts both tyrosine 8 and arginine 10 in neighboring α4 [18]. A salt bridge is formed between the carboxylate group of aspartate 9 in α3 and the guanidinium group of arginine 10 in α4, simultaneous with a hydrogen bond linking aspartate 9 of α3 with tyrosine 8 of α4 [18]. Similar

8 Regulation of Proteolysis at the Proteasome Portal

Fig. 3 A gate-and-latch system determines open and closed states of the substrate channel. Yeast 20S CP in a latent closed conformation (A) and in an open state imposed by the PA26S activator (B). At left a top view of the seven-member " ring of the 20S CP is shown. All " subunits are color coded, starting with "1 (light blue) on top and running counterclockwise to "7 (red). On the right, a side view focusing only on two opposing subunits – "3 (pink) and "7 (red) – highlights the conformational

switch that occurs in the N-termini (black) between the closed state (A) to an open state (B) upon binding of PA26. Note that the N-terminal tails of "3 and "7 adopt different conformations in the closed state and point inwards to block access through the channel (A). In contrast, in the open state induced by PA26S, the tails adopt a new ordered conformation, pointing away from the channel region. A portion of the N-terminus (up to the blue arrow in Figure 2) is disordered and invisible in the crystal structure.

bonds probably link the analogous residues in mammalian 20S CPs [19]. Embedded within the N-terminal segments of most α subunits is a short consensus sequence: Tyr8-Asp9-Arg10 or “the YDR motif ” (Figure 2). Conservation of tyrosine at position 8 is absolute among subunits in yeast (and is invariable at this location in most subunits of other organisms as well); aspartate at position 9 is present in the tail of all yeast subunits except for α2, while conservation of arginine as residue number 10 is less strict. The direct contacts formed between these residues in adjacent subunits may explain their correlated evolutionary conservation. Interactions involving YDR residues could be critical for maintaining distinct open and closed conformations of 20S proteasomes. Finding YDR residues in all subunits that form the α ring makes it somewhat puzzling that crystal structure determination did not pick out similar contacts between other neighboring subunits in the pore region. This raises the possibility that the interaction between α3 and α4 plays a unique and central role in maintaining the closed conformation of the proteasome. Support for the pivotal role of α3 in gating the 20S CP channel was provided upon truncation of the tail region of α3. Truncation of the N-terminus of the α3 subunit in yeast (the α3N mutant) resulted in a puriﬁed 20S CP that was found in the open pore conformation

3 A Porthole into the Proteasome 9

(Figures 1 and 3). Furthermore, removal of the N-terminus of α3 caused disordering in neighboring subunits concomitant with stimulation of 20S CP peptidase activity. These results indicate that the tail of α3 is important for stabilizing neighboring tails in the closed conformation. Moreover, an aspartate-to-alanine substitution in the YDR motif of α3 (the α3 D9A mutant) appears to increase peptidase activity of puriﬁed 20S CP in vitro, on par with the activation observed upon deletion of the entire tail region in α3N [18]. Both mutations associated with α3 point to a functional signiﬁcance of the YDR motif in stabilization of the closed state of the gate. Interestingly, the YDR sequence is found intact in the α subunits from various archaea, even though the tail regions are not anchored in the latent state of 20S proteasomes from these organisms [16, 57]. This observation points to a wider role for α-subunit tail interactions in deﬁning proteasome conformation. 3.2 The Open State

In order for substrates to enter the proteolytic chamber, and most likely for products to exit as well, the blocking N-terminal residues of the α subunits in the closed state must be rearranged. Rearrangement obviously necessitates breaking of the interactions that anchor the tails in the closed conformation, while forming competing interactions to stabilize them in an open conformation. For example, removal of the ﬁrst nine residues at the N-terminus of the α3 subunit in yeast breaks stabilizing interactions with neighboring subunits, causing signiﬁcant disordering in neighboring tails (Figure 2) and leading to an open pore roughly 13 Å across at the center of the α-ring surface (Figure 1F). Disordering alone is insufﬁcient to allow unobstructed entry through the pore, as no enhancement of protein degradation rates has been observed so far with proteasomes puriﬁed from this mutant [18, 22]. Each α subunit plays a unique role in gating. Deletion of the equivalent N-terminal tail of α7 does not signiﬁcantly increase the peptidase activity compared to wild type, pointing to a signiﬁcant role for α3 [22]. Because of its peripheral location at the α-ring surface, truncation of α7 alone may not result in sufﬁcient loss of order in neighboring tails to generate an opening wide enough for entry of small peptides. However, deletion of tails of two opposing α subunits (the α3α7N strain; [22]) may act synergistically to relieve hindrance of entry of proteins into the proteolytic chamber. Thus, gating of the proteolytic channel emerges as a pivotal property in regulating proteolysis rates. In fact, activated proteasomes may require more than just a disordered state of channel gating residues. Blocking residues may need to be removed (as in the truncation studies described in the preceding paragraph) or anchored into a stable, open conformation to relieve obstruction of the channel. Studies on archaeal proteasomes point to interactions involving YDR residues as inﬂuencing channel opening rather than stabilizing the closed state. While the N-terminal tails (12 amino acids) of each subunit of the α ring are disordered, they nevertheless occupy the pore region and impose a partial barrier to passage of protein substrates [16, 57]. The lack of deﬁned electron density reﬂects that under the experimental conditions

10 Regulation of Proteolysis at the Proteasome Portal

mutual interactions were not strong enough to anchor these tails into a single stable conformation, resulting in multiple possible conformations of these segments between individual molecules in the sample. Evidence that the YDR motif may play a role in stabilizing the open state was provided by a combined mutagenesis and biochemical study of archaeal proteasomes. Puriﬁed archaeal 20S proteasomes slowly hydrolyze GFP tagged with a short C-terminal extension, GFP-ssrA. The rate is accelerated upon addition of the archaeal proteasome activator PAN. However, PAN was unable to activate proteolysis of GFP-ssrA by 20S proteasomes from the archaeon T. acidophilum that were mutated at positions tyrosine 8 or arginine 9 (of the YDR motif in the N-tail regions) of their α subunit [55]. It was suggested that these residues are required to stabilize the tails in an open conformation that is preferred upon attachment of PAN. A stably open conformation was observed in proteasome assembly precursors as well. During proteasome biogenesis, the seven α subunits form an intermediate homomeric ring, the α 7 ring, which only then interacts with β subunits to yield the mature complex with α 7 β 7 composition [62]. In contrast to mature archaeal 20S CPs, structure determination of such an α 7 ring precursor from the archaeon Archaeoglobus fulgidus found the N-terminal segments anchored in a stable, open state [57]. In this conformation, the tail regions that contain the YDR motif adopted a helical structure motif and pointed away from the ring surface: tyrosine 8 of each subunit made a hydrogen bond with aspartate 9 of the preceding α subunit, whereas arginine 10 pointed inwards towards the central channel and did not partake in anchoring neighboring tails. The region N-terminal to tyrosine 8 of each subunit was disordered, creating the appearance of a pore roughly 13 Å in diameter contiguous with the substrate channel. In mature archaeal 20S CPs, the interactions between the N-termini of the α subunits appear to be broken, causing disordering in a greater portion of the tail regions up to residue number 12 (inclusive). These disordered tails, which are not stably anchored, partially block the pore and interfere with passage of proteins into the proteasome lumen. Eukaryotic proteasomes can be found in an open state as well. Crystallography analysis of a PA26–20S CP complex depicted the α ring in an ordered, open, symmetric conformation [55]. Attachment of PA26 induces all seven of the α subunit N-terminal tails to adopt an ordered conformation for residues 7–12 away from the center of the ring. A cluster of four highly conserved residues, Tyr8 and Asp9 (part of the YDR motif in the tail region) together with downstream residues Pro17 and Tyr26 (in the ﬁrst stable alpha helix; HO), is critical for the open state. Attachment of PA26 to the surface of the α ring repositions Pro17, which in turn induces a conformational change in tail segments to lift up and away from the center of the ring (Figure 2). In stark contrast to the closed state, the open state is remarkably symmetric, with all tails conforming to a similar structure held in place by similar interactions. In this open state, Asp9 forms a hydrogen bond with Tyr26 of the same subunit. Cooperativity between subunits is communicated via an additional hydrogen bond linking Asp9 of one subunit and Tyr8 of the preceding (clockwise) tail [55]. Consequently, the hydrogen bond that holds Asp9 and Tyr8 of α3 and α4, respectively, in the closed state must be broken and rearranged to allow for the open state in which

4 Facilitating Trafﬁc Through the Gated Channel 11

Asp9 of α3 now interacts with Tyr8 of α2. The term “proteasome gating” refers to switching between these two conformations. A principal role of regulatory particles is to promote channel opening by stabilizing one conformation over the other. The interactions occurring between different subunits in the pore region of the eukaryotic PA26–20S CP complex are quite similar to those found in the α 7 -ring precursor complex of archaea, which is also found in a stable, open conformation [57]. Asp9 in each subunit interacts with Tyr8 of the preceding (clockwise) tail in α 7 rings, leading to a symmetric, open structure. Asp9 of each α subunit also interacts with the downstream Tyr26 residue of the same subunit, repositioning the tail away from the center of the ring and up into the cavity of the docking proteasome activator complex (PA26 in this case). This conformation can be seen clearly in a side view of the 20S CP complex shown in Figure 3. The mesh of internal interactions among α subunits described in the preceding paragraph and the apparent lack of stable interactions between α-subunit tail residues with the proteasome activator suggest that a stable, open conformation is an intrinsic property of proteasomes and may explain how, under some conditions, 20S core particles can spontaneously adopt the activated form without need for attachment of an activator complex [60]. Nevertheless, comparative studies show that the 20S CP from Rhodococcus is also found in an open conformation [59], even though the α subunits in this organism do not contain any of the signature YD or PY residues in the tail or HO regions (Figures 1, 2). Apparently, the semblance of an open state does not absolutely require interactions among these residues, suggesting additional mechanisms for stabilizing the closed or open states and switching between them. Additional studies will have to be completed for a clear understanding of gating mechanisms.

4 Facilitating Trafﬁc Through the Gated Channel 4.1 Regulatory Complexes

As mentioned above, a number of ATP-independent activators are known to attach to the 20S CP and activate its peptidase and protease activities. These include the 11S Reg/PA28, PA26, and PA200 [46–53]. Attachment of PA28, for example, increases Vmax for hydrolysis of certain peptides by the 20S CP by up to 100-fold, but in contrast to ATPase-containing activators (such as the 19S RP or PAN), PA28 does not promote protein degradation by the 20S CP [46, 63]. Activation of peptidase activity by non-ATPase-containing regulators can be attributed to imposing an open-channel conformation and facilitated substrate entry upon attachment of the regulatory complex. For example, activation has been documented for the PA26–20S hybrid complex, formed in vitro from 20S CP from S. cerevisiae and PA26 from T. brucei [55]. S. cerevisiae apparently lacks natural homologues of this class of activators, but it is assumed that attachment of other symmetric activators induces

12 Regulation of Proteolysis at the Proteasome Portal

a similar conformational change. For example, the symmetric PAN complex found in archaea appears to drive formation of the open conformation of archaeal proteasomes in much the same manner [55]. Attachment of PAN expedites proteolysis by wild-type archaeal 20S proteasomes, yet is unable to activate proteasomes mutant in any of the channel cluster residues (Tyr8, Asp9, Pro17 and Tyr26; see Section 3.2). This result suggests that each residue in the conserved YD-P-Y cluster is critical for stabilizing the open conformation in activated proteasomes. Mutations in these residues may lead to disordered tails that are unable to “lock” into an open conformation even upon PAN attachment, thus impinging on substrate entry. Activation is not achieved merely by realigning the residues that obstruct trafﬁc through the alpha ring in the closed state, but necessitates locking the α tails into a stable, open conformation. So far, it is unclear whether asymmetric activators, such as the 19S RP, open the channel in a manner similar to that of the symmetric examples given by PA26 and PAN. However, evidence linking the 19S RP to gating can be deduced from a substitution mutation in the ATP-binding site of a single ATPase (RPT2) that severely affects peptidase activity of the proteasome, probably due to hampering the ability of the RP to properly gate the channel into the CP [56, 64]. This observation indicates that even the entry of small peptides – which do not need to be unfolded – can be controlled by the RP. Furthermore, a constitutively open-channel 20S CP generated upon deletions of tail residues in the α3 and α7 subunits (α3α7N mutant) exhibits activated peptidase activity, similar to that measured for 26S proteasome holoenzymes [22]. The implication is that attachment of the 19S RP realigns the α-subunit tails to facilitate passage of substrates, thus enhancing proteolysis rates. It should be emphasized, however, that in contrast to the homomeric rings found in PA26 or PAN, the 19S RP is a heterogeneous complex that contains six different ATPases. The two classes of regulators may induce different conformational changes on the sevenfold symmetry of the α ring. Furthermore, it has not yet been veriﬁed that the six ATPases indeed form a six-member ring at the base of the 19S RP. A limited subset of Rpt subunits have been found to come in direct contact with an α subunit (α2–Rpt4, α2–Rpt5, α4–Rpt4, α7–Rpt4, α1–Rpt6, α2–Rpt6, α4–Rpt2, α6–Rpt4; [65– 68]). The pair Rpt2–α3 has been shown to be involved in gating the channel into the CP [18, 56, 64], though gating may be controlled by additional Rpt–α subunit interactions. As there does not appear to be simple pairing of each Rpt with a single α subunit, it is possible that the conformation adopted by α-subunit tails in the 26S proteasome holoenzyme will differ from the “symmetric” open state observed for the PA26–20S CP hybrid. 4.2 Substrate-facilitated Trafﬁc

Recent studies show that some natively disordered proteins can enter the proteasome without assistance of ATP-dependent activators [26–28, 30]. The ability of latent 20S CP to catalyze cleavage of some unfolded proteins suggests that they may directly interact with the α ring to promote gating to facilitate their own entry.

5 Conclusions 13

This mechanism is not general: latent 20S CPs degrade most substrates slower than activated proteasomes [22]. Nevertheless, some substrates with unfolded domains or hydrophobic patches are degraded rapidly by latent 20S CP, faster even than by 26S holoenzymes [23]. Presumably, sequence motifs in the substrate interact with channel-gating residues in α subunits and aid in channel opening. For example, p21 and α-synuclein facilitate their own degradation and, when fused to stable and hard-to-degrade proteins such as GFP, promote their degradation as well indicating that gating sequences are transferable. Support for such a mechanism can be deduced from certain peptides that interact with the proteasome in a noncompetitive way to modulate the proteolytic activity of the proteasome [24, 69]. This stimulation was not observed for open-channel proteasomes (such as α3N or the PA26–20S CP complex), suggesting that they speciﬁcally interact with channel-gating residues and promote channel opening. Whether these interactions involve the YDR motif or the YD-P-Y cluster in α subunits, similar to the manner by which regulatory particles activate proteasome activity, has not been elucidated. Interestingly, the pore region of the 20S CP, in the closed state, exposes only hydrophobic or negatively charged side chains. No positively charged groups are present on the surface of the α ring in eukaryotes (Figure 1). Thus, as a substrate approaches the surface of the α ring, it “sees” a predominantly negatively charged surface. This may explain how the latent 20S CP discriminates between substrates and why certain unstructured substrates with hydrophobic or positively charge stretches may interact with gating residues in the pore region and facilitate their own translocation inwards, whereas others do not [23]. For example, hydrolysis of casein – which is highly phosphorylated and carries multiple negatively charged groups – is remarkably slow by latent 20S CP, yet can be dramatically accelerated upon channel opening possibly by removal of tail residues as in Figure 1E [22].

5 Conclusions

It appears that proteasome-dependent proteolysis is a regulated process that can be enhanced or inhibited under certain conditions. There are reports that the pro-teasome itself can be a target of such regulation [22, 29, 61, 70]. Indeed, enhancement of overall in vivo proteolysis rates observed in the open-channel mutant indicates that the proteasome may be partially rate limiting in the overall cascade of ubiquitin-dependent protein degradation [22]. Polyubiquitinated substrates must be stable enough, even if only transiently, to allow for competition between degradation and reversal of fate. Channel gating within 26S holoenzymes may participate in the delicate balance between proteolysis and rescue. A function of a gated channel leading into the CP is to impose inhibition during assembly of the mature CP. In the ﬁnal stage of CP assembly, selfcompartmentalization is achieved by the association of two α 7 β 7 half-CPs at the β-β interface. These half-CPs are inactive due to propeptides in the critical β subunits that mask their active site. As these half-CPs are joined, inhibition

14 Regulation of Proteolysis at the Proteasome Portal

by β-subunit N-termini is relieved by autolysis [17, 62], while inhibition by the blocking N-termini of the α subunits is imposed. Binding of the regulatory particles relieves this inhibition by opening the channel and thus activating proteolysis. There is increasing evidence that (at least in yeast) certain stress constitutions such as prolonged starvation or severe heat shock naturally promote proteasome dissociation into separate 20S CP and 19S RP subcomponents [22, 29, 61, 70]. These conditions may require repressed proteasome-dependent degradation for survival. One manner by which proteasome activity could be downregulated is by reinstating autoinhibition of the dissociated 20S CP. Indeed, the open-channel mutant that lacks the ability to enter the closed conformation exhibits low viability under conditions that promote proteasome disassembly [22]. An additional reason for a gated channel could be to regulate exit of products from the proteasome. It is possible that under normal conditions product release is slowed down by a gated channel in order to increase processivity or to decrease average peptide length. Most of these short peptides are quickly removed from the cytoplasm. Under certain conditions (such as during immune response) it might be beneﬁcial to produce peptides with other lengths or properties. For example, upon interferon-γ induction, attachment of PA28/11S Reg plays a role in antigen processing by altering the makeup of peptides generated by the hybrid proteasome 19S RP–20S CP–11S Reg complexes [52, 53, 71, 72]. In analogy to the distantly related PA26, PA28 probably attaches to the α-ring surface and rearranges the blocking N-termini, promoting the open-channel conformation. It is possible that the open state increases the exit rate of peptides generated in the proteolytic chamber and alters their makeup to ﬁt better antigen-presentation requirements.

References 1 GLICKMAN, M. H. and A. CIECHANOVER, The Ubiquitin-proteasome Proteolytic Pathway: Destruction for the sake of construction. Physiol. Rev., 2002. 82: p. 373–428. 2 PICKART, C. M., Mechanisms Underlying Ubiquitination. Annu. Rev. Biochem., 2001. 70: p. 503–533. 3 WEISSMAN, A. M., Themes and variations on ubiquitylation. Nature Rev. Cell Mol. Biol., 2001. 2 (3): p. 169–179. 4 MAUPIN-FURLOW, J. A., et al., Proteasomes in the archaea: from structure to function. Front. Biosci., 2000. 5: p. D837–D865. 5 De MOT, R., et al., Proteasomes and other self-compartmentalizing proeases in prokaryotes. Trends Microbiol., 1999. 7 (2): p. 88–92. 6 PICKART, C. M. and R. E. COHEN, Proteasomes and their kin: Proteases in

7

8

9

10

11

the machine Age. Nat. Rev. Mol. Cell Biol., 2004. 5 (3): p. 177–187. BAJOREK, M. and M. H. GLICKMAN, Proteasome regulatory particles: Keepers of the gates. Cell. Mol. Life Sci., 2004. 61 (13): p. 1579–1588. GUTERMAN, A. and M. H. GLICKMAN, Deubiquitinating enzymes are IN(trinsic to proteasome function). Curr. Prot. Pep. Sci., 2004. 5 (3): p. 201–210. GROLL, M. and T. CLAUSEN, Molecular shredders: how proteasomes fulﬁll their role. Current Opinion in Structural Biology, 2003. 13 (6): p. 665–673. GROLL, M. and R. HUBER, Substrate access and processing by the 20S proteasome core particle. Int. J. Biochem. Cell Biol., 2003. 35 (5): p. 606–616. GLICKMAN, M. H. and V. MAYTAL, Regulating the 26S proteasome., in Curr.

References

12

13

14

15

16

17

18

19

20

21

22

Top. Microbiol. and Immunol., P. ZWICKL and W. BAUMEISTER, Editors. 2002, Springer-Verlag: Germany. p. 43–72. ZWICKL, P. and E. SEEMUELLER, 20S proteasomes, in Curr. Topics Microbiol. Immunol., P. ZWICKL and W. BAUMEISTER, Editors. 2002, Springer-Verlag: Germany. p. 23–41. HEINEMEYER, W. and D. H. WOLF, Active sites and assembly of the 20S proteasome., in Proteasomes: The world of regulatory proteolysis, D. H. WOLF and W. HILT, Editors. 2000, Eurekah.com/ LANDES BIOSCIENCE Publishing Company: Georgetown, Texas USA. p. 48–70. BOCHTLER, M., et al., The Proteasome. Annu. Rev. Biophys. Biomol. Struct., 1999. 28: p. 295–317. VOGES, D., P. ZWICKL, and W. BAUMEISTER, THE 26S PROTEA-SOME: A Molecular Machine Designed for Controlled Proteolysis. Annu. Rev. Biochem, 1999. 68: p. 1015–1068. LOEWE, J., et al., Crystal structure of the 20S proteasome from the archeon T. acidophilum at 3.4 Angstrom resolution. Science, 1995. 268: p. 533–539. GROLL, M., et al., Structure of 20S proteasome from yeast at a 2.4 Angstrom resolution. Nature, 1997. 386: p. 463–471. GROLL, M., et al., A gated channel into the core particle of the proteasome. Nat. Struct. Biol., 2000. 7: p. 1062–1067. UNNO, M., et al., The structure of the mammalian 20S proteasome at 2.75A resolution. Structure, 2002. 10: p. 609–618. VOLKER, C. and A. LUPAS, Molecular Evolution of Proteasomes, in Curr. Top. Microbiol. and Immunol., P. ZWICKL and W. BAUMEISTER, Editors. 2002, Springer-Verlag: Berlin Heidelberg. p. 1–22. GILLE, C., et al., A Comprehensive View on Proteasomal Sequences: Implications for the Evolution of the Proteasome. Journal of Molecular Biology, 2003. 326 (5): p. 1437–1448. BAJOREK, M., D. FINLEY, and M. H. GLICKMAN, Proteasome Disassembly and Downregulation Is Correlated with Viability during Stationary Phase. Current Biology, 2003. 13 (13): p. 1140–1144.

23 LIU, C.-W., et al., Endoproteolytic activity of the proteasome. Science, 2003. 299: p. 408–411. 24 KISSELEV, A. F., D. KAGANOVICH, and A. L. GOLDBERG, Binding of hydrophobic peptides to several non-catalytic sites promotes peptide hydrolysis by all active sites of 20 S proteasomes. Evidence for peptide-induced channel opening in the alpha-rings. J. Biol. Chem., 2002. 277: p. 22260–22270. 25 GLICKMAN, M. H., et al., A subcomplex of the proteasome regulatory particle required for ubiquitin-conjugate degradation and related to the COP9/Signalosome and eIF3. Cell, 1998. 94 (5): p. 615–623. 26 ORLOWSKI, M. and S. WILK, Ubiquitin-independent proteolytic functions of the proteasome. Archives of Biochemistry and Biophysics, 2003. 415 (1): p. 1–5. 27 FORSTER, A. and C. P. HILL, Proteasome degradation: enter the substrate. Trends in Cell Biology, 2003. 13 (11): p. 550–553. 28 GRUNE, T., et al., Selective degradation of oxidatively modiﬁed protein substrates by the proteasome. Biochemical and Biophysical Research Communications, 2003. 305 (3): p. 709–718. 29 SHRINGARPURE, R., et al., Ubiquitin Conjugation Is Not Required for the Degradation of Oxidized Proteins by Proteasome. J. Biol. Chem., 2003. 278 (1): p. 311–318. 30 ASHER, G., et al., Mdm-2 and ubiquitin-independent p53 proteasomal degradation regulated by NQO1. PNAS, 2002. 99 (20): p. 13125–13130. 31 HOYT, M. A. and P. COFFINO, Ubiquitin-free routes into the proteasome. Cell. Mol. Life Sci., 2004. 61 (13): p. 1596–1600. 32 GUTERMAN, A. and M. H. GLICKMAN, Complementary roles for Rpn11 and Ubp6 in deubiquitination and proteolysis by the proteasome. J. Biol. Chem., 2004. 279 (3): p. 1729–1738. 33 ADAMS, G. M., et al., Formation of proteasome-PA700 complexes directly correlates with activation of peptidase activity. Biochem., 1998. 37: p. 12927–12932.

15

16 Regulation of Proteolysis at the Proteasome Portal

34 GLICKMAN, M. H., et al., The regulatory particle of the S. cerevisiae proteasome. Mol. Cell. Biol., 1998. 18: p. 3149– 3162. 35 FROHLICH, K. U., An AAA family tree. J. Cell Sci., 2001. 114: p. 1601–1602. 36 ZWICKL, P., et al., An archaebacterial ATPase, homologous to ATPases in the eukaryotic 26 S proteasome, activates protein breakdown by 20 S proteasomes. J. Biol. Chem., 1999. 274: p. 26008– 26014. 37 WOLF, S., et al., Characterization of ARC, a divergent member of the AAA ATPase family from Rhodococcus erythropolis. J. Mol. Biol., 1998. 277: p. 13–25. 38 WOLLENBERG, K. and J. C. SWAFFIELD, Evolution of proteasomal ATPases. Mol. Biol. Evol., 2001. 18 (6): p. 962–974. 39 BRAUN, B. C., et al., The base of the proteasome regulatory particle exhibits chaperone-like activity. Nat. Cell Biol., 1999. 1: p. 221–226. 40 STRICKLAND, E., et al., Recognition of misfolding proteins by PA700, the regulatory subcomplex of the 26S proteasome. J. Biol. Chem., 2000. 275: p. 5565–5572. 41 NAVON, A. and A. L. GOLDBERG, Proteins are unfolded on the surface of the ATPase ring before transport into the proteasome. Mol. Cell, 2001. 8 (6): p. 1339–1349. 42 LIU, C.-W., et al., Conformational remodeling of proteasomal substrates by PA700, the 19S regulatory complex of the 26S Proteasome. J. Biol. Chem., 2002. 277 (30): p. 26815–26820. 43 BENAROUDJ, N., et al., ATP hydrolysis by the proteasome regulatory complex PAN serves multiple functions in protein degradation. Mol. Cell, 2003. 11: p. 69–78. 44 OGURA, T. and K. TANAKA, Dissecting various ATP-dependent steps involved in proteasomal degradation. Mol. Cell, 2003. 11 (1): p. 3–5. 45 KENNISTON, J. A., et al., Studies with a prokaryotic protease show that the cost of translocation consumes much of the energy that is used in degradation. Cell, 2003. 114: p. 511–520. 46 MA, C. P., C. A. SLAUGHTER, and G. N. DEMARTINO, Identiﬁcation, puriﬁcation,

47

48

49

50

51

52

53

54

55

56

57

and characterization of a protein activator (PA28) of the 20S proteasome. J. Biol. Chem., 1992. 267: p. 10515–10523. REALINI, C., et al., Characterization of recombinant REGα, REGβ, and REGγ proteasome activators. J. Biol. Chem., 1997. 272: p. 25483–25492. HENDIL, K. B., S. KHAN, and K. TANAKA, Simultaneous binding of PA28 and PA700 activators to 20S proteasomes. Biochem. J., 1998. 332: p. 749–754. YAO, Y., et al., Structural and functional characterizations of the proteasome-activating protein PA26 from Trypanosoma brucei. J. Biol. Chem., 1999. 274 (48): p. 33921–33930. MCCUTCHEN-MALONEY, S. L., et al., cDNA cloning, expression, and functional characterization of PI31, a proline-rich inhibitor of the proteasome. J. Biol. Chem., 2000. 275: p. 18557– 18565. USTRELL, V., et al., PA200, a nuclear proteasome activator involved in DNA repair. EMBO J., 2002. 21 (13): p. 3516–3525. RECHSTEINER, M., C. REALINI, and V. USTRELL, The proteasome activator 11S REG (PA28) and class I antigen presentation. Biochem. J., 2000. 345: p. 1–15. STOHWASSER, R., et al., Kinetic evidences for facilitation of peptide channeling by the proteasomal activator PA28. Eur. J. Biochem., 2000. 267: p. 6221–6230. KOHLER, A., et al., The substrate translocation channel of the proteasome. Biochimie, 2001. 83(3–4): p. 325–332. FORSTER, A., F. G. WHITBY, and C. P. HILL, The pore of activated 20S proteasomes has an ordered 7-fold symmetric conformation. EMBO J., 2003. 22 (17): p. 4356–4364. KOEHLER, A., et al., The axial channel of the proteasome core particle is gated by the Rpt2 ATPase and controls both substrate entry and product release. Mol. Cell, 2001. 7: p. 1143–1152. GROLL, M., et al., Investigations on the Maturation and Regulation of Archaebacterial Proteasomes+ Journal of Molecular Biology, 2003. 327(1 SU): p. 75–83.

References 58 WILSON, H. L., et al., Biochemical and physical properties of the M. jannaschii 20S proteasome and PAN, a homolog of the ATPase (Rpt) subunits of the eucaryal 26S proteasome. J. Bacteriol., 2000. 186 (6): p. 1680–1692. 59 KWON, Y. D., et al., Crystal structures of the Rhodococcus proteasome with and without its pro-peptides: implications for the role of the pro-peptide in proteasome assembly. J. Mol. Biol., 2004. 335: p. 233–245. 60 OSMULSKI, P. A. and M. GACZYNSKA, Nanoenzymology of the 20S proteasome: Proteasomal actions are controlled by the allosteric transition. Biochem., 2002. 41: p. 7047–7053. 61 IMAI, J., et al., The molecular chaperone Hsp90 plays a role in the assembly and maintenance of the 26S proteasome. EMBO J., 2003. 22 (14): p. 3557–3567. 62 CHEN, P. and M. HOCHSTRASSER, Autocatalytic subunit processing couples active site formation in the 20S proteasome to completion of assembly. Cell, 1996. 86: p. 961–972. 63 DUBIEL, W., et al., Puriﬁcation of an 11S regulator of the multicatalytic protease. J. Biol. Chem., 1992. 267 (31): p. 22369–22377. 64 RUBIN, D. M., et al., Active site mutants in the six regulatory particle ATPases reveal multiple roles for ATP in the proteasome. EMBO J., 1998. 17 (17): p. 4909–4919.

65 COUX, O., An interaction map of proteasome subunits. Biochem. Soc. Trans., 2003. 31: p. 465–469. 66 DAVY, A., et al., A protein-protein map of the C. elegans 26S proteasome. EMBO Rep., 2001. 2 (9): p. 821–828. 67 FU, H. Y., et al., Subunit interaction maps for the regulatory particle of the 26s proteasome and the cop9 signalosome reveal a conserved core structure. EMBO J., 2001. 20 (24): p. 7096–7107. 68 FERRELL, K., et al., Regulatory subunit interactions of the 26S proteasome, a complex problem. Trends Biochem. Sci., 2000. 25 (2): p. 83–88. 69 PAPAPOSTOLOU, D., O. COUX, and M. REBOUD-RAVAUX, Regulation of the 26S proteasome activities by peptides mimicking cleavage products*1. Biochemical and Biophysical Research Communications, 2002. 295 (5): p. 1090–1095. 70 ZHANG, F., et al., O-GlcNAc modiﬁcation is an endogenous inhibitor of the proteasome. Cell, 2003. 115 (6): p. 715–725. 71 GROETTRUP, M., et al., A role for the proteasome regulator PA28a in antigen presentation. Nature, 1996. 381: p. 166–168. 72 CASCIO, P., et al., Properties of the hybrid form of the 26S proteasome containing both 19S and PA28 complexes. EMBO J., 2002. 21 (11): p. 2636–2645.

17

1

Tailoring Protein Scaffolds for Ligand Recognition A. Skerra

Originally published in: Protein-Ligand Interactions.Edited by Hans-Joachim B¨ohm, Gisbert Schneider. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30521-6

1 Introduction

A major goal of medicinal chemistry is the design of low-molecular-weight ligands that bind to target proteins in a tight and speciﬁc manner. In the case of enzymes, these ligands act as inhibitors or allosteric effectors, while in the case of transmembrane receptors, they serve as agonistic or antagonistic signaling molecules. Ligands of these types have conventionally been derived from natural compound libraries and, more recently, via combinatorial synthesis. The quickly growing number of proteins with known three-dimensional structure and the signiﬁcant methodological improvements in the structural elucidation of proteins during the past decade [1] – employing X-ray crystallography or nuclear magnetic resonance (NMR) techniques – have strongly promoted the computer-aided drug-design approach. Especially enzyme inhibitors can now be readily constructed on the basis of structural information about the target macromolecule [2]. Nevertheless, in the case of receptor targets, the rational prediction of cognate compounds is still hampered due to the inherent difﬁculties associated with their crystallization or NMR study. An inverse task is given when there is demand for a macromolecule that specifically binds a small ligand. This question has only recently been addressed by peptide chemistry. For example, antiparallel bundles of four α-helices, which were assembled on a cyclic peptide structure as template, have been used to create hydrophobic cavities for heme as a low-molecular-weight compound [3]. The speciﬁc complexation of FeIII ·protoporphyrin IX was facilitated by the proper positioning of liganding His residues. While this approach could be interesting from the perspective of rational protein design, it may be limited to special applications, and detailed structural information about the complex is not yet available. Deeper mechanistic insight into the molecular recognition of small molecules has been gained from antibodies, a class of natural proteins that have traditionally Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Tailoring Protein Scaffolds for Ligand Recognition

served as speciﬁc binding agents for a variety of “hapten” ligands [4]. Numerous practical applications exist for such antibodies in the ﬁelds of medical diagnostics as well as bioanalytics, where so-called immunochemical methods provide a quick, inexpensive, and reliable method for the sensitive detection of metabolites and even xenobiotic compounds [5]. There are also examples of hapten-binding antibodies with clinical use, e.g., in the therapeutic treatment of poisoning with cardiac steroids like digoxin. In this case an Fab fragment of an antibody with high afﬁnity towards the small molecule is administered, which removes the free compound from blood circulation, prevents it from binding to the cell surface receptor, and makes it amenable to renal ﬁltration or degradation in the liver [6]. With respect to the generation of cognate ligand-receptor proteins, one disadvantage is that low-molecular-weight compounds as such cannot be directly used for the immunization of animals. Rather, these haptens must be conjugated to macromolecular carriers in order to elicit an effective immune response. Nevertheless, attempts to generate antibodies with high afﬁnities and speciﬁcities against small ligands have often remained unsuccessful. Two potential problems need to be considered in this context. First, when antibodies are to be raised against metabolically occurring substances, they might interfere with physiological processes. Furthermore, if the compounds are toxic, immunization may not be possible at all. Second, and more generally, antibodies were probably evolved by the immune system mainly for the recognition of proteins or other macromolecular targets (like nucleic acids or oligosaccharides) rather than low-molecular-weight compounds. This notion is supported by the crystal structures of various antibody fragments in complex with either antigens or haptens. In the ﬁrst case an extended interface is formed between the antigen-binding site of the antibody (the paratope) and the macromolecular target. Typically, a surface of approximately 800 Å2 is buried and at least ﬁve of the six hypervariable loops (complementarity-determining regions, CDRs) – possibly even together with residues from the structurally conserved framework regions of the antibody variable domains – are involved [7, 8]. The shape of the combining site is often ﬂat but also can be slightly concave or convex. In contrast, in the case of haptens the mode of interaction with the paratope is much more restricted because a pocket for the ligand needs to be formed in order to provide a sufﬁcient number of interactions that ensure tight complex formation. This pocket is usually located at the interface between the pair of variable domains from the light and heavy chains of the antibody. Hence, a cleft must be formed whose shape is mainly determined by the two CDR-3 loops protruding from the VH and VL domains, which are related by a pseudo C2 -symmetry axis. Because a minimal hydrophobic contact area between VH and VL is required in order to maintain the non-covalent domain association, the size of this pocket is limited, and many ligands therefore become just partially buried when bound to an antibody (see Section 4). In addition to this structural consideration, there are empirical observations from recombinant antibody technology indicating that it is difﬁcult to generate an antibody fragment with exquisite speciﬁcity towards a hapten using combinatorial libraries cloned from unimmunized donor gene pools or derived from synthetic genes. In contrast, the in vitro selection of high-afﬁnity antibodies against proteins is

1 Introduction 3

Fig. 1 Three types of scaffolds, with a convex, ﬂat, and concave interface, respectively. (Left) Camel VH H domain, here presenting its extended CDR-3 loop towards the active site of an enzyme (PDB entry 1MEL). (Middle) Protein A with its interface made of two "-helices directed against an Ig constant domain (PDB entry 1BDD). (Right) Bilin-

binding protein with its pocket formed by four loops at the open end of the $-barrel structure for the complexation of biliverdin IX( , (PDB entry 1BBP). Loops or amino acid positions that are important for the molecular recognition of the respective target and that may be amenable to side chain exchanges are colored.

nowadays a routine procedure. Consequently, the recruitment of alternative protein classes for the generation of small ligand receptors has attracted attention [4]. To this end, the concept of using a scaffold – which means a protein architecture with high intrinsic stability – to create a binding site for the speciﬁc interaction with the target molecule has gained interest. An appropriate protein scaffold should provide a rigid folding unit that spatially brings together several exposed loops that form a continuous and extended interface such that multiple interaction with the target and hence tight binding are ensured. Ideally, such a scaffold should have structurally partitioned the generic information and stability of its polypeptide fold on the one hand and the local shape and molecular recognition function of its active site on the other (Fig. 1). Initially, this approach has had remarkable success in the generation of artiﬁcial binding proteins towards “protein antigens” (for a general review, see [9]). Several single domain proteins that belong to the generic immunoglobulin (Ig) fold, thus supporting a set of two or three hypervariable peptide loops on one end of a sandwich of β-sheets, have proven to be suitable for the recognition of such macromolecular targets. Prominent examples include an individual ﬁbronectin III domain [10] as well as certain VH domains derived from camel or llama Ig [11], which constitute soluble globular proteins even in the absence of a cognate VL domain. Typically, these scaffold proteins exhibit a wedge-shaped structure with the set of variable loops located at the tip in close mutual neighborhood (Fig. 1). Therefore, they seem to be particularly suited for complex formation at a groove on the surface of the target protein. In many cases this corresponds to the active site, and, indeed, effective enzyme inhibitors have been generated using the camel VH -domain approach. In contrast, the structural complexation of small molecules is difﬁcult to achieve with this scaffold. So far only cameloid antibodies recognizing rather large azo dye compounds as haptens have been described [12].

4 Tailoring Protein Scaffolds for Ligand Recognition

Another type of scaffold that has been successfully used for the recognition of prescribed target proteins originates from the bacterial immunoglobulin receptor protein A. So-called “afﬁbodies” were obtained by reshaping the natural Ig-binding interface of the Z domain of protein A, which is formed by a side-by-side pair of α-helices [13]. An essentially ﬂat surface is generated in this manner (Fig. 1), which can probably pack against a patch with low curvature on the target protein. Compared with the generation of recombinant receptor proteins against macromolecular targets, the recognition of small, hapten-like compounds obviously poses a greater challenge. To this end, a pocket with complementary shape must be created in order to enable the burial of a signiﬁcant area of hydrophobic surface and to provide a sufﬁcient number of protein-ligand interactions – van der Waals contacts, hydrogen bonds, and possibly salt bridges – such that practically useful dissociation constants in the nanomolar range result. In fact, these stringent demands still make it rather difﬁcult to apply rational design principles to the creation of cognate receptor proteins; instead, their construction has to rely on the powerful methodology of combinatorial biochemistry that is available today. Nevertheless, there exist only a few protein families in nature whose function lies in the plain complex formation with small molecules – as opposed to their biochemical conversion by enzymes or to the triggering of cellular signals via membrane or nuclear receptors. One example is given by the periplasmic nutrient-binding proteins that are found in Escherichia coli and other Gram-negative bacteria, comprising a variety of proteins with speciﬁcities for sugars, amino acids, and essential inorganic ions such as phosphate and sulfate [14]. These proteins serve for the transient complexation of their cognate compounds, followed by controlled delivery to transporter proteins that reside in the inner bacterial plasma membrane. Yet, despite their similarity in function, their sizes vary considerably and the mechanism of ligand complexation usually involves several distinct globular domains. Similarly, streptavidin from Streptomyces avidinii – and also its eukaryotic counterpart avidin, which occurs in chicken egg white – has evolved only in order to tightly complex biotin, a small vitamin compound [15]. In this case the complexation is kinetically almost irreversible, which makes sense for its role as a bacterial antibiotic protein and has led to its widespread use as a biochemical reagent (for references, see [16]). The lipocalins constitute another family of secretory ligand-binding proteins, which are typical for higher organisms. Initially, they were discovered in vertebrates, such as the retinol-binding protein (RBP) in man [17], but in fact lipocalins are found in a variety of eukaryotes and even in bacteria [18–20]. Generally, they serve for the transport or storage of poorly soluble or chemically sensitive compounds. Although their primary structures mostly lack detectable homology, structural analyses revealed a common fold for these proteins, comprising a rigid β-barrel as the central element of the lipocalin architecture [21]. The ligand is bound at the open end of this supersecondary structure, where a set of four loops forms the entrance to a structurally well-deﬁned pocket. Consequently, lipocalins have emerged as an attractive scaffold with potential for the engineering of artiﬁcial ligand-binding proteins.

2 Lipocalins: A Class of Natural Compound Carriers

2 Lipocalins: A Class of Natural Compound Carriers

The ﬁrst lipocalin whose 3-D structure was solved and reﬁned at high resolution was the human plasma retinol-binding protein (RBP) [22, 23]. RBP acts as a natural transporter of vitamin A (retinol) in the blood of vertebrates. Upon complexation in a hydrophobic cavity with complementary shape, the poorly soluble terpenoid alcohol becomes packaged by the protein and protected from oxidation or double-bond isomerization. RBP is synthesized in the liver and directly loaded with the ligand in the hepatocyte, where retinol is stored. Furthermore, the holo-RBP forms a structurally deﬁned ternary complex with transthyretin [24], also known as prealbumin. After delivery of the retinol ligand to a target tissue, the complex decomposes and the monomeric apo-RBP becomes ﬁltered out by the kidney and degraded. In the crystal structure, RBP folds into a single globular domain of approximately 40 Å in diameter (Fig. 2) whose central part is made of an eight-stranded, up- anddown β-barrel. At the amino-terminal end, the β-sheet region is ﬂanked by a coiled peptide segment, and at the carboxy-terminal end, it is followed by an α-helix and an amino acid stretch in a more or less extended conformation. Within the β-barrel the anti-parallel strands (assigned A to H) are arranged in a (+1)7 topology. They wind in a right-handed and conical manner around a central axis such that part of the ﬁrst strand A is hydrogen bonded via its backbone to the last strand H again. One end of the β-barrel is closed by the amino-terminal peptide segment that runs across its bottom between the two short loops connecting strands B/C and F/G, respectively, before it enters into β-strand A. Dense packing of side chains in this region and within the adjacent interior of the barrel structure leads to the formation of a hydrophobic core. The other end of the β-barrel is open to the solvent and forms a characteristic pocket. In the case of RBP, retinol is encapsulated as a ligand and protrudes into the barrel by almost half of its depth. The entrance to the pocket is formed by a set of four loops, which connect the eight antiparallel β-strands in a pairwise fashion. Because of the chalice-like shape of the protein (Fig. 2) and since many members of this family complex lipophilic compounds, the term “lipocalins” was proposed [25]. Several other lipocalins whose tertiary structures have been elucidated adopt a very similar fold. These were dubbed “prototypic” lipocalins [21] in order to distinguish them from more distantly related members of the family [18]. Within this subset, especially the β-barrel with the attached α-helix is highly conserved. In contrast, the four loops that form the entrance to the ligand pocket vary considerably in sequence, conformation, and length, thus effecting the differing ligand speciﬁcities (Fig. 2). However, not all lipocalins need to complex a small ligand in order to fulﬁll their physiological role. In aphrodisin, for example, which acts as a strong pheromone on male hamsters, the polypeptide itself seems to be responsible for the biological activity, thus requiring transfer of the non-volatile macromolecule by physical contact [26]. Even though a potential ligand pocket was recently detected in its crystal

5

6 Tailoring Protein Scaffolds for Ligand Recognition

Fig. 2 Generic structure of lipocalins. (Top) Ribbon representation of the retinol-binding protein with the bound vitamin A (PDB entry 1RBP). The four loops are shown in dark red at the open end of the $-barrel, and the three characteristic disulﬁde bonds of the

RBP are highlighted. (Bottom) Superposition of six natural lipocalins with diverse ligand speciﬁcities (PDB entries 1BBP, 1BEB, 1BJ7, 1EPA, 1MUP, 1RBP). The $-barrel is colored black (for detailed description, see [21]).

2 Lipocalins: A Class of Natural Compound Carriers

structure [27], attempts to identify a cognate low-molecular-weight molecule have failed. For some other lipocalins promiscuous binding of hydrophobic ligands was assumed [18, 28]. In the case of human apolipoprotein D (ApoD), for example, which occurs as a peripheral protein of the high-density lipoprotein (HDL) complex in serum, a whole series of potential ligands has been discussed [29]. Yet, thorough binding studies with the rigorously puriﬁed recombinant protein revealed just two ligands to be complexed with approximately micromolar dissociation constants: progesterone and arachidonic acid [30]. Because progesterone is well discriminated by ApoD against related steroids, such as pregnenolone and testosterone, it seems likely that arachidonic acid is recognized at a different binding site. Lipocalins are typical secretory proteins containing disulﬁde bonds. Human RBP possesses the maximal number of three disulﬁde cross-links that was observed so far. One of them joins the carboxy-terminal end of the polypeptide chain to the βbarrel (Cys70 –Cys174 ). Another one ﬁxes the amino-terminal segment of the protein to the carboxy-terminal end of the α-helix (Cys4 –Cys160 ). The third disulﬁde bond (Cys120 –Cys129 ) links the two neighboring strands G and H just underneath loop # 4 at the open end of the β-barrel (cf Fig. 2). The latter two disulﬁde bridges are characteristic for RBP. Although a Cys residue close to the amino-terminus is also found in several other lipocalins, it usually forms a disulﬁde bond with a Cys residue in strand G of the β-barrel, as in the bilin-binding protein (BBP), which carries two disulﬁde bonds [31, 32]. The C-terminal disulﬁde bond is obviously conserved in the lipocalin family, especially in those members that possess just one of them, e.g., the human neutrophil gelatinase-associated lipocalin (hNGAL) [33, 34]. Yet, there are certain local deviations as in the BBP, where this link is made to a Cys residue in strand B instead of strand D, as in RBP. Some lipocalins do not possess disulﬁde bonds at all, e.g., the bacterial lipocalin [20]. Hence, it seems that stabilization of the lipocalin architecture does not generally necessitate disulﬁde cross-links, contrasting with the immunoglobulin fold [35]. Many lipocalins are abundant in serum or tissue ﬂuids. However, their glycosylation status varies. Human RBP, for example, is not glycosylated, whereas ApoD from human plasma was shown to be glycosylated at both of its potential N-glycosylation sites, Asn45 and Asn78 [36] – possibly in contrast with other tissues where this lipocalin is also expressed. Nevertheless, when synthesized as a recombinant protein in E. coli, via secretion into the bacterial periplasm, the unglycosylated ApoD can be isolated as a soluble and functional protein [30]. Similarly, hNGAL, which is normally glycosylated at a single position, can be obtained as an unglycosylated protein from E. coli and adopts its proper tertiary structure, as elucidated by NMR analysis [33]. Finally, many lipocalins exist as soluble monomers. RBP, for example, which participates in a reversible complex formation with transthyretin, can be isolated as a fully stable monomer, either in complex with retinol or in the absence of the ligand [37]. Interestingly, several mammalian lipocalins appear to be linked via a disulﬁde bond to other functional protein complexes. For example, human ApoD carries

7

8 Tailoring Protein Scaffolds for Ligand Recognition

a ﬁfth Cys residue in addition to those giving rise to its two intra-chain disulﬁde bonds. Its unpaired thiol side chain is connected to a Cys residue of apolipoprotein A-II, which is an integral lipoprotein of the HDL particle. When this residue is removed by site-directed mutagenesis, the recombinant ApoD can be isolated in a soluble monomeric state [30]. Moreover, human ApoD is naturally produced as an individual protein in some other tissue ﬂuids [29], and in other organisms, such as rabbits, the unpaired Cys residue is even missing. Similarly, hNGAL normally occurs cross-linked with gelatinase B (known as matrix metalloproteinase 9), but also as a monomeric or homodimeric serum protein, and it was successfully produced in a monomeric state for structural studies [33]. Taken together, lipocalins provide attractive candidates in order to engineer novel ligand speciﬁcities. Features like their small size (typically between 150 and 180 residues), monomeric polypeptide composition, dispensable posttranslational modiﬁcation, and robust protein fold not only facilitate protein-engineering studies but also provide advantages for practical applications.

3 Anticalins: Lipocalins Reshaped via Combinatorial Biotechnology

In a ﬁrst attempt to tailor the ligand pocket of a lipocalin, the bilin-binding protein (BBP) served as a biochemically well-characterized model protein. The BBP originally occurs as a secretory protein in the butterﬂy Pieris brassicae, where it complexes biliverdin IXγ , a metabolic oxidation product of protoporphyrin IX. Hence, it serves for coloration as well as photoprotection, especially at the larval state. Natural BBP is found in two isoforms [32]. BBP-I forms a dimer in solution, whereas BBP-II, which likely arises from deamidation of the amino-terminal Asn residue of BBP-I, adopts a stable monomeric state. Genetic analysis revealed that only BBP-I is encoded on the insect chromosome [38]. After fusion of the mature part of the polypeptide chain to a bacterial leader peptide, the apo-BBP could be produced in the periplasm of E. coli as a recombinant protein in a functional state. For this purpose the amino-terminal Asn residue was directly exchanged by Asp at the genetic level and a monomeric BBP with full ligand-binding capability was obtained [38]. The crystal structure of the natural holo-BBP was elucidated at high resolution [31, 32]. It revealed the characteristic β-barrel fold with the tetrapyrrole ligand bound in a helical conformation (Fig. 3). Compared with human RBP, the four loops give room to a wider and shallower pocket for biliverdin. Consequently, BBP appeared to be a promising candidate for the reshaping of its ligand-binding site towards a variety of target compounds. To this end, the methodology of combinatorial biochemistry was applied (Fig. 3), comprising steps of (1) directed random mutagenesis of the loop regions in order to create a molecular library and (2) selection of cognate binding proteins from this library against a prescribed ligand. Based on the 3-D structure of BBP, a total of 16 amino acid positions were identiﬁed within the four loop segments – as well as adjoining regions of the β-strands – that dominate the interface with the natural ligand (Fig. 3). These

3 Anticalins: Lipocalins Reshaped via Combinatorial Biotechnology

Fig. 3 Generation of anticalins by randomization and selection. (Top) Randomized amino acid positions (corresponding side chains are shown in light blue) in the BBP, depicted here with the bound biliverdin in dark blue. (Bottom) Principle of the molecular library selection.

positions were chosen to fulﬁll two criteria: ﬁrst, they could be expected to tolerate both small and large side chain substitutions, and second, they appeared to reach as deeply as possible into the binding pocket. Hence, one could assess whether the hydrophobic core in the deeper part of the β-barrel would still be functional with respect to the stable folding of corresponding mutants. The 16 positions in the cloned BBP cDNA [38] were subjected to site-directed random mutagenesis using a two-step assembly polymerase chain reaction (PCR) with the help of primer oligodeoxynucleotides that were synthesized with mixed bases at the mutagenic codon positions. In order to introduce unique restriction

9

10 Tailoring Protein Scaffolds for Ligand Recognition

sites at both ends of the ampliﬁed central fragment of the BBP structural gene (both for BstXI, but with mutually non-compatible overhangs), two amino acids had to be exchanged at positions belonging to the β-barrel: Asn21 → Gln and Lys135 → Met. In addition, the recombinant BBP carried the mutation Asn1 → Asp mentioned above and Lys87 → Ser, which was introduced in order to remove a proteolytic cleavage site [39]. Thus, there were altogether four ﬁxed amino acid replacements in addition to the randomized side chains. The mutagenized gene cassette was then inserted into an appropriate E. coli vector and a genetic library comprising 3.7 × 108 variants was prepared [40]. The phagemid-display technique [41] was employed in order to select BBP variants with novel binding speciﬁcities from the resulting library [39]. For this purpose the BBP variants were produced as fusion proteins with a bacterial signal peptide at the amino-terminus and with the Strep-tag II, followed by a truncated pIII phage coat protein, at the carboxy-terminus [40]. In this case the amino acids 217 to 406 of the gene III product from ﬁlamentous phage M13 were used. The whole fusion gene was cloned on a phasmid vector under the tight transcriptional control of the chemically inducible tetracycline promoter [42] so that phagemid particles displaying BBP variants on their surface were efﬁciently produced under appropriate conditions. Fluorescein, a well known immunological hapten [43] with many applications in biochemistry and biophysics and a collection of commercially available derivatives, served as the prescribed ligand for BBP variants in the ﬁrst selection study. The phagemid random library was used for panning on a plastic surface coated with a covalent conjugate of ﬂuorescein with bovine serum albumin (BSA). After six cycles of adsorption, acid elution, and phagemid re-ampliﬁcation, the speciﬁc enrichment of a mutant phagemid fraction was observed. From DNA sequence analysis of 10 arbitrarily chosen clones, it became apparent that just four different BBP variants were still present in this population. Three of them – dubbed FluA, FluB, and FluC – gave rise to strong signals for the binding of several ﬂuorescein conjugates when produced as soluble proteins and investigated in an ELISA. In each of these variants, all 16 randomized amino acids had been exchanged when compared with the wild-type BBP [40]. Interestingly, four substitutions were identical among the three selected variants: Arg58 , Arg95 , Arg116 , and His127 . The corresponding preponderance of positively charged side chains was in agreement with the several negative charge centers of the ﬂuorescein derivative that was used in the selection. A similar effect had been observed in attempts to select recombinant antibody fragments against the same hapten from a semi-synthetic combinatorial library [44]. The BBP variant FluA was subjected to detailed biochemical characterization. The engineered lipocalin could be produced at high yield in the periplasm of E. coli (9.1 mg per 2 L culture compared with 1.2 mg for wild-type BBP) and isolated to homogeneity in one step via the Strep-tag method [45]. According to the relative shift in electrophoretic mobility between oxidized and reduced state of the protein, the two disulﬁde bonds of the BBP scaffold were correctly formed. Furthermore, the far UV circular dichroism spectrum of FluA revealed no marked differences

3 Anticalins: Lipocalins Reshaped via Combinatorial Biotechnology

from the wild-type protein. Consequently, the BBP had tolerated 16 amino acid exchanges within its ligand pocket – plus the four rationally introduced mutations mentioned above – without losing its folding properties as a lipocalin. The novel ligand-binding activity of FluA was studied in ELISA experiments with ﬂuorescein coupled to different carrier proteins. In each case steep saturation curves were observed with half-maximal concentrations in the nanomolar range, so that the recognition of the hapten appeared to be independent of the macromolecular context. Thermodynamic dissociation constants for the complex formation between FluA and ﬂuorescein and some related compounds were determined by ﬂuorescence titration in solution, measuring the emission of the protein’s Tyr and Trp residues. As a result, ﬂuorescein was bound slightly stronger than its two derivatives 4-aminoﬂuorescein and 4-glutarylamidoﬂuorescein, the compounds that actually had been used in the synthesis of the protein conjugates for the selection experiments. In contrast, pyrogallol red, a chemically similar triphenylmethane compound, was bound two orders of magnitude less tightly, while no binding at all could be detected for the related dyes phenolphthalein or rhodamine B. Interestingly, when the titration was performed such that the hapten’s own characteristic ﬂuorescence was measured, almost complete quenching was observed. From this very accurate titration experiment, a K D value of 35.2 ± 3.2 nM was determined for the FluA·ﬂuorescein complex [40]. Using the same spectroscopic effect, the association kinetics between ﬂuorescein and FluA could be measured by rapid mixing, yielding a K on value of 5.28 ±0.05 × 106 M−1 s−1 (G. Beste and A. Skerra, unpublished). The phenomenon of almost complete ﬂuorescence quenching upon complex formation between ﬂuorescein and the engineered lipocalin FluA was elucidated in a series of time-resolved light-absorption measurements after pulse activation of the bound ligand [46]. These experiments revealed an ultrafast electron transfer between the ﬂuorescein and an aromatic side chain (Tyr or Trp) in its close proximity. The excited ﬂuorescein dianion within the ligand pocket abstracts an electron from the neighboring amino acid at a rate of 400 fs. The resulting radical trianion is deactivated in a radiationless process – with a larger time constant of 4 ps – to the spectroscopic ground state of ﬂuorescein by back transfer of the electron. The observed monoexponentiality in the formation of the excited state, and also of its subsequent decay, points toward a high structural deﬁnition of the hapten-binding site and explains the highly efﬁcient quenching effect that becomes apparent under stationary conditions. Clearly, this spectroscopic phenomenon was a serendipitous event because there was no corresponding selection applied during the generation of FluA. Also, the other mutants that were selected along with it did not show ﬂuorescence quenching to the same high extent. Nevertheless, it is remarkable that such an efﬁcient electron transfer process, which is even faster (by a factor 3–4) than the one measured between bacteriochlorophyll and bacteriopheophytin in the bacterial reaction center of Rhodobacter sphaeroides, can be achieved by combinatorial protein design (see the discussion in [46]). The ﬂuorescence-quenching effect observed with FluA is also signiﬁcantly more pronounced than for antibodies that were raised against

11

12 Tailoring Protein Scaffolds for Ligand Recognition

ﬂuorescein by immunization. Hence, the engineered lipocalin may be of interest as a reagent in biophysical studies where the speciﬁc masking of ﬂuorescein groups is desired. The same random library of BBP variants was used in selection experiments with several other haptens. The cardiac steroid digoxigenin served as a molecular target of practical relevance [47]. In this case the library was screened by combining phagemid display with a ﬁlter-sandwich colony-screening assay in order to rapidly identify individual BBP variants with corresponding ligand-binding activity. As a result, one variant with speciﬁcity for digoxigenin was isolated whose dissociation constant was determined to be 295 ± 37 nM by means of protein ﬂuorescence titration [47]. In an attempt to further improve the ligand afﬁnity of this engineered lipocalin, dubbed DigA, an in vitro afﬁnity maturation was performed. Inspection of the primary sequence of the BBP variant revealed that the ﬁrst of the four loops mainly had charged side chains acquired during the selection. However, these amino acids did not appear to be optimal for the complexation of the hydrophilic, though uncharged, steroid. In this respect it should be noted that the BBP random library with its complexity of 3.7 × 108 was by far too small to represent all possible combinations of the 16 randomized amino acids. Therefore, in principle, considerable room remained for further sequence optimization in parts of the binding pocket, once molecular recognition of a speciﬁc ligand was achieved. Consequently, six amino acid positions within loop # 1 of DigA were selectively subjected to oligodeoxynucleotide-directed random mutagenesis, again followed by selection for the binding of digoxigenin groups via phagemid display and colony screening. In this way, the variant DigA16 was obtained, which binds digoxigenin signiﬁcantly tighter, with a K D value of 30.2 ± 3.6 nM [47]. Remarkably, the glycosylated natural compound digoxin, which has three sugar molecules attached to C-3 of the steroid system [48], is bound with precisely the same afﬁnity. Likewise, digoxigenin conjugates with several different carrier proteins – bovine serum albumin, ovalbumin, or ribonuclease A – that were covalently linked via an aliphatic spacer to the same steroid ring position gave rise to indistinguishable binding signals in solid-phase assays. Thus, the BBP variant DigA16 recognized the digoxigenin group as a true hapten, without detectable context dependence. Subsequent ligand-binding studies [47] revealed that a chemically similar cardiac glycoside, digitoxin, which differs from digoxin just by a single missing hydroxyl group, is bound stronger still, with a K D value equal to 3.2 ±0.54 nM. In contrast, complex formation with the related steroid ouabain, which often shows cross-reactivity with antibodies raised against digoxin [48], was not detectable. In addition, no complex formation was observed with the steroid testosterone or with 4-aminoﬂuorescein, the ligand that had served before in the selection of the BBP variant FluA. Hence, DigA16 represents an engineered lipocalin with high afﬁnity and pronounced speciﬁcity towards a rather hydrophilic steroid. The dramatic alteration in the ligand-binding function of this lipocalin was achieved by exchanging a total of 17 amino acids [47], which form most of the pocket in the natural BBP, together

4 Structural Aspects of Ligand Recognition by Engineered Lipocalins

with the four site-directed amino acid replacements that were introduced into the scaffold in order to make it better amenable to protein-engineering experiments (see above). Attempts were made to raise the afﬁnity of DigA16 for the digoxigenin group even further by applying additional cycles of targeted random mutagenesis at loops# 3 and # 4 [49]. The resulting variant DigA16/19, which carries several new mutations in loop# 4, exhibits improved afﬁnity for digoxigenin, with K D =12.4±1.3 nM. In addition, DigA16/19 possesses enhanced ligand speciﬁcity and also recognizes part of the linker that was used for ﬁxing the steroid group to the carrier protein. During these experiments the randomized residues were still restricted to the original set of positions chosen within the four loops. However, from recent structural analyses (see Section 4) it appeared that there are additional, so far nonmutated amino acids that contribute to the shape of the ligand pocket and may therefore govern afﬁnity and speciﬁcity for the steroids. Thus, future improvement in molecular recognition by this engineered lipocalin may be guided by rational principles. Nevertheless, the successful construction of a digoxigenin-binding lipocalin provides a novel and useful tool in biochemistry. Digoxigenin and digitoxigenin are medically important compounds, either as potentially poisonous substances or as drugs with therapeutic value – as long as they are applied at a precisely adjusted dose [50]. Furthermore, the digoxigenin group has acquired recent popularity in biochemistry as a non-radioactive label for a variety of biomolecules. Several chemically activated derivatives are available for the selective labeling of proteins or nucleic acids so that digoxigenin can be used independently from the commonly employed biotin group, with the advantage of very low background-staining activity [51]. The generation of BBP variants with novel binding speciﬁcities for ﬂuorescein or digoxigenin, respectively, has demonstrated for the ﬁrst time that a lipocalin can be tailored to recognize non-natural ligands. In order to illustrate the antibody-like binding function of the engineered lipocalins, this new class of proteins was termed “anticalins” [40].

4 Structural Aspects of Ligand Recognition by Engineered Lipocalins

The 3-D structures of the ﬂuorescein- and digoxigenin-binding BBP variants have recently been analyzed by X-ray crystallography and compared with the original bilin-binding protein. The crystal structures were determined in different space groups and, for one variant, in both the presence and absence of the hapten, thus giving insight into the structural mechanisms of speciﬁc ligand recognition by the engineered lipocalins. In the case of the ﬂuorescein-binding variant FluA, crystals were obtained in the presence of the ligand at pH 8.1 with two FluA·ﬂuorescein complexes in the asymmetric unit, which were reﬁned to a resolution of 2.0 Å [52]. The two molecules

13

14 Tailoring Protein Scaffolds for Ligand Recognition

Fig. 4 Crystal structure of the anticalin FluA with the bound ﬂuorescein (green). C" positions of the 16 randomized amino acids are shown as gray spheres. Trp129 is depicted with its side chain in magenta.

were highly similar in structure, with a root mean square difference (rmsd) of 0.33 Å for 173 mutually superimposed Ca positions. The overall topology of the βbarrel with the α-helix attached to it, both of which are characteristic features of the lipocalin architecture (see Section 2), was found to be conserved (Fig. 4). Both disulﬁde bonds of the BBP scaffold, one between Cys18 and Cys115 and one between Cys42 and Cys47 , were also clearly visible. Upon superposition with the BBP crystal structure (molecule A from the Protein Data Base entry 1BBP [32]), an rmsd of 1.2 Å was calculated for 159 superimposed Cα positions. The largest structural differences were seen at the four loops that form the entrance to the binding site. The most prominent conformational changes occurred in loops# 1 and # 2. Loop# 1 had adopted a more extended conformation and moved away from the center of the ligand-binding site towards the bulk phase of the solvent. The Cα position of the mutated residue Asn36 (Val in BBP) at its tip was concomitantly displaced by approximately 8 Å. Loop# 3, which was also involved in the contacts with the structural neighbor in the asymmetric unit, had moved away from the barrel axis, thus opening the cavity for the bound ligand, with the Cα position o the non-mutated residue Gly92 at its tip shifted by about 6.6 Å. Fluorescein is bound at the bottom of the cleft that harbors biliverdin IXγ , in the wild-type BBP structure (Fig. 2). Its xanthenolone moiety is located close to the center of the β-barrel, while the carboxyphenyl group is oriented towards the entrance of the pocket. The para ring position (with respect to the central carbon

4 Structural Aspects of Ligand Recognition by Engineered Lipocalins

atom of the triphenylmethane dye), which carried the linker group during the selection experiments for this anticalin [40], is accessible from the solvent via a narrow channel. An area of 454 Å2 , corresponding to 91% of the solvent-accessible surface of ﬂuorescein, became buried in the complex. Approximately 50% of the buried area from the side of the protein is contributed by 6 of the 16 residues that were mutated in the generation of FluA from BBP. The remaining buried surface belongs to 10 residues that were not mutated. Unexpectedly, when compared with the complexation of biliverdin by BBP, ﬂuorescein was found to be inserted even more deeply into the hydrophobic core of the β-barrel. In the central region of the protein, where no mutations had been introduced, the necessary space was created by the movement of loop# 3 and by rearrangement of several side chains. In particular, there is a ladder of residues comprising His86 , Phe99 , His127 , and Trp129 , which have undergone a concerted reorientation of their aromatic side chains. Of those, only His127 , which participates in a packing interaction with one of the phenolic rings of ﬂuorescein, was mutated in the generation of FluA from BBP. The non-mutated residue Trp129 is located directly underneath. It has shown major side chain reorientation and gives rise to an extended π stacking interaction at the middle of the xanthenolone ring system of ﬂuorescein via coplanar arrangement in van der Waals distance. It seems that the introduction of the imidazole side chain at position 127 has triggered the whole movement, including those of the non-mutated residues His86 and Phe99 at the bottom of the pocket. As a result of these changes, original residues of BBP form a signiﬁcant part of the reshaped ligand pocket. Hence, it appears that in addition to the loop region, the hydrophobic core of the lipocalin displays considerable plasticity as well. Residues close to the hydrophobic core that had been thought to be crucial for proper folding of the protein have adopted completely new side chain orientations in order to allow for the binding of the new ligand. Future design of anticalins with even higher afﬁnities for small haptens may therefore also include residues from the central part of the β-barrel. When considering the side chains that make up the binding site for ﬂuorescein, both mutated and non-mutated, it is found that they are predominantly polar in nature. Thus, the lipocalin pocket is by no means restricted to lipophilic ligands, as was anticipated before and as the name of this protein family may suggest. Furthermore, the crystal structure conﬁrms the crucial role of the mutated basic residues Arg58 and His127 in FluA for the tight binding of ﬂuorescein, which was previously demonstrated by site-directed mutagenesis experiments [40]. However, the mechanism of interaction is different from the earlier assumption that was based on the crystal structure of the anti-ﬂuorescein antibody 4–4–20 [53]. There, the xanthenolone group is oriented such that an Arg and a His residue (with Cα distance similar to BBP) are each in contact with one of the phenolic moieties and form hydrogen bonds. In contrast, Arg58 and His127 of FluA contact the same phenolic group, but from opposite sides. This arrangement is made possible by the extensive structural reorganization at the open end of the β-barrel (see above). Three Arg residues at positions 88, 95, and 116, which were shown to have a

15

16 Tailoring Protein Scaffolds for Ligand Recognition

favorable effect on the ligand afﬁnity via their positively charged side chains, do not form direct contacts with the bound ﬂuorescein and should therefore mainly exert an electrostatic inﬂuence. Yet, the reasons for their peculiar arrangement at the entrance to the ligand pocket and their strong conservation among the several selected ﬂuorescein-binding BBP variants [40] are not obvious at present. Taken together, the crystallographic analysis of the anticalin FluA, which was generated by combinatorial design from a prototypic lipocalin, reveals that mutated residues within the loop region and adjoining parts of the β-barrel can give rise to three different effects. First, they can contribute direct contacts with the bound ligand or at least provide an appropriate electrostatic environment. Second, they may induce novel backbone conformations in the loops and thus lead to the formation of a pocket with generic shape complementarity with the prescribed ligand. Finally, there are certain amino acids that inﬂuence the side chain conformations of neighboring residues and thus reshape the pocket in an indirect manner. Similar phenomena are known from antibodies where, apart from amino acids that contact the antigen or hapten, key residues within the hypervariable loops are responsible for their canonical backbone conformations [54] and framework residues indirectly ﬁne-tune the shape of the combining site [55]. The crystal structure of the FluA·ﬂuorescein complex also provides a structural explanation for the strong quenching effect of this particular anticalin that was mentioned before. Tight coplanar packing of the indole ring of Trp129 against the xanthenolone system of the bound ﬂuorescein was observed (Fig. 4). Consequently, this aromatic residue is the likely candidate for the highly efﬁcient electron-transfer process and is optimally positioned in this respect. In the case of the anticalin DigA16, crystal structures were solved not only for the bound digoxigenin but also for the complex with the related steroid digitoxigenin and for the uncomplexed apo-protein [56]. The crystals, which were grown at pH 7.6–8.0 and whose structures were reﬁned to resolutions between 1.8 and 1.9 Å, were essentially isomorphous and contained one monomer per asymmetric unit. In addition, crystals were obtained in another space group for the uncomplexed original BBP variant DigA, although with poorer diffraction quality. Again, the overall topology of the lipocalin, comprising the β-barrel with the α-helix attached to it, remained conserved, whereas the set of four loops at the entrance to the ligand pocket revealed clear structural differences in comparison with the BBP. The most prominent conformational change was observed for loop# 1, where an α-helical segment of seven amino acids appeared in DigA16 (residues 33 through 39), in both the presence and absence of the steroid ligand (Fig. 5). Notably, the non-mutated residue Tyr39 within the new helix – which faces the solvent in the BBP – has shifted in all three DigA16 structures such that it packs with its side chain against the bound steroid, if present. This dramatic conformational change results in a displacement of its Cα atom by 9 Å with respect to the position in the BBP. The α-helical loop conformation seems to be essentially stabilized by two speciﬁc interactions. First, the side chains of the mutated residues Arg58 and Ser60 in loop# 2 form hydrogen bonds with the carbonyl oxygen of Tyr39 at the carboxy-terminal end

4 Structural Aspects of Ligand Recognition by Engineered Lipocalins

Fig. 5 Molecular recognition of haptens by engineered lipocalins versus antibodies. (Top) Crystal structure of the anticalin DigA16 with the bound digoxigenin (yellow). The arrow points to the hydroxyl substituent of steroid ring A, which had served for covalent attachment – via a ﬂexible spacer – to a solid support during the selection process for this BBP variant. The C" posi-

tions of the 16 initially randomized amino acids are shown as gray spheres. (Bottom) Crystal structure of the anti-digoxigenin Fab fragment 26–10 (PDB entry 1IGJ) with the VL and VH domains colored cyan and magenta, respectively. The bound di-goxigenin group is shown in yellow, while the digitoxose sugar attached to it (at the same ring position as above) is colored light gray.

17

18 Tailoring Protein Scaffolds for Ligand Recognition

of the helix. Second, the newly introduced side chain of His35 at the amino-terminal end of the helix becomes packed between this Tyr residue and the side chain of Leu127 in loop# 4, which also resulted from the mutagenesis. Thus, a small pocket for the imidazole group is formed, which is closed at the bottom by the side chains of Trp129 (already present in the BBP) and Gln28 . The latter residue was introduced during the afﬁnity maturation from DigA to DigA16, leading to a 10-fold improved ligand afﬁnity (see Section 3). In the complex with DigA16, digoxigenin is bound at the bottom of the cleft that otherwise harbors biliverdin IXγ , in the wild-type BBP, and it roughly replaces the space previously occupied by one of the tetrapyrrole rings [32]. An area of 514 Å, corresponding to 95% of the solvent-accessible surface of digoxigenin have thus become buried. Approximately half of the buried surface is contributed by 9 of the 17 residues that were mutated in the generation of DigA16, while the remainder is due to 10 residues that have not been mutated. Similarly, as deduced for the anticalin FluA, speciﬁc recognition of the steroid ligand is achieved by preformed shape complementarity of the ligand pocket in apo-DigA16 as a result of (1) side chain replacements and (2) indirect effects of mutated positions on wild-type residues. The latter effect can especially be seen for the shifted residue Tyr39 , which is part of the newly formed helix in loop# 1, and for altered side chain conformations of Phe99 and Trp129 at the bottom of the ligand pocket. These two conserved residues have rotated their aromatic side chains by approximately 120◦ compared with BBP, thus enabling accommodation of the new bulky steroid ligand. Their conformation appears to be similar in all three DigA16 structures, even in the absence of the ligand. Apparently, the side chain rotation of Trp129 is in concert with the extensive rearrangement in the upper part of the binding cleft, particularly within loop# 1 due to the mutated residues. The steroid ligand is bound mainly via van der Waals interactions but also via hydrogen bonds with its polar substituents. Several water molecules have become buried in the ligand pocket as well and participate in hydrogen bond interactions with the hydroxyl and lactone groups of the hydrophilic steroid. Of the buried protein surface in the digoxigenin complex, 34% is provided by non-hydrocarbon groups (for comparison, 33% in the BBP). Hence, the binding site has considerable polar character, which is in contrast with the almost entirely hydrophobic nature of the ligand pocket in other lipocalins, such as the retinol-binding protein (with a corresponding value of 16%). Similarly, the steroid ligand is almost fully trapped within the binding site, with a remaining accessible surface of 5%. Indeed, there is a small gap in the protein shell that permits accessibility of the steroid position C-3, which carried the linker group during the selection procedure [47]. Thus, as a result of the combinatorial protein design experiment, the corresponding steroid derivative must have been almost perfectly shielded from solvent. From a comparison between the DigA16 complexes with digoxigenin versus digitoxigenin, the structural mechanism of the ﬁne speciﬁcity in the steroid recognition – and discrimination between these closely related ligands – became apparent. The non-mutated residue His86 at the bottom of the ligand pocket, which is close to the hydrophobic core within the β-barrel, plays a crucial role in this

4 Structural Aspects of Ligand Recognition by Engineered Lipocalins

respect and is involved in an induced ﬁt during ligand complexation. Upon binding of digoxigenin, the side chain of His86 , which points into the empty cavity in apo-DigA16, is displaced by the ligand and forms a hydrogen bond with the hydroxyl group HO-12 of the steroid. Thus, it becomes rotated towards Tyr22 , which is part of a loop at the closed end of the β-barrel. The side chain of Tyr22 itself rotates away from its original hydrogen-bonding partner Thr104 in apo-DigA16 into the direction of His86 and forms a hydrogen bond with the imidazole side chain instead. Digitoxigenin, which is bound essentially at the same position and with the same orientation as digoxigenin, lacks the OH group at the steroid position C-12. Consequently, a hydrogen bond between His86 and digitoxigenin is missing in the complex with DigA16, and its imidazole side chain packs closer to the steroid ring system. Thus, compared with the digoxigenin complex it is partially rotated back into the position that it has assumed in the apo-protein. Accordingly, the side chain of Tyr22 still forms a hydrogen bond with Thr104 . Instead, a water molecule appears at the position that is occupied by the phenolic hydroxyl group of Tyr22 in the DigA16·digoxigenin complex, which is weakly hydrogen-bonded to His86 in the case of the bound digitoxigenin. The set of available crystal structures also provides an explanation for the effect of the afﬁnity maturation that led from the original DigA anticalin to the DigA16 mutant. During this step, several amino acids were randomized in loop# 1 [47]. While most of the corresponding side chains are solvent exposed in the tertiary structure, two residues are probably relevant for the loop conformation and improved ligand afﬁnity of DigA16. First, His35 remained conserved with respect to the DigA sequence, which underlines its role in the helix conformation as described above, together with the invariant residue Tyr39 . Second, Glu28 was consistently replaced by Gln (in several mutants that were selected along with DigA16; see [47]). In the apo-DigA structure, the side chain of Glu28 adopts a different conformation compared with Gln28 in apo-DigA16 and forms a salt bridge with the Arg58 guanidinium group. As a consequence, the carboxylate moiety of Glu28 may sterically interfere with the position of the lactone substituent at ring D of the bound steroid ligand. In addition, due to a corresponding shift of the Arg side chain, the helix – which covers the bound ligand – probably undergoes minor repositioning. Hence, it seems that the introduction of Gln28 is mainly responsible for the 10-fold enhanced afﬁnity of DigA16. Notably, the mechanism of molecular recognition by the anticalin is different from the manner in which digoxigenin is bound by immunoglobulins. Two crystal structures have been described for Fab fragments of monoclonal antibodies that were raised against digoxin as a hapten: the Fab “26–10” in complex with digoxin (PDB accession code 1IGJ; [57]) and the Fab “40–50” in complex with ouabain (PDB accession code 1IBG; [58]). In both cases only between 60% and 70% of the steroid became buried from solvent during complex formation (Fig. 5). The ring system is covered by protein residues mostly from one side, probably due to the limited capacity of the Ig architecture to create a deep pocket at the interface of the VH and VL domains. Even though high afﬁnities have been achieved by these antibodies, their speciﬁcities are poor because both signiﬁcantly cross-react with ouabain, a

19

20 Tailoring Protein Scaffolds for Ligand Recognition

Fig. 6 Plasticity of the engineered lipocalin-binding site: superposition of the natural BBP with its variants FluA and DigA16 (with loops colored blue, green, and yellow, respectively).

cardiac steroid related to digoxin [48, 58]. In contrast, the engineered lipocalin DigA16 clearly distinguishes between these two compounds [47], thus providing a functional advantage. In summary, a phenomenon of pronounced structural plasticity was observed in the engineered BBP variants FluA and DigA or DigA16, which means that the backbone conformation of the lipocalin loop region was strongly inﬂuenced by the side chain replacements (Fig. 6). However, in the context of a given amino acid sequence, the conformational ﬂexibility of these loops seems to be rather low because no signiﬁcant differences were observed in the two independently reﬁned FluA·ﬂuorescein complexes or when comparing the DigA16 structures in the absence or presence of the ligand. This effect was not entirely expected a priori, because from a superposition of natural lipocalins with known tertiary structures [21], it was not clear to which extent the loop conformation is governed by its own distinct sequence versus individually variable features of the β-barrel structure that provides the support.

5 Prospects and Future Applications of Anticalins

The functional and structural data that have been gathered during the engineering of lipocalins for the recognition of two unrelated low-molecular-weight ligands

5 Prospects and Future Applications of Anticalins 21

clearly demonstrate the potential of this scaffold for the generation of artiﬁcial receptor proteins with high afﬁnity and speciﬁcity for prescribed target molecules. Our ﬁndings conﬁrm that the β-barrel architecture of lipocalins constitutes a remarkably stable scaffold. Even though amino acids were replaced in at least 20 different positions, most of them within the binding site of the BBP, the overall topology and the β-barrel structure itself were retained. Structural changes were merely observed at a local level and essentially restricted to the loop regions. It seems that lipocalins indeed provide a partitioned protein architecture, wherein the β-barrel – together with the ﬁxed loops at its closed end and the α-helix attached to it – provides a rigid framework that is structurally conserved among the “prototypic” members of this family [21], while the set of four loops at its open end can be hypervariable. This situation is reminiscent of antibodies, where a set of six CDRs presented on top of a largely constant framework region is responsible for the speciﬁc binding of the antigen or hapten. However, compared with recombinant antibody fragments, engineered lipocalins should provide signiﬁcant beneﬁts because they are composed of one instead of two polypeptide chains, they have a much smaller size, and their set of four loops can be easily manipulated simultaneously at the genetic level. Consequently, lipocalins provide a promising alternative to antibody fragments for the engineering of artiﬁcial ligand-binding proteins, therefore called “anticalins,” using the methods of combinatorial biochemistry [21, 40]. The most striking property of the lipocalin scaffold is its ability to provide a well-deﬁned and conformationally rigid cavity for the ligand. The ligand pocket is signiﬁcantly deeper than the hapten-binding sites found in antibodies and may even reach down into the hydrophobic core of the lipocalin, as was seen in the case of the ﬂuorescein-binding anticalin FluA. A similar mode of complexation would not be possible for an antibody fragment because of the detrimental effect on the non-covalent association between VH and VL . Hence, the lipocalin-bound ligand can be trapped from the solvent and becomes almost fully surrounded by protein residues, which explains the pronounced speciﬁcity, especially observed for the steroid ligands. Thus, an extended linker structure, which should also include hydrophilic groups, seems to be required for the functional immobilization of the target compound during the selection procedure for the cognate anticalin. It would be nice to devise selection techniques that no longer necessitate the covalent ﬁxation of the target because under such conditions anticalins might be generated that fully encapsulate their ligands. Remarkably, the anticalins obtained up to now recognize their low-molecularweight ligands independently from the carrier – usually a stable globular protein like BSA or RNAse – that was employed for target display. This was shown for both the ﬂuorescein group [40] and digoxigenin [47]. In this respect, anticalins distinguish themselves from many antibodies, especially when derived from synthetic libraries [4], and also from different protein scaffolds that have been tested for similar purposes. For example, when mutants of cytochrome b562 with two randomized loop regions were selected against an organic target compound, the hapten was only recognized – with a weak micromolar afﬁnity – as long as it remained linked to the original BSA carrier [59].

22 Tailoring Protein Scaffolds for Ligand Recognition

The well-deﬁned binding properties for the small ligand in combination with the lack of cross-reactivity with the macromolecular conjugate partner are probably due to the choice of the randomized positions in the lipocalin within an inner zone at the open end of the β-barrel but below the exposed tips of the loop region (Fig. 3). Consequently, just a few of the randomized residues are potentially accessible from outside the pocket, most likely only after signiﬁcant changes in the backbone conformation – as observed for loop # 1, which adopts an α-helical structure in DigA16 (see Section 4). However, the scope of molecular recognition by engineered lipocalins should not be restricted to low-molecular-weight compounds. Indeed, the more exposed side chains of the four loops may be speciﬁcally randomized for the generation of another sort of anticalin libraries that could be useful for selection towards macromolecular targets. Preliminary data from our laboratory suggest that mutants of the BBP can be obtained in this manner, which exhibit speciﬁc binding activity for prescribed proteins with dissociation constants in the nanomolar range. The proof of concept for the generation of anticalins as a novel class of receptor proteins with deﬁned ligand-binding properties was realized using the BBP as a model lipocalin. It has been shown that, because of their simple and robust architecture, these anticalins provide several practical advantages. For example, they are remarkably stable against denaturation. Thermal unfolding studies revealed a melting temperature of 61.3 ◦ C for the recombinant BBP and an even higher T m value of 72.8 ◦ C for the anticalin DigA, although this variant had not been selected for enhanced folding stability [49]. Another advantage of the lipocalin architecture relates to the fact that both ends of the polypeptide chain are sterically accessible at the outside of the β-barrel and should normally not interfere with the structure of the ligand-binding site. Thus, anticalins are amenable to the construction of functional fusion proteins at both their amino- and carboxy-termini. This was demonstrated in the case of DigA16 for alkaline phosphatase, which could serve directly as a reporter enzyme for the detection of digoxigenin groups after fusion with either end of the anticalin [47]. Anticalins may even be fused to each other, leading to so-called “duocalins,” a novel class of bifunctional ligand-binding proteins [60]. The insights that were gained so far from the anticalin approach illustrate once again the huge potential of polypeptides to adopt diverse molecular shapes, seen here for the ligand pockets of engineered lipocalins. Given the high plasticity of the loop region, which probably constitutes a speciﬁc functional advantage of this protein family, the rational prediction of the inﬂuence of amino acid substitutions on the structure of the binding site will probably remain difﬁcult in the near future. But even when applying the powerful methods of combinatorial biochemistry, one should be aware of the still limited options for the realization of novel functional active sites, which is caused by the vast number of possible sequence combinations on the one hand and the restricted number of molecules that can be physically generated and applied to a selection experiment on the other. We have tried to address this generic problem in protein engineering by making a careful choice of amino acids for random mutagenesis – in fact, just less then half of the positions that one could actually consider – in order to reduce the combinatorial complexity and thus create a potent molecular library in the functional sense. The fact that speciﬁc hapten-binding activities were immediately derived from

References

this library conﬁrms the validity of this concept. Nevertheless, additional steps of afﬁnity maturation may be needed, as demonstrated for DigA16 [47, 49], in order to ﬁne-tune the shape complementarity of the binding site after initial ligand recognition property for the ligand was imprinted. The rational choice of positions to be modiﬁed, in combination with repeated cycles of targeted randomization and selection – corresponding to a kind of molecular evolution – is probably the best general strategy for obtaining novel proteins with well-deﬁned ligand-binding function, at least for the moment. In conclusion, speciﬁcally engineered anticalins open numerous areas of application as ligand-binding proteins not only in bioanalytics and separation technology but also in medical diagnostics and possibly even therapy. Especially for the latter purpose, it could be advantageous to generate anticalins based on a human lipocalin framework [21]. Hence, immunogenic side effects will be minimized upon repeated administration to patients. The preparation of appropriate fusion proteins should permit the introduction of useful effector functions – as already demonstrated with enzymes or certain binding modules, such as the albumin-binding domain [47]. The ﬁeld of engineered protein scaffolds for molecular recognition has rapidly emerged during the past few years (for reviews, see [9, 61]). Among the several scaffold structures that are currently being exploited, immunoglobulins and lipocalins certainly stand out. Both families are utilized by nature itself in order to provide speciﬁc binding proteins based on a stable tertiary fold that supports hypervariable loops. In the case of immunoglobulins, hundreds of millions of different antibodies are constantly being created in each individual’s immune system using mechanisms of genetic recombination and somatic hypermutation. In contrast, lipocalins are much smaller in number and have been stably evolved in many organisms in order to serve more specialized physiological functions. Whereas antibodies must predominantly recognize macromolecular antigens – such as proteins and carbohydrates – in their defense against microbial or viral pathogens, lipocalins seem to mainly serve for the transport and storage of low-molecular-weight compounds. Especially in this respect, lipocalins are distinct from other protein scaffolds that have been subject to protein engineering and successfully used for the generation of binding modules against protein targets. The promising results obtained here from the tailoring of a natural lipocalin for the recognition of hapten-like ligands emphasizes the unique potential of this protein family for the generation of corresponding receptor proteins. Apart from the interesting practical applications, this ﬁeld of research will also offer conceptual insight into the mechanisms of molecular recognition between proteins and small molecules in general.

References 1 R. C. STEVENS, S. YOKOYAMA, I. A. WILSON, Science 2001, 294, 89–92. 2 H. GOHLKE, G. KLEBE, Angew. Chem. Int. Ed. 2002, 41, 2644–2676.

3 H. K. RAU, N. DEJONGE, W. HAEHNEL, Angew. Chem. Int. Ed. Engl. 2000 39, 250–253.

23

24 Tailoring Protein Scaffolds for Ligand Recognition

4 G. A. WEISS, H. B. LOWMAN, Chem. Biol. 2000, 7, R177–R184. 5 B. HARRIS, Trends Biotechnol. 1999, 17, 290–296. 6 M. R. UJHELYI, S. ROBERT, Clin. Pharmacokinet. 1995, 28, 483–493. 7 D. R. DAVIES, S. CHACKO, Acc. Chem. Res. 1993, 26, 421–427. 8 E. A. PADLAN, Mol. Immunol. 1994, 31, 169–217. 9 A. SKERRA, J. Mol. Recognit. 2000, 13, 167–187. 10 A. KOIDE, C. W. BAILEY, X. HUANG, S. KOIDE, J. Mol. Biol. 1998, 284, 1141–1151. 11 S. MUYLDERMANS, M. LAUWEREYS, J. Mol. Recognit. 1999, 12, 131–140. 12 S. SPINELLI, M. TEGONI, L. FRENKEN, C. van VLIET, C. CAMBILLAU, J. Mol. Biol. 2001, 311, 123–129. 13 K. NORD, E. GUNNERIUSSON, J. RINGDAHL, S. STAHL, M. UHLE´ N, P.-Å. NYGREN, Nature Biotechnol. 1997, 15, 772–777. 14 F. A. QUIOCHO, P. S. LEDVINA, Mol. Microbiol. 1996, 20, 17–25. 15 Y. LINDQUIST, G. SCHNEIDER, Curr. Opin. Struct. Biol. 1996, 6, 798–803. 16 M. WILCHEK, E. A. BAYER, Biomol. Eng. 1999, 16, 1–4. 17 M. E. NEWCOMER, D. E. ONG, Biochim. Biophys. Acta 2000, 1482, 57–64. 18 D. R. FLOWER, Biochem. J. 1996, 318, 1–14. 19 G. GUTIE´ RREZ, M.D. GANFORNINA, D. S´ANCHEZ, Biochim. Biophys. Acta 2000, 1482, 35–45. 20 R. E. BISHOP, Biochim. Biophys. Acta 2000, 1482, 73–83. 21 A. SKERRA, Biochim. Biophys. Acta 2000, 1482, 337–350. 22 M. E. NEWCOMER, T. A. JONES, J. ÅQVIST, J. SUNDELIN, U. ERIKSSON, L. RASK, P. A. PETERSON, EMBO J. 1984, 3, 1451–1454. 23 S. W. COWAN, M. E. NEWCOMER, T. A. JONES, Proteins: Struct. Funct. Genet. 1990, 8, 44–61. 24 H. L. MONACO, M. RIZZI, A. CODA, Science 1995, 268, 1039–1041. 25 S. PERVAIZ, K. BREW, Fed. Am. Soc. Exp. Biol. 1987, 1, 209–214. 26 W. J. HENZEL, H. RODRIGUEZ, A. G. SINGER, J. T. STULTS, F. MACRIDES, W. C. AGOSTA, H. NIALL, J. Biol. Chem. 1988, 263, 16682–16687.

27 F. VINCENT, D. L¨OBEL, K. BROWN, S. SPINELLI, P. GROTE, H. BREER, C. CAMBILLAU, M. TEGONI, J. Mol. Biol. 2001, 305, 459–469. 28 D. R. FLOWER, J. Mol. Recognit. 1995, 8, 185–195. 29 E. RASSART, A. BEDIRIAN, S. DO CARMO, O. GUINARD, J. SIROIS, L. TERRISSE, R. MILNE, Biochim. Biophys. Acta 2000, 1482, 185–198. 30 M. VOGT, A. SKERRA, J. Mol. Recognit. 2001, 14, 79–86. 31 R. HUBER, M. SCHNEIDER, O. EPP, I. MAYR, A. MESSERSCHMIDT, J. PFLUGRATH, H. KAYSER, J. Mol. Biol. 1987, 195, 423–434. 32 R. HUBER, M. SCHNEIDER, I. MAYR, R. M¨ULLER, R. DEUTZMANN, F. SUTER, H. ZUBER, H. FALK, H. KAYSER, J. Mol. Biol. 1987, 198, 499–513. 33 M. COLES, T. DIERCKS, B. MUEHLENWEG, S. BARTSCH, V. Z¨OLZER, H. TSCHESCHE, H. KESSLER, J. Mol. Biol. 1999, 289, 139–157. 34 D. H. GOETZ, S. T. WILLIE, R. S. ARMEN, T. BRATT, N. BORREGAARD, R. K. STRONG, Biochemistry 2000, 39, 1935–1941. 35 R. GLOCKSHUBER, T. SCHMIDT, A. PLU¨ CKTHUN, Biochemistry 1992, 31, 1270–1279. 36 P. A. SCHINDLER, C. A. SETTINERI, X. COLLET, C. J. FIELDING, A. L. BURLINGAME, Protein Sci. 1995, 4, 791–803. 37 H.N. M¨ULLER, A. SKERRA, J. Mol. Biol. 1993, 230, 725–732. 38 F. S. SCHMIDT, A. SKERRA, Eur. J. Biochem. 1994, 219, 855–863. 39 A. SKERRA, Rev. Mol. Biotechnol. 2001, 74, 257–275. 40 G. BESTE, F. S. SCHMIDT, T. STIBORA, A. SKERRA, Proc. Natl. Acad. Sci. USA 1999, 96, 1898–1903. 41 G. P. SMITH, V. A. PETRENKO, Chem. Rev. 1997, 97, 391–410. 42 A. SKERRA, Gene 1994, 151, 131–135. 43 E. W. VOSS, Jr., Fluorescein Hapten: An Immunological Probe, CRC Press, Boca Raton, FL, 1984. 44 C. F. BARBAS III, J.D. BAIN, D. M. HOEKSTRA, R. A. LERNER, Proc. Natl. Acad. Sci. USA 1992, 89, 4457–4461. 45 A. SKERRA, T. G. M. SCHMIDT, Methods Enzymol. 2000, 326, 271–304.

References 46 M. G¨OTZ, S. HESS, G. BESTE, A. SKERRA, M. E. MICHEL-BEYERLE, Biochemistry 2002, 41, 4156–4164. 47 S. SCHLEHUBER, G. BESTE, A. SKERRA, J. Mol. Biol. 2000, 297, 1105–1120. 48 M. MUDGETT HUNTER M. N. MARGOLIES, A. JU, E. HABER, J. Immunol. 1982, 192, 1165–1172. 49 S. SCHLEHUBER, A. SKERRA, Biophys. Chem. 2002, 96, 213–228. 50 A. R. HASTREITER, E. G. JOHN, R. L. van der HORST, Clin. Perinatol. 1988, 15, 491–522. 51 T. MCCREERY, Mol. Biotechnol. 1997, 7, 121–124. 52 I. KORNDO¨ RFER, G. BESTE, A. SKERRA, in preparation. 53 M. WHITLOW, A. J. HOWARD, J. F. WOOD, E. W. VOSS, Jr., K. D. HARDMAN, Protein Eng. 1995, 8, 749–761.

54 C. CHOTHIA, A. M. LESK, J. Mol. Biol. 1987, 196, 901–917. 55 J. FOOTE, G. WINTER, J. Mol. Biol. 1992, 224, 487–499. 56 I. KORNDO¨ RFER, S. SCHLEHUBER, A. SKERRA, in preparation. 57 P. D. JEFFREY, R. K. STRONG, L. C. SIEKER, CY. Y. CHANG, R. L. CAMPBELL, G. A. PETSKO, E. HABER, M. N. MARGOLIES, S. SHERIFF, Proc. Natl. Acad. Sci. USA 1993, 90, 10310–10314. 58 P. D. JEFFREY, J. F. SCHILDBACH, CY. Y. CHANG, P. H. KUSSIE, M. N. MARGOLIES, S. SHERIFF, J. Mol. Biol. 1995, 248, 344–360. 59 J. KU, P. G. SCHULTZ, Proc. Natl. Acad. Sci. USA 1995, 92, 6552–6556. 60 S. SCHLEHUBER, A. SKERRA, Biol. Chem. 2001, 382, 1335–1342. 61 P.-Å. NYGREN, M. UHLE´ N, Curr. Opin. Struct. Biol. 1997, 7, 463–469.

25

1

Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities Uwe T. Bornscheuer

Ernst-Moritz-Arndt-Universit¨at, Greifswald, Germany

Originally published in: Directed Molecular Evolution of Proteins. Edited by Susanne Brakmann, Kai Johnsson. Copyright ľ 2002 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30423-1

1 Introduction

The application of enzymes is now well documented. They are used in organic chemistry – mainly for the production of optically pure compounds – synthesis of detergents and emulsiﬁers, lipid modiﬁcation, laundry formulations, food and beverage production etc. [1–6]. Due to their high chemo-, regio- and stereoselectivity, activity at ambient temperatures and pH, biocatalysts are often superior to chemical ones and this might explain that a considerable number of industrial bioprocesses have been established in recent years [2, 7]. Traditional methods to identify new enzymes are based on screening of e.g. soil samples or strain collections by enrichment cultures. Once a suitable biocatalyst is identiﬁed, strain improvement as well as cloning and expression of the encoding gene(s) enable production on a large scale. Unfortunately, not all microorganisms can be cultured using common fermentation technology and the number of accessible microorganisms from a sample is estimated to 0.001–1% depending on their origin [8]. In addition, not all enzymes found in nature are suitable for a speciﬁc synthetic problem and the useful properties mentioned above are not always satisfactory in practice. Until recently, these limitations were overcome by i) screening for alternative biocatalysts, ii) changes of the reaction system (e.g., from hydrolysis to esteriﬁcation) or iii) by rational protein design followed by site-directed mutagenesis, if the gene encoding the enzyme and its three-dimensional structure are available [9]. All these methods are very time-consuming and in case of rational protein design also information-intensive. A very promising alternative for the generation of enzymes with improved or altered properties is the strategy of directed evolution (also called evolutive biotechnology or molecular evolution) which came to the fore in the early 90s. In most Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities

Fig. 1 Principle of directed evolution.

cases, directed evolution is comprised of a random mutagenesis of the gene encoding the catalyst by error-prone polymerase chain reaction (epPCR) resulting in large libraries of mutants. Another method is based on DNA- (or gene-) shufﬂing and related techniques. Libraries thus created are then assayed by using high-throughput technologies to identify improved variants (Fig. 1). Further details on the principles and applications of directed evolution can be found elsewhere in this book and in a considerable number of recent reviews dealing with various aspects of directed evolution [8–21]. This chapter focuses on the creation of novel substrate speciﬁcities of enzymes.

2 General Considerations

During natural evolution, a broad variety of enzymes has been developed, which are classiﬁed according to the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (IUBMB). Thus, for each type of

2 General Considerations Table 1 Examples for the evolutionary generation of enzymes with novel substrate

speciﬁcities Wild-type activity

Method

Assay

Main Results

Ref.

Pseudomonas ﬂuorescent esterase E. coli IGPS

MS

growth/pHindicator

[22, 24, 26]

SDM

growth assay

E. coli ProFARI

shufﬂing

growth assay

E. coli β-galactosidase

DNA-shufﬂing

p-/o-nitro-phenyl glycosides

Bacillus megaterium P450 MO Pseudomonas SY77 glutaryl acylase

epPCR/SM

visual inspection

active double mutant acting on sterically hindered substrate evolved PRAI activity from IGPS scaffold evolved PRAI activity from ProFARI 300–1000-fold increased speciﬁcity for fucosides generation of indigo/indirubin

epPCR/SM

growth assay

increased Vmax /Km for adipyl-7-ADCA

[34]

Pseudomonas putida P450 MO

epPCR/StEP

digital image screening

[35, 37]

Staphylococcus aureus lipase

epPCR/shufﬂing

chromogenic assay

enhanced naphthalene hydroxylation/formation of dihydroxy compounds 12-fold increase in phospholipase activity

Bacillus thermocatenulatus lipase

epPCR/SDM

visual inspection

17-fold increase in phospholipase activity

[39]

Saccharomyces cerevisiae peroxidase E. coli aldolase

shufﬂing

visual inspection

300-fold increased activity for guaiacol

[40]

epPCR/shufﬂing

UV assay

acceptance of phosphate free substrates

[41]

[28]

[29]

[31]

[33]

[38]

Abbreviations: ADCA, adipyl-cephalosporic acid; epPCR, error-prone PCR; IGPS, indole-3-glycerol phosphate synthase; MO, monooxygenase; MS, mutator strain; PRAI, phosphoribosylanthranilate isomerase; ProFARI, N -[5-phosphoribosyl)formimino]-5-aminoimidazole-4-carboxylamide ribonucleotide isomerase; SDM, site-directed mutagenesis, StEP, staggered extension process

characterized enzyme an EC (Enzyme Commission) number has been provided (see: www.expasy.ch/enzyme/). For instance, all hydrolases have EC number 3 and further subdivisions are provided by three additional digits, e.g. all lipases (ofﬁcial name: triacylglycerol lipases) have the EC number 3.1.1.3 and are thus distinguished from esterases (ofﬁcial name: carboxyl esterases) having the EC number

3

4 Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities

3.1.1.1. This classiﬁcation is based on the substrate (and cofactor) speciﬁcity of an enzyme only, however often very similar amino acid sequences and also related three-dimensional structures can be observed. This suggests that the creation of novel enzyme speciﬁcity has a high probability if a closely related catalytic machinery and/or architecture is present for both types of activities, i.e. both substrates should be converted by the Ser-His-Asp triad present in lipases and amidases if an amidase substrate should be accepted by a lipase or vice versa. The chance for creating activity is usually enhanced if both enzymes share high sequence and/or structural homology. In other words, most readers will agree that it will be rather impossible to convert an esterase into a monooxygenase as these two enzymes differ substantially in their architecture, substrate and cofactor-requirement and consequently mode of action. With this in mind, it is not surprising that the examples reported in the literature (Table 1) for novel substrate speciﬁcities by directed evolution can be subdivided into the following three groups (Scheme 1): 1. Conversion of substrates, which in principle should be acceptable by the enzyme under investigation, but do not react due to e.g. steric hindrance, electronic effects etc.

Scheme 1 Selected examples illustrating possible targets for novel substrate speciﬁcities of enzymes. Esterases do not act on esters of tertiary alcohols and this problem thus belongs to group 1. Galactosidases only accept galactosides as substrates, but not glucosides and therefore this example

resembles group 2. Some lipases – e.g from Rhizopus sp., Bacillus sp. or Staphylococcus sp. – exhibit low activity towards the sn1-group of phospholipids. However, this activity is signiﬁcantly lower compared to phospholipase A1 (PLA1 ) and consequently it can be regarded as a group 3 target.

3 Examples

2. Conversion of substrates which are usually converted by another (sometimes closely related) enzyme. 3. The compounds are already converted by the enzyme of interest, but at very low activity compared to well accepted (natural) substrates.

It should be noted that a clear-cut distinction between these groups is not always feasible.

3 Examples 3.1 Group 1

The ﬁrst example in which a substrate not accepted by the wild-type enzyme could be converted by a mutant obtained by directed evolution was published by my group [22]. A sterically hindered 3-hydroxy ester (Scheme 2, 1a, b) resembling a building block in the envisaged total synthesis of epothilones was not accepted by 20 wild-type hydrolases (18 lipases, 2 esterases). Although closely related 3-hydroxy esters could be stereoselectively converted by a range of commercial lipases, this speciﬁc compound bearing two methyl groups at C4 did not react in hydrolysis of the corresponding ethyl ester as well as in acylation attempts at the 3-hydroxy group using vinyl acetate in toluene [23]. As the basic structure of the 3-hydroxy ester was set by the epothilone scaffold, the most promising alternative to obtain the compound in optically pure form was the creation of a hydrolase mutant exhibiting activity towards this substrate, preferentially in a stereoselective manner. Thus, a mutant library of an esterase from Pseudomonas ﬂuorescens (PFE) was created using the mutator strain Epicurian coli XL1-Red (available from Stratagene). Key to the identiﬁcation of improved variants from these libraries was an agar plate assay system based on pH indicators, thus leading to a change in color upon hydrolysis of the ethyl ester 1a (Scheme 3). Parallel assaying of replica-plated colonies on agar plates supplemented with the glycerol derivative 1b was used to reﬁne the identiﬁcation, because only E. coli colonies producing active esterases had access to the carbon source glycerol, thus leading to enhanced growth and in turn larger colonies [22, 24]. By this strategy, a double mutant (Leu181Val, Ala209Asp) was obtained, which indeed hydrolyzed the sterically hindered substrate. This was conﬁrmed by preparative scale reactions followed by gas chromatographic analysis to calculate conversion and enantiomeric excess as well as by determination of the optical rotation by polarimetry. The best mutant gave an enantiomeric excess of 25% ee which corresponds to an enantioselectivity of E ∼5.0, which is in accordance with the, in general, rather low selectivity of PFE. Several other mutants were identiﬁed, but enantiomeric excess was usually around 3–4 % ee. Interestingly, the activity towards acetic acid ethyl ester – the standard substrate to determine esterase activity by a pH-stat assay – was decreased considerably for various mutants. Whereas

5

6 Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities

Scheme 2 Structures of compounds used (1a,b, 4) and obtained (2, 3) by directed evolution of biocatalysts aiming at novel substrate speciﬁcities.

wild-type PFE showed an activity of 6 Umg−1 , the activity of the Leu181Val/ Ala209Asp mutant was 2.3 Umg−1 and some variants showed only activities <1.0Umg−1 . The 3D-structure of PFE is unknown, however it was possible to perform homology modeling based on the solved 3D-structure of a haloperoxidase from Streptomyces aureofaciers [25]. This revealed that both mutations are not close to the binding pocket but solely at the periphery of the enzyme [24]. Although this method was useful to turn an inactive wild-type enzyme into an active esterase, the best mutant showed only modest enantioselectivity. Further work showed that the enantioselectivity of PFE could be increased from E = 3.5 to E = 5.2–6.6 in a single round of mutation [26]. Libraries were created by epPCR as well as by using the mutator strain from Epicurian coli. An extremely accurate determination of the enantioselectivity was achieved by using resoruﬁn esters of (R)- or (S)-3-phenylbutyric acid, which allowed measurement of ﬂuorescence in microtiter plates (MTP) avoiding problems with interfering compounds present in the culture medium. A further increase in enantioselectivity up to E = 12 was achieved by saturation mutagenesis at all three positions identiﬁed for the ﬁrst

Scheme 3 Principle of the assay system used to identify active variants of esterase from Pseudomonasfuorescens (PFE) acting on the sterically hindered 3-hydroxy ester 1. Both substrates 1a,b yield the free acid leading to a color change; 1b also releases the carbon source glycerol leading to enhanced growth of esterase mutant producing E. coli clones.

3 Examples

Fig. 2 Mutants obtained by directed evolution and saturation mutagenesis showing enhanced enantioselectivity in the resolution of 3-phenylbutyric acid derivatives. MTP: E-values (Eapp ) determined in microtiter plates using the corresponding (R)- and (S)-resoruﬁn ester; GC:E-values (Etrue ) cal-

culated using the equation E = [ln(1-c)*(1ees )]/[ln(1-c)*(1+ ees )] from data determined by gas chromatographic analysis on a chiral column from samples obtained after esterase-catalyzed hydrolysis of (R,S)-3phenyl butyric acid ethyl ester [24].

generation mutants (E. Henke and U.T. Bornscheuer, unpublished). However, combinations of best variants by site-directed mutagenesis gave no further improvement (Fig. 2). It should be noted, that Eapp values determined in MTPs were quite close to Etrue values, which have been determined by gas chromatographic analysis of samples from resolutions of racemic mixtures of the corresponding ethylesters of 3-phenylbutyric acids. Here, also competition between the two enantiomers is included, whereas the MTP-assay delivers only apparent E values. Signiﬁcantly higher increases in enantioselectivity were achieved for the resolution of methyl 3-bromo-2-methylpropionate. Using the homology model of PFE, Trp29 and Phe199 were identiﬁed as positions for random mutagenesis. Screening of libraries using Quick E [27] identiﬁed a mutant Trp29Leu exhibiting an E = 90 compared to E = 12 for wild-type PFE. This is the highest selectivity ever achieved up to now by directed evolution techniques (U.T. Bornscheuer and R.J. Kazlauskas, unpublished), and allows the synthesis of optically pure substrate and product of this compound.

7

8 Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities

Scheme 4 Substrate speciﬁcities of ProFARI (involved in histidine biosynthesis) and PRAI (involved in tryptophan biosynthesis).

The knowledge about mutations affecting enantioselectivity and altered substrate speciﬁcity derived from these directed evolution studies of PFE is currently used as a basis to design variants combining both properties. In an excellent contribution, Fersht and co-workers evolved phosphoribosylanthranilate isomerase (PRAI) activity from the (βα)8 -barrel scaffold of indole-3glycerol phosphate synthase (IGPS) [28]. Their success provides a striking example for testing the “conserved scaffold” hypothesis of enzyme evolution. Careful design of the basic PRAI loop system into IGPS created the scaffold for directed evolution by DNA-shufﬂing and StEP. Active variants were identiﬁed by an agar plate assay using an E. coli strain that does not grow in the absence of tryptophan, which in turn is the product of PRAI activity. The mutant obtained had 28% identity to PRAI and 90% identity to IGPS. The same hypothesis that several different enzymes having the (βα)8 -barrel scaffold evolved from a common ancestor was used as a basis for the alteration of the substrate speciﬁcity of another isomerase having only 10% sequence homology [29]. This showed that a single round of gene-shufﬂing turned ProFARI (N -[5-phosphoribosyl)formimino]-5-aminoimidazole-4-carboxylamide ribonucleotide isomerase) – involved in histidine biosynthesis – into a mutant with PRAI activity catalyzing the same reaction in tryptophan biosynthesis (Scheme 4). In vivo selection of mutants in an E. coli strain lacking PRAI activity yielded two variants each with only a single amino acid substitution. One mutant catalyzed both reactions, although with only low residual wild-type activity (<4%). Both examples have been already discussed extensively in a recent review [30]. 3.2 Group 2

DNA-shufﬂing was successfully used to turn an E. coli lacZ β-galactosidase into a fucosidase [31]. Screening of ∼10 000 colonies per round based on chromogenic fucose substrates resulted in the identiﬁcation of evolved fucosidases with 10–20-fold increased kcat /K m for the fucose substrate compared to the wild-type. Compared to the native β-galactosidase, the ﬁnal evolved enzyme showed a 1000-fold increased substrate speciﬁcity for o-nitrophenyl fucopyranoside versus o-nitrophenyl

3 Examples

galactopyranoside and a 300-fold increase for the corresponding p-nitrophenol derivatives. In a related manner, attempts were made to convert the E. coli lacZ β-galactosidase into a β-glucosidase, however so far only minor changes in the substrate speciﬁcity have been reported [32]. In another example, some mutants of the P450-monooxygenase from Bacillus megaterium (P450 BM-3, expressed in E. coli) obtained by directed evolution using epPCR were found to hydroxylate indole at two different positions (Scheme 2, 2a, b) which upon air oxidation and dimerization yield indigo and indirubin [33]. So far, both compounds were known to be formed by the action of naphthalene dioxygenases or styrene monooxygenases, but are not produced by wild-type P450 BM3 and are thus a result of altered substrate speciﬁcity. Sequencing of several individual clones revealed that three positions (Phe87, Leu 188, Ala74) contained mutations. Next, simultaneous saturation mutagenesis at all three positions was performed to identify best “replacements” which led to the identiﬁcation of a triple mutant (Phe87Val, Leu188Gln, Ala74Gly) enabling formation of indigo and indirubin in milligram amounts. In a very recent paper the activity of a glutaryl acylase from Pseudomonas SY77 towards adipyl-7-ADCA – an important precursor in the synthesis of semi-synthetic cephalosporins – was altered by directed evolution [34]. Random mutagenesis of the α-subunit, growth selection of active variants followed by saturation mutagenesis identiﬁed three mutants showing decreased Vmax /Km values for glutaryl substrates and higher values towards the adipyl analog. The authors used a growth selection based on adipyl-leucine or adipyl serine to identify acylase mutants acting on adipic acid instead of glutaric acid side-chains in cephalosporins. 3.3 Group 3

A P450 monooxygenase from Pseudomonas putida primarily catalyzes the hydroxylation of camphor (P450cam ) utilizing the cofactor NADPH. In contrast, hydroxylation of naphthalene (Scheme 2, 3) occurs only to a minor extent. Coexpression of P450cam with horseradish peroxidase in E. coli in the presence of hydrogen peroxide allowed hydroxylation in the absence of NADPH via a “peroxide shunt” pathway. Moreover, under these reaction conditions, hydroxylated naphthalenes form ﬂuorescent dimers amenable to digital image screening [35]. Thus, approximately 200 000 mutants obtained by epPCR and the staggered extension process (StEP) [36] were easily screened for increased ﬂuorescence suggesting enhanced activity of P450cam . In addition, mutations could be identiﬁed having altered regiospeciﬁcities. Thus variants of P450cam were generated showing ∼20-fold higher activity compared to the wild-type as well as mutants catalyzing the formation of dihydroxy compounds [37]. Phospholipase activity was introduced into a Staphylococcus aureus lipase mutant previously created by site-directed mutagenesis, which displayed very low phospholipase activity. Next, directed evolution using epPCR and gene shufﬂing led to a substantial increase in the phospholipase/lipase activity ratio [38]. Mutants

9

10 Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities

were ﬁrst screened in a qualitative agar plate assay followed by high-throughput chromogenic assay based on a thio-derivative of a phospholipid. Upon phospholipase activity, the free SH-group was reacted with 5,5 -dithiobis(2-nitrobenzoic acid) yielding 5-thio-2-nitrobenzoic acid amenable to spectrophotometric quantiﬁcation. The best variant contained six mutations and displayed a 11.6-fold increase in phospholipase activity and a 11.5-fold increased phospholipase/lipase ratio compared to the starting mutant. Recently, similar ﬁndings were reported for a lipase from Bacillus thermocatenulatus expressed in E. coli [39]. A single round of random mutagenesis followed by screening of 6000 transformants on egg-yolk identiﬁed three variants with ∼10–12-fold phospholipase activity. Further rounds of random mutagenesis gave no improvement, but site-directed mutagenesis (Leu353Ser) gave a 17-fold enhanced phospholipase activity. The substrate speciﬁcity of a cytochrom c peroxidase (CCP) from Saccharomyces cerevisiae towards guaiacol (Scheme 2, 4) was increased 300-fold by means of DNAshufﬂing [40]. Screening was performed by spreading colonies onto nitrocellulose membranes. Mutants showing enhanced activity towards guaiacol were identiﬁed by visual inspection of fast-staining colonies due to a brownish color based on the formation of tetraguaiacol. Interestingly, sequencing revealed that the – in the super-family of peroxidase – fully conversed residue Arg48 was changed to His. The authors believe that this mutation plays a key role in the increase in activity toward the phenolic substrate guaiacol. Wild-type aldolase from E. coli poorly accepts non-phosphorylated D-2-keto-3deoxygluconate (KDG). Several rounds of epPCR and shufﬂing identiﬁed mutants with signiﬁcantly enhanced activity towards KDG [41]. The best mutant showed a 70fold increased catalytic efﬁciency with concomitant decrease in the activity towards the phosphorylated analog. Assaying was performed in microtiter plates based on reduction of released pyruvate by lactate dehydrogenase yielding a decrease in NADH concentration. Furthermore, library screening identiﬁed mutants accepting L-glyceraldehyde as substrate, which is converted by the wild-type aldolase to only a negligible extent.

4 Conclusions

The examples provided in this Chapter demonstrate that directed evolution resembles a very useful tool to create enzyme activities hardly accessible by means of rational protein design (Table 1). Even if the desired substrate speciﬁcity is known from other biocatalysts – e.g. phospholipase A1 activity – the advantage of the directed evolution approach resides in the already achieved functional expression of a particular protein. Thus bottlenecks arising from the identiﬁcation of enzymes by traditional screening and cultivation methods can be circumvented. In addition, directed evolution can dramatically reduce the time required for the provision of a suitable tailor-made enzyme, also because cloning and functional expression of the biocatalyst has already been achieved.

References

The striking examples by the Fersht and Sterner groups for the evolution of βα 8 -barrel enzymes also demonstrate that even functional and structural barriers between different enzymes can be overcome by means of directed evolution. Acknowledgment

I am especially grateful to Prof. Romas Kazlauskas (McGill University, Canada) and Dipl.-Chem. Erik Henke (Greifswald University) for their support. References 1 BORNSCHEUER, U. T., KAZLAUSKAS, R. J. (1999), Hydrolases in organic synthesis -regio- and stereoselective biotransformations, Weinheim: Wiley-VCH. 2 BORNSCHEUER, U. T. (2000), Enzymes in Lipid Modiﬁcation, Weinheim: Wiley-VCH. 3 DRAUZ, K., WALDMANN, H. (1995), Enzyme catalysis in organic synthesis, Vol. 1&2 Weinheim: VCH. 4 FABER, K. (2000), Biotransformations in Organic Chemistry, Berlin: Springer. 5 PATEL, R. N. (2000), Stereoselective Biocatalysis, New York: Marcel Dekker. 6 WONG, C.-H., WHITESIDES, G. M. (1994), Enzymes in synthetic organic chemistry, Oxford: Pergamon Press. 7 LIESE, A., SEELBACH, K., WANDREY, C. (2000), Industrial Biotransformations, Weinheim: Wiley-VCH. 8 MILLER, C. A. (2000), Advances in enzyme discovery, Inform 11, 489–495. 9 BORNSCHEUER, U. T., POHL, M. (2001), Improved biocatalysts by directed evolution and rational protein design, Curr. Opin. Chem. Biol. 5, 137–143. 10 ARNOLD, F. H. (1998), Design by directed evolution, Acc. Chem. Res. 31, 125–131. 11 ARNOLD, F. H., VOLKOV, A. A. (1999), Directed evolution of biocatalysts, Curr. Opin. Chem. Biol. 3, 54–59. 12 ARNOLD, F. H. (1999), Unnatural selection: molecular sex for fun and proﬁt, Engineering Science 40–50. 13 BORNSCHEUER, U. T. (1998), Directed evolution of enzymes, Angew. Chem. Int. Ed. 37, 3105–3108. 14 ARNOLD, F. H., MOORE, J. C. (1997), Optimizing industrial enzymes by

15

16

17

18

19

20

21

22

23

directed evolution, Adv. Biochem. Eng./Biotechnol. 58, 1–14. SCHMIDT-DANNERT, C., ARNOLD, F. H. (1999), Directed evolution of industrial enzymes, Trends Biotechnol. 17, 135–136. REETZ, M. T., JAEGER, K.-E. (1999), Superior biocatalysts by directed evolution, Topic Curr. Chem. 200, 31–57. PETROUNIA, LP., ARNOLD, F. H. (2000), Designed evolution of enzymatic properties, Curr. Opin. Biotechnol. 11, 325–330. TOBIN, M. B., GUSTAFSSON, C., HUISMAN, G. W. (2000), Directed evolution: the “rational” basis for “irrational”; design, Curr. Opin. Struct. Biol. 10, 421–427. SUTHERLAND, J. D. (2000), Evolutionary optimisation of enzymes, Curr. Opin. Chem. Biol. 4, 263–269. JAEGER, K. E., REETZ, M. T. (2000), Directed evolution of enantioselective enzymes for organic chemistry, Curr. Opin. Chem. Biol. 4, 68–73. REETZ, M. T. (2001), Combinatorial and evolution-based methods in the creation of enantioselective catalysts, Angew. Chem. Int. Ed. 40, 284–310. BORNSCHEUER, U. T., ALTENBUCHNER, J., MEYER, H. H. (1998), Directed evolution of an esterase for the stereoselective resolution of a key intermediate in the synthesis of Epothilones, Biotechnol. Bioeng. 58, 554–559. BORNSCHEUER, U., HERAR, A., KREYE L. WENDEL, V., CAPEWELL, A., MEYER, H. H., SCHEPER T. KOLISIS, F. N. (1993), Factors affecting the lipase catalyzed transesteriﬁcation reactions of 3-hydroxy

11

12 Evolutionary Generation of Enzymes with Novel Substrate Speciﬁcities

24

25

26

27

28

29

30

31

32

esters in organic solvents, Tetrahedron: Asymmetry 4, 1007–1016. BORNSCHEUER, U. T., ALTENBUCHNER, J., MEYER, H. H. (1999), Directed evolution of an esterase: Screening of enzyme libraries based on pH-indicators and a growth assay, Bioorg. Med. Chem. 7, 2169–2173. HECHT, H. J., SOBEK, H., HAAG, T., PFEIFER, O., P´EE, K. H. v. (1994), The metal-ion-free oxidoreductase from Streptomyces aureoﬁaciens has an a/B hydrolase fold, Nature Struct. Biol. 1, 532–537. HENKE, E., BORNSCHEUER, U. T. (1999), Directed evolution of an esterase from Pseudomonas ﬂuorescens. Random mutagenesis by error-prone PCR or a mutator strain and identiﬁcation of mutants showing enhanced enantioselectivity by a resorufm-based ﬂuorescence assay, Biol. Chem. 380, 1029–1033. JANES, L. E., KAZLAUSKAS, R. J. (1997), Quick E. A fast spectrophotometric method to measure the enantioselectivity of hydrolases, J. Org. Chem. 62, 4560–4561. ALTAMIRANO, M. M., BLACKBURN, J. M., AGUAYO C., FERSHT, A. R. (2000), Directed evolution of new catalytic activity using the a/B-barrel scaffold, Nature 403, 617–622. J¨URGENS, C., STROM, A., WEGENER, D., HETTWER, S., WILMANNS, M., STERNER, R. (2000), Directed evolution of a (αβ)8 -barrel enzyme to catalyze related reactions in two different metabolic pathways, Proc. Natl. Acad. Sci. USA 97, 9925–9930. STEVENSON, J. D., LUTZ, S., BENKOVIC, S. J. (2001), Retracing enzyme evolution in the (αβ)8 -barrel scaffold, Angew. Chem. Int. Ed. 113, 1906–1908. ZHANG, J. H., DAWES, G., STEMMER, W. P. (1997), Directed evolution of a fucosidase from a galactosidase by DNA shufﬂing and screening, Proc. Natl. Acad. Sci. USA 94, 4504–4509. STEFAN, A., RADEGHIERI, A., GONZALEZ VARAY RODRIGUEZ, A., HOCHKOEPPLER, A.

33

34

35

36

37

38

39

40

41

(2001), Directed evolution of β-galactosidase from Escherichia coli by mutator strains defective in the 3 →5 exonuclease activity of DNA polymerase III, FEBS Lett. 493, 139–143. LI, Q.-S., SCHWANEBERG, U., FISCHER, P., SCHMID, R. D. (2000), Directed evolution of the fatty acid hydroxylase P450 BM-3 into an indole-hydroxylating catalyst, Chem. Eur. J. 6, 1531–1536. SIO, C. F., LAAN, J. M. v. d., RIEMENS, A. M., QUAX, W. J. (2000), Altering substrate speciﬁcity of Pseudomonas SY77 glutaryl acylase towards adipyl-7-ADCA, Biotechnology 2000 meeting, Berlin JOO, H., ARISAWA, A., LIN, Z., ARNOLD, F. H. (1999), A high-throughput digital imaging screen for the discovery and directed evolution of oxygenases, Chem. Biol. 6, 699–706. ZHAO, H., GIVER L. AFFHOLTER, J. A., ARNOLD, F. H. (1998), Molecular evolution by staggered extension process (StEP) in vitro recombination, Nature Biotechnol. 16, 258–261. JOO, H., LIN, Z., ARNOLD, F. H. (1999), Laboratory evolution of peroxide-mediated cytochrome P450 hydroxylation, Nature 399, 670–673. KAMPEN, M. D. v., EGMOND, M. R. (2000), Directed evolution: From a staphylococcal lipase to a phospholipase, Eur. J. Lipid Sci. Technol. 102, 717–726. KAUFFMANN, I., SCHMIDT-DANNERT, C. (2001), Conversion of Bacillus thermocatenulatus lipase into an efﬁcient phospholipase with increased activity towards long chain fatty acyl substrates by directed evolution and rational design, Protein Eng 14, 919–928. IFFLAND, A., TAFELMEYER, P., SAUDAN C. JOHNSSON, K. (2000), Directed molecular evolution of cytochrome c peroxidase, Biochemistry 39, 10790–10798. FONG, S., MACHAJEWSKI, T. D., MAK, C. C., WONG, C.-H. (2000), Directed evolution of D-2-keto-3-deoxy-6-phosphogluconate aldolase to new variants for the efﬁcient synthesis of D- and L-sugars, Chem. Biol. 7, 873–883.

1

Design of Restriction Endonucleases with Novel Speciﬁcity Thomas Lanio, Albert Jeltsch, and Alfred Pingoud

Originally published in: Directed Molecular Evolution of Proteins. Edited by Susanne Brakmann, Kai Johnsson. Copyright ľ 2002 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30423-1

1 Introduction 1.1 Biology of restriction/modiﬁcation systems

Restriction/modiﬁcation systems (RM systems) can be regarded as an “immune” system to protect the host against foreign DNA, in particular bacteriophage DNA [1]. RM systems comprise two different enzyme activities directed against the same short, often palindromic, DNA sequence – (i) a methyltransferase activity that protects the host DNA by methylation of the recognition sequences and (ii) a restriction endonuclease activity, that cuts unmethylated recognition sequences usually present in foreign DNA (Fig. 1). Since erroneous cleavage of the host DNA would be highly toxic for the cell, restriction enzymes have evolved a remarkable speciﬁcity for their canonical recognition site. They are known to cleave “star” sites, which are DNA sequences differing only in one base pair from the canonical sequence, and also methylated recognition sites, by at least 5–6 orders of magnitude more slowly than their canonical recognition sequence [2–6]. With this speciﬁcity, restriction endonucleases are among the most speciﬁc enzymes known that do not employ an energy-dependent proofreading step. RM systems are very effective against bacteriophages and are ubiquitously distributed over the eubacterial and archaeal kingdoms. The observation that genes encoding RM systems may comprise up to 4% of the genome of a prokaryotic organism emphasizes the importance of these enzymes for their hosts [7]. In addition to their protective functions, the involvement of RM systems in recombination and transposition is still being discussed [8–10], as well as the concept that genes encoding these systems could be regarded as selﬁsh genetic elements [11]. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Design of Restriction Endonucleases with Novel Speciﬁcity

Fig. 1 Biological function of restriction/modiﬁcation systems: RM systems recognize and act on short palindromic DNA sequences. While the host genome is protected by the methyltransferase activity of the system, invading phage DNA is cleaved by the endonuclease activity.

1.2 Biochemical properties of type II restriction endonucleases

RM systems are usually divided into three different types (I, II and III), and then into subtypes, according to certain properties like cofactor requirement, subunit composition and mode of DNA cleavage. Among these three types, the type II restriction enzymes are the most important ones for molecular biology and biotechnology. Today, more than 3300 restriction endonucleases with more than 200 different speciﬁcities are known and their number is steadily increasing [12]. These enzymes have been studied over the last 30 years as model systems for speciﬁc protein-DNA interactions, in general, and enzymes acting on DNA, in particular (for reviews see [13, 14]). Besides many biochemical studies that were carried out to analyze the mechanism of DNA recognition and cleavage by these enzymes, crystal structures of 12 different type II restriction enzymes have been determined, in part in complex with DNA allowing for a detailed insight into the structure and function of this class of enzyme (for a review see [15]). The orthodox type II restriction endonuclease is a homodimer of around 2 × 30 kDa. It recognizes a palindromic sequence of 4–8 base pairs and cleaves both strands of its DNA substrate in the presence of Mg2+ ions within or directly adjacent to the recognition site, leading to 5 phosphate and 3 OH ends. Depending on the enzyme employed, the cleavage reaction results either in blunt ends (for example EcoRV), or sticky ends with a 5 overhang (for example EcoRI) or a 3 overhang (for example BglI, Fig. 2). An interesting subgroup is represented by the type IIS restriction endonucleases. These enzymes recognize an asymmetric site and cleave this sequence at a deﬁned distance after dimerizing on the substrate (for example FokI). Restriction enzymes constitute one of the largest families of related enzymes known so far and therefore exemplify how natural evolution has generated many different speciﬁcities from one (or a few) ancestral protein(s). A detailed comparison

1 Introduction 3

Fig. 2 Cleavage of the speciﬁc recognition sites by the type II restriction endonucleases EcoRV, EcoRI and BglI: The cleavage reaction, which requires Mg2+ as cofactor, leads to 5 phosphate and 3 OH ends. While EcoRV cleavage results in blunt ends, EcoRI and BglI generate sticky ends with a 5 and 3 overhang, respectively.

of the structures and functions of these enzymes will not only improve our understanding of natural evolution but also guide design approaches. Information regarding all known RM systems are available at REBASE (www.rebase.neb.com). 1.3 Applications for type II restriction endonucleases

The discovery of restriction endonucleases was one of the key events leading to and being a prerequisite for the development of modern gene technology. These enzymes are indispensable tools for DNA analysis and cloning as well as for medical applications where the study of restriction fragment length polymorphisms (RFLPs) provided ﬁrst insights into the genetic diversity of man as well as into the genetic basis of some diseases. Recently the use of restriction enzymes has been partly replaced by the PCR technique for certain applications but restriction enzymes still are the work horses in the molecular biology laboratory with a worldwide market of approximately 100–200 million US$. Today, an almost full set of enzymes with different speciﬁcities is commercially available for the 4 base pair and 6 base pair cutters (16 or 64, respectively), but only 9 out of 256 possible 8 base pair cutters were found. This under-representation of restriction enzymes with long recognition sites can be understood, because palindromic sites comprising 8 base pairs statistically occur only once every 65 536 base pairs and phage genomes usually are short (104 –105 base pairs). Therefore, a restriction enzyme with an 8 or even 10 base pair recognition site could not fulﬁll its biological role efﬁciently.

4 Design of Restriction Endonucleases with Novel Speciﬁcity

Restriction endonucleases having a recognition sequence of 8 or more base pairs, therefore, in general provide only a limited selective advantage for the bacterial host and in consequence to date only a few 8 base pair cutters were isolated from natural sources and most likely not many more will be found by screening bacterial strains. On the other hand these “rare cutters” with recognition sites longer than 8 base pairs would be very helpful for the manipulation of large DNA fragments such as human genes which are often larger than 5kb. The availability of such “rare cutters” would facilitate the generation of artiﬁcial chromosomes and perhaps make the manipulation of such constructs as easy as plasmid cloning is today. Moreover, such enzymes could be extremely useful for gene targeting and gene replacement, not only for experimental genetics but also for biomedical applications [16–19]. Currently, the only strategy available to introduce single cuts in large DNA fragments is the Achilles’ heel cleavage procedure. In this method, an enzyme is converted into a “rare cutter” by allowing speciﬁc cleavage only at one recognition site within a large substrate by protecting all other speciﬁc sites via methylation. This is achieved by incubating the DNA substrate with an oligodeoxyribonucleotide, which forms a triple-helix with the recognition site to be cleaved, thereby protecting the site. In the next step the DNA is incubated with the methyltransferase corresponding to the restriction endonuclease to be employed. The methyltransferase methylates all speciﬁc recognition sites of the plasmid except the single site protected by triple-helix formation. After inactivation of the methyltransferase the triple-helix is disrupted and the substrate is washed extensively. The single nonmethylated site is then cut by the restriction endonuclease. More recently instead of oligodeoxyribonucleotides, PNAs (peptide nucleic acids) have been used to improve triple-helix formation by sequence-speciﬁc strand invasion even at shorter recognition sequences. With PNA-assisted rare cleavage [20] a restriction enzyme was demonstrated to cut genomic DNA into a relatively small number of fragments in the range of Mbp. Instead of triplex-forming oligonucleotides or PNAs also other DNA binding protein (like transcription factors) may be used to protect one site from methylation. Employing this method it was for example, possible to cleave a single site in the yeast genome [21]. A major disadvantage of this method is the requirement for complete methylation of the substrate and, moreover, to prevent methylation of the single site of interest. In consequence the Achilles’ heel approach, in the absence of better methods, might be quite useful for DNA analysis, but not for the manipulation of whole genomes. 1.4 Setting the stage for protein engineering of type II restriction endonucleases

From what has been said above it is obvious that there is a great demand for restriction enzymes with long recognition sequences of 8 base pairs and more. In the last few years several projects were initiated towards achieving this goal by protein engineering, taking advantage of the fact that restriction endonucleases have not only been studied over the last 30 years but also introduced as model

2 Design of Restriction Endonucleases with New Speciﬁcities

systems for protein engineering in the late 1980s [22]. Whereas initial efforts were devoted to rational protein design, more recently the focus has shifted to directed evolution to change (or extend) the speciﬁcity of restriction endonucleases. In the present review we will summarize what has been achieved so far using these different approaches and present an outlook on future developments.

2 Design of Restriction Endonucleases with New Speciﬁcities 2.1 Rational design 2.1.1 Attempts to employ rational design to change the speciﬁcity of restriction enzymes A prerequisite for the rational design of proteins is detailed knowledge about the structure, or, even better, about the apoenzyme, the enzyme in the speciﬁc complex with its cognate substrate, and the enzyme in complex with competitive inhibitors (transition state analogs) (Fig. 3). Until now 12 different type II restriction endonucleases have been studied crystallographically (for a review see [15]). Of special interest are those studies which reveal the differences between the structures of the enzyme bound to its speciﬁc DNA, the enzyme bound to non-speciﬁc DNA, and the free enzyme. Even more valuable are crystallographic studies of enzymes complexed with slightly different (with respect to sequences ﬂanking the recognition site) cognate and near-cognate (with respect to the recognition site) substrates. The restriction endonuclease EcoRW (recognition site: GATATC) is one of the best characterized type II restriction enzymes. For this enzyme the structure of the free enzyme [23, 24], the structure of the enzyme bound non-speciﬁcally to DNA [23], and the structure of the enzyme bound to various cognate substrates [23, 25–27] as well as the structure of a product complex [25] are known. In addition, structures of EcoRV variants bound to speciﬁc substrates [28] and of EcoRV bound to modiﬁed substrates [29] have been determined. All these different structures, and those of other restriction endonucieases, show that the recognition of the speciﬁc site and the discrimination between the speciﬁc and non-speciﬁc sites is achieved by multiple contacts between enzyme and substrates. These contacts are not only formed between the enzyme and the bases of the recognition sequence (direct readout) [30] but also between the enzyme and the phosphodiester backbone of the DNA (indirect readout) [31]. With detailed structural knowledge available it was suggested that it should be possible to produce enzymes with new speciﬁcities by altering some of these contacts. However, all mutagenesis studies carried out in particular with EcoRI and EcoRV revealed that none of the mutants generated by rational design displayed the desired change in speciﬁcity (for a review see [32]). Instead, the substitution of amino acid residues involved in substrate recognition usually resulted in a severe loss of activity. Obviously, a complex interwoven network in the protein-DNA interface is formed cooperatively, leading to the accurate

5

6 Design of Restriction Endonucleases with Novel Speciﬁcity

Fig. 3 Correlation of required mechanistic information, required structural information, importance of screening and number of possible enzyme variants in protein engineering by rational protein design and directed evolution.

positioning of the catalytic amino acid residues of both enzyme subunits, which explains the very high level of speciﬁcity and accuracy. Any manipulation in this network seems to lead inevitably to a strong reduction in the catalytic rate [33, 34] and/or a relaxation of speciﬁcity [35], as shown for EcoRI. The observation that it was not possible to change the substrate speciﬁcity of any restriction enzyme might be attributed to our limited understanding of the process of DNA recognition by enzymes for two reasons: (i) we do not fully understand the dynamic structures of the speciﬁc enzyme-DNA complexes, which means that we have no reasonable estimates for the energies associated with any of the observed contacts and, more importantly, we have no quantitative information regarding the inﬂuence of one contact on all other contacts, i.e. the cooperativity of the interactions, (ii) We do not have sufﬁcient knowledge of alternative conformations of the enzyme-DNA complexes that might be adopted if one or more positions of the target sequence have been exchanged. Only a detailed understanding of these alternative conformations, which do not support catalytic activity, will allow the elucidation of the enthalpic and entropic origins of the extreme speciﬁcity of restriction endonucleases. However, the lack of success of rational design might also in part be due to the biology of these systems: restriction endonucleases, like all proteins, are synthesized by protein biosynthesis on the ribosomes in the cell. This process has a limited accuracy: about one wrong amino acid residue is incorporated per 103 –104 residues [36–38]. Assuming that there are 10 copies of a restriction enzyme in the cell (2 × 300 amino acid residues) more than 99 % of all cells will contain at least one restriction enzyme variant with at least one amino acid

2 Design of Restriction Endonucleases with New Speciﬁcities

exchange. Given the possibility that even one copy of an enzyme with an altered or relaxed speciﬁcity will cause severe problems, there must have been a strong evolutionary pressure to make restriction enzymes not only accurate but also secure in the sense that variants produced by errors in protein biosynthesis do not cleave an altered DNA sequence, i.e. a sequence unprotected by the methyltransferase of the given RM system. Rational design was applied with notable success only to the generation of variants which recognize and cleave modiﬁed substrates. For example, an EcoRI variant was obtained that had lost the ability to discriminate between thymine and uracil at the forth position of the EcoRI recognition sequence (GAAUTC and GAATTC) [39]. An EcoRV variant was produced in which a single enzyme-substrate contact was altered by site-directed mutagenesis leading to a 220-fold higher selectivity for GATAUC over GATATC modiﬁed substrates [40]. The largest alteration of the speciﬁcity of a restriction enzyme achieved to date was obtained with another EcoRV variant. This variant cleaves a modiﬁed substrate in which the GATATpC phosphate group is replaced by a methylphosphonate, with the same rate as the wild-type enzyme, which cleaves modiﬁed and non-modiﬁed substrate at almost the same rate. The variant, however, prefers the modiﬁed over the non-modiﬁed substrates by more than three orders of magnitude [41]. As none of the modiﬁcations targeted by these variants occurs in regular DNA, the described variants can only serve as the proof of principle that new enzyme-substrate contacts can be generated by rational design. Changing the target speciﬁcity of a restriction enzyme, however, would require the reengineering of 3–4 contacts of the enzyme to the DNA (co-crystal structure analyses of speciﬁc restriction enzyme-DNA complexes tell us that on average each base pair of the recognition sequence is involved in four speciﬁcity-determining contacts) against the evolutionary pressure that exists to prevent the generation of variants with new speciﬁcities by chance! 2.1.2 Changing the substrate speciﬁcity of type IIs restriction enzymes by domain fusion Type IIs restriction endonucleses are composed of a DNA recognition domain and a DNA cleavage domain. Whereas the DNA cleavage domain of FokI, a type IIs enzyme, has a typical restriction endonuclease fold [42], the structure of the DNA recognition domain is completely unrelated and resembles that of the catabolite activator protein [43]. The separation of recognition and cleavage in two different subunits can be employed to generate a chimeric nuclease with altered speciﬁcity by genetically fusing other non-related DNA binding domains to the FokI catalytic domain [44]. The most successful class of chimeric nucleases is based on the linkage of the cleavage domain from FokI and a DNA binding domain consisting of three zinc ﬁngers [45]. Since each of the zinc ﬁngers binds to three base pairs, the length and sequence of the DNA to be recognized can be addressed by modulation of the number and nature of the amino acids residues in contact with the DNA. Similar fusions have been prepared with transcription factors of the homeodomain [46] and Gal4-type [47]. Efﬁcient double strand cleavage is only observed if two copies of the recognition sequence are in close proximity and in the correct orientation,

7

8 Design of Restriction Endonucleases with Novel Speciﬁcity

reﬂecting the requirement for dimerization of the cleavage domain of FokI [48]. While native FokI cleaves 9 and 13 base pairs away from its recognition site the dimerized cleavage domains sometimes produce cuts with overhangs other than four base pairs, which could be explained by the uncoupling of substrate recognition and catalysis. A potential application of these chimeric nucleases could be the stimulation of homologous recombination in vivo by introducing a double strand break and thereby producing recombinogenic ends [49]. Similar approaches have been explored for other non-speciﬁc nucleases, like DNasel. However, none of the alternative chimeric nucleases showed a similar degree of speciﬁcity as the FokI chimera although it was possible to change the cleavage preferences of DNasel signiﬁcantly [50]. The reason for this difference might be that DNasel is a very active nuclease whereas the catalytic domain of FokI only has a low catalytic activity. Therefore, with DNasel DNA cleavage always occurs also at non-targeted sites, which does not happen as easily with FokI. 2.1.3 Rational design to extend speciﬁcities of type II restriction enzymes A much more promising approach than to employ rational design for the change of a speciﬁcity is the extension of existing speciﬁcities. As we know from the published co-crystal structures and biochemical studies, restriction endonucleases contact not only their recognition sequence but also some additional base pairs. In the case of EcoRV it was shown that, besides the contacts to the nucleotides of the recognition sequence, the protein interacts non-speciﬁcally with the backbone and the bases upstream and downstream of the GATATC site [23]. These contacts are largely water-mediated and are considered to be responsible for the dependence of the cleavage rate on the sequences ﬂanking the recognition sequence. This ﬂanking sequence effect suggested a promising starting point for a rational protein design project. It seemed to be possible to engineer new speciﬁc protein-DNA contacts outside of the GATATC site enabling the nuclease to recognize an extended site of up to 10 base pairs. This idea was put forward and tested by Schottler et al. [51]: they exchanged an amino acid residue involved in a water-mediated contact to the base on the 5 -side of the GATATC recognition sequence against all naturally occurring amino acid residues. Some variants, having amino acid residues with long or bulky side chains at that position show altered preferences, namely an extended speciﬁcity for PuGATATCPy (Fig. 4). More recently, on the basis of crystal structures obtained with EcoRV and oligonucleotides with a canonical site but different ﬂanking sequences, amino acid residues which are involved in contacts to the ﬂanking sequences were identiﬁed, and substitutions suggested which could possibly discriminate between different sequences [28]. However, the detailed biochemical characterization of the mutants led to disappointing results, because only one had a considerably altered selectivity, which, however, was not the predicted one [52]. Obviously it is very difﬁcult to predict amino acid substitutions that would lead to an extended speciﬁcity of a restriction enzyme. This does not necessarily mean that additional speciﬁc contacts cannot be designed, but more likely that the starting point of the design is not well-deﬁned. In most of the co-crystals obtained so far,

2 Design of Restriction Endonucleases with New Speciﬁcities

Fig. 4 Contacts of EcoRV to the base pairs ﬂanking the recognition sequence. Exchange of Ala 181 (top) to Lys (bottom) leads to a variant with a strong preference for purine residues 5 to the recognition sequence [51].

the enzymes cannot be activated by Mg2+ ions soaked into the crystal. Thus, the structures do not really represent the speciﬁc complex as it is formed before DNA cleavage. Moreover, no structural information on the transition state is available with respect to the speciﬁcity determining step and in consequence, predictions regarding the effects of amino acid residue substitutions are unreliable. The lack of our understanding of the key steps of the different conformational changes that are required to couple recognition to catalysis, has so far prevented successful rational design, not only of enzyme variants with an altered speciﬁcity but also of those with an extended speciﬁcity. 2.2 Evolutionary design of extended speciﬁcities

A possible way to overcome these limitations might be a “non-rational” approach by repertoire selection methods which have been shown to be extremely useful for developing new enzyme speciﬁcities. With a combination of random mutagenesis and selection it was possible to change enzyme speciﬁcities for a wide range of different purposes, even in the absence of detailed structural information (Fig. 3). One of the most successful techniques proved to be the shufﬂing method which was

9

10 Design of Restriction Endonucleases with Novel Speciﬁcity

introduced by Stemmer in 1994 [53] leading to the molecular breeding approach, which was published a few years later by the same group [54]. In the last few years a variety of enzymes was improved regarding stability, solubility, substrate speciﬁcity and other properties using this and other methods of directed evolution [55], which all make use of large libraries of up to 109 –1010 different variants. To ﬁnd the variants of interest, namely those with the desired improved or altered speciﬁcity, a reliable high-throughput assay or a stringent in vivo assay must be available (Fig. 3). Even more important is the coupling of phenotype to genotype in order to facilitate recovery of the genes of interest. This can be achieved in vivo, because in the cell the synthesis of each protein is instructed by the cellular DNA, or in vitro by using phage display [56], ribosome display [57] or coupled transcription/translation in water-oil emulsions [58]. The application of directed evolution approaches for the change or extension of the speciﬁcity of a restriction enzyme is hampered by the fact that an in vivo selection assay is not available and that examination of endonuclease activity in vitro usually requires puriﬁcation of the enzymes to avoid background activity by other nucleases prevalent in all cells. This means that an altered or extended speciﬁcity can only be observed with sufﬁciently puriﬁed protein preparations, thereby unfortunately separating genotype and phenotype. As neither a reliable in vivo test nor the secretion of restriction endonucleases followed by a colony assay could be established in the past, experiments with larger libraries of these enzymes require a solid clone management, a fast and uncomplicated, nevertheless speciﬁc, puriﬁcation scheme and a fast and robust assay method. Such an approach was chosen to extend the recognition sequence of EcoRV, for example from GATATC to GGATATCC or to AGATATCT. For this enzyme, the expression of His6 -tagged fusion proteins is established, enabling protein puriﬁcation via afﬁnity chromatography on a Ni2+ -NTA column or Ni2+ -NTA covered magnetic beads, and assays have been worked out to test for extended speciﬁcities using substrates with two differently ﬂanked GATATC sites. As randomization of a whole gene results in libraries far too large to be screened in vitro, for EcoRV a structure guided semi-rational approach was employed. The cocrystal structure of EcoRV reveals different regions of the enzyme which are in close proximity (<0.5 mm) to the nucleotide directly ﬂanking the recognition site on the 5 end. The three regions identiﬁed are between four and eight amino acid residues in length and located in different parts of the enzyme (Fig. 5). To limit the number of possible variants and to make an evolutionary approach possible, the amino acid residues in the three regions were randomized and screened for the desired speciﬁcity in three subsequent rounds of mutagenesis and selection, employing the genes of the selected variants of the ﬁrst cycle as template for the second mutagenesis step and so forth (Fig. 6). For that purpose, a mutagenesis protocol was developed to ensure that each mutation cycle resulted in additional mutations instead of only accumulating variants that originated from the template pool [59]. Employing this method and screening more than 500 different variants, generated in three subsequent cycles, several variants were found which cleave differently ﬂanked EcoRV recognition sites with more than 10-fold different rates. Compared to the wild-type enzyme, the

2 Design of Restriction Endonucleases with New Speciﬁcities

Fig. 5 Structure of a speciﬁc EcoRV-DNA complex. The regions subjected to mutagenesis as described in Lanio et al [59] are highlighted in red (amino acid residues 95 to 104), green (amino acid residues 180 to 184) and blue (amino acid residues 219 to 226). In B only the regions subjected to mutagenesis are displayed, together with the DNA.

11

12 Design of Restriction Endonucleases with Novel Speciﬁcity

Fig. 6 Schematic overview of the in vitro evolution cycle employed. The locations of the regions subjected to mutagenesis are shown in Fig. 5. The substrate employed to assay the activity and preference of the variants was a prelinearized plasmid harboring two differently ﬂanked EcoRV

recognition sites (CGGGATATCGTC and AAAGATATCTTT). Both sites are cleaved by wild-type EcoRV with identical rates, such that two cleavage intermediates are formed. Mutants that prefer one of the sites produce both intermediates in unequal amounts.

selectivity for sites with differently ﬂanked sequences has been increased by two orders of magnitude [60]. During the experiment not only the desired preferences increased, but also the number of variants displaying this preference. To further improve these results, the genes of the most successful variants were mixed and cleaved with two restriction endonucieases, thereby separating the three mutated regions. Religation of the fragments yielded recombinants with new combinations of beneﬁcial mutations. Interestingly, some of the resulting variants had lost their extended speciﬁcity, indicating that individual beneﬁcial substitutions can neutralize each other. However, this last shufﬂing step also resulted in EcoRV variants with further improved properties, namely variants which differ in their activity towards differently ﬂanked GATATC sites by more than two orders of magnitude: the non-preferred site is only attacked after complete cleavage of the preferred AGATATCT site (Fig. 7). A careful look at the wild-type enzyme cocrystal structure did not reveal an obvious explanation for the effects of the different amino acid substitutions, neither for a single substitution itself, nor for the combination of several different substitutions. As these substitutions most probably affect the transition state whose formation is presumed to be the speciﬁcity determining step, even a cocrystal structure of EcoRV variants with differently ﬂanked substrates might not really improve our ideas about how to extend speciﬁcity.

2 Design of Restriction Endonucleases with New Speciﬁcities

Fig. 7 Cleavage experiments with prelinearized substrate and the EcoRV variant with the most pronounced preference for AT ﬂanked recognition sites (Y95H; K98E; E99V; N100T; S183A; Q224K). Substrate (4nM) (see Fig. 6) was incubated with enzyme (3 nM). Lane 0 = length marker, 1–13 = 1 , 3 , 5 , 6 , 10 , 30 , 60 , 90 , 120 , 150 , 170 —addition of new substrate to

exclude that the variant lost activity during the extended incubation time – 180 , 190 . Note that the substrate ABC is cleaved very rapidly into the two intermediate products AB and C – the CC-ﬂanked recognition site in intermediate AB is only cleaved after complete cleavage of the preferred ATﬂanked recognition site, to give the ﬁnal products A, B and C.

Although in the course of the experiments described only single and double mutants were introduced per cycle, the number of variants screened was several orders of magnitude lower than the number of variants generated. Since the bottleneck of the whole procedure was the isolation and puriﬁcation of the enzyme variants, a high-throughput process was developed in order to allow parallel protein expression, puriﬁcation and subsequent characterization of 96 protein variants in an automated protocol in micro-plate format [61] (Fig. 8). To test the reliability of the system three codons of the EcoRV gene were randomized using a spiked oligodeoxynucleotide yielding a library of about 1200 different clones. 1056 different clones, corresponding to an almost complete representation of the library, were inoculated and expressed in 96-well blocks. The puriﬁcation of the proteins after cell lysis is based on the interaction between the His6 -tagged EcoRV variants and Ni2+ -NTA coated microplate allowing for the puriﬁcation of 10 pmol protein per well. The functional characterization of the variants was performed with the assays previously described (Figs. 6 and 7). Since the whole process is done automatically by a pipetting robot it is possible to characterize 103 different clones within a week and 104 –105 clones in a reasonable period of time. In the relatively small library described several variants displayed a more than 10-fold extended speciﬁcity against differently ﬂanked EcoRV recognition sites [61]. In the future, similar approaches will be made easier by the availability of an assay that is suited for high-throughput sample analysis [62]. Rather than using macromolecular DNA substrates and analyzing their cleavage by electrophoresis,

13

14 Design of Restriction Endonucleases with Novel Speciﬁcity

Fig. 8 Puriﬁcation of 96 different enzyme variants in a highthroughput approach. Cells are grown, harvested and lysed in microplates. His6 -tagged variants are isolated by transferring the lysate to Ni-NTA coated microplates from which they are eluted after washing. Speciﬁc enzyme activity can be checked by suitable assays.

a 96-well microplate-based ﬂuorescence assay has been developed employing two oligodeoxynucleotides with differently ﬂanked recognition sites and labeled with different ﬂuorescence markers. Before cleavage the ﬂuorescence is quenched. Upon cleavage, the ﬂuorescence of the preferentially cleaved substrate increases. Recording the ﬂuorescence of the two differently labeled substrates allows a rapid automated analysis of variants with an extended speciﬁcity (Fig. 9).

3 Conclusions 15

Fig. 9 Schematic overview of a ﬂuorescence-based cleavage assay for determination of cleavage preferences of restriction enzymes. The two substrates employed harbor differently ﬂanked recognition

sites and carry different labels whose ﬂuorescence is quenched before cleavage. Upon cleavage the ﬂuorescence of the preferentially cleaved substrate increases.

3 Conclusions

In the past two decades different strategies have been employed to change or to extend the speciﬁcity of restriction enzymes. Since substrate recognition is highly redundant and recognition itself coupled to catalysis via a multitude of intra- and intermolecular interactions, which are not really understood, all rational design projects, by and large, have failed so far. The rare positive results concerned variants with preferences for artiﬁcial substrates. Taking these results into account rational design turns out to be ill-suited for the alteration of substrate speciﬁcity of restriction enzymes at present. Promising results were obtained with chimeric nucleases, in which combination of the cleavage domain of the type IIs restriction endonuclease FokI was fused to different DNA binding proteins. This technique was successfully employed to initiate homologous recombination in vivo, but since the uncoupling of recognition and catalysis leads to different single strand overhangs these chimeric endonucleases are not suitable for cloning experiments which require a precise double strand break at a deﬁned position with deﬁned overhangs. As this can only be achieved by restriction enzymes, the Achilles’ heel approach is the superior technique, especially for cloning experiments. The requirement for complete methylation of the DNA at all sites that should not be cleaved by the restriction enzyme, prevents the usage of this for DNA manipulation in vivo. The extension of recognition sequences of type II restriction endonucleases also turned out to be a more than difﬁcult job. The function of the different contacts within the protein DNA interface is still poorly understood, as is the mechanism of how substrate recognition is coupled to catalysis. Presumably, the most serious deﬁcit for attempts to alter the speciﬁcity of restriction enzymes is the lack of detailed knowledge of the transition state of the DNA cleavage reaction. In

16 Design of Restriction Endonucleases with Novel Speciﬁcity

consequence, one does not really know what the speciﬁcity determining step looks like and how the catalytic centers are activated during the recognition process. This lack of information could be bypassed by an evolutionary design approach which does not presuppose this information. Unfortunately, for directed evolution large libraries of clones have to be generated, which demands a stringent selection of the clones coding for the gene product with the desired property. As screening for restriction enzyme variants with extended cleavage speciﬁcities is not yet possible in vivo, a protocol for expression, puriﬁcation and characterization of the different variants is required. This limits the size of the library. It was shown recently, however, that by employing nucleoside analogues in an error-prone PCR, even relatively small libraries of variants that carry many amino acid exchanges could yield the same variants that are otherwise only generated by gene shufﬂing [63]. Libraries comprising up to 105 different clones could be expressed and screened with recently established methods [61] and the high-throughput generation and characterization of restriction enzyme variants with extended speciﬁcities could be further improved. Acknowledgements

Research on restriction enzymes in the authors’ laboratory was supported by the Deutsche Forschungsgemeinschaft, the Bundesministerium fur Bildung, Wissenschaft, Forschung und Technologie, the European Union, and by the Fonds der Chemischen Industrie.

References 1 T. A. BICKLE, D. H. KRUGER, Microbiol. Rev. 1993, 57, 434. 2 D. R. LESSER, M. R. KURPIEWSKI, L. JEN-JACOBSON, Science 1990, 250, 776. 3 V. THIELKING, J. ALVES, A. FLIESS, G. MAASS, A. PINGOUD, Biochemistry 1990, 29, 4682. 4 J. ALVES, U. SELENT, H. WOLFES, Biochemistry 1995, 34, 11191. 5 M. NELSON, E. RASCHKE, M. MCCLELLAND, Nucleic Acids Res. 1993, 21, 3139. 6 L. JEN-JACOBSON, L. E. ENGLER, D. R. LESSER, M. R. KURPIEWSKI, C. YEE, B. MCVERRY, Emb J 1996, 25, 2870. 7 L. F. LIN, J. POSFAI, R. J. ROBERTS, H. KONG, Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 2740. 8 J. HEITMAN, Genet. Eng. 1993, 15, 57. 9 M. MCKANE, R. MILKMAN, Genetics 1995, 139, 35. 10 K. CARLSON, L. D. KOSTURKO, Mol. Microbiol. 1998, 27, 671.

11 T. NAITO, K. KUSANO, I. KOBAYASHI, Science 1995, 267, 897. 12 R. J. ROBERTS, D. MACELIS, Nucleic Acids Res. 2001, 29, 268. 13 R. J. ROBERTS, S. E. HALFORD, in Nucleases, 2nd edition (Eds.: S. M. LINN, R. S. LLOYD, R. J. ROBERTS), Cold Spring Harbor Laboratory Press, Cold Spring Harbor, 1993, pp. 35. 14 A. PINGOUD, A. JELTSCH, Eur. J. Biochem. 1997, 246, 1. 326 13 Evolutionary Generation versus Rational Design of Restriction Endonucleases with Novel Speciﬁcity 15 A. PINGOUD, A. JELTSCH, Nucleic. Acids Res. 2001, 29, 3705. 16 A. CHOULIKA, A. PERRIN, B. DUJON, J. F. NICOLAS, Mol. Cell. Biol. 1995, 15, 1968. 17 B. ELLIOTT, C. RICHARDSON, J. WINDERBAUM, J. A. NICKOLOFF, M. JASIN, Mol. Cell. Biol. 1998, 18, 93.

References 18 M. COHEN-TANNOUDJI, S. ROBINE, A. CHOULIKA, D. PINTO, F. EL MARJOU, C. BABINET, D. LOUVARD, F. JAISSER, Mol. Cell. Biol. 1998, 18, 1444. 19 C. RICHARDSON, M. E. MOYNAHAN, M. JASIN, Genes Dev. 1998, 12, 3831. 20 K. I. IZVOLSKY, V. V. DEMIDOV, N. O. BUKANOV, M. D. FRANK-KAMENETSKII, Nucleic Acids Res. 1998, 26, 5011. 21 M. KOOB, W. SZYBALSKI, Science 1990, 250, 271. 22 J. ROSENBERG, B. WANG, C. FREDERICK, N. REICH, P. GREENE, J. GRABLE, J. MCCLARIN, in Protein Engineering, (Eds.: D. OXENDER, C. F. FOX), AR. Liss Inc., 1987, pp. 237. 23 F. K. WINKLER, D. W. BANNER, C. OEFNER, D. TSERNOGLOU, R. S. BROWN, S. P. HEATHMAN, R. K. BRYAN, P. D. MARTIN, K. PETRATOS, K. S. WILSON, EMBO J. 1993, 12, 1781. 24 J. J. PERONA, A. M. MARTIN, J. Mol. Biol. 1997, 273, 207. 25 D. KOSTREWA, F. K. WINKLER, Biochemistry 1995, 34, 683. 26 N. C. HORTON, J. J. PERONA, J. Mol. Biol. 1998, 277, 779. 27 N. C. HORTON, J. J. PERONA, Proc Natl. Acad. Sci. U. S. A. 2000, 97, 5729. 28 N. C. HORTON, J. J. PERONA, J. Biol. Chem. 1998, 273, 21721. 29 N. C. HORTON, M. D. SAM, B. A. CONNOLLY, J. J. PERONA, Biophys. J. 2000, 78, 417A. 30 N. C. SEEMAN, J. M. ROSENBERG, A. RICH, Proc Natl. Acad. Sci. U. S. A. 1976, 73, 804. 31 Z. OTWINOWSKI, R. W. SCHEVITZ, R. G. ZHANG, C. L. LAWSON, A. JOACHIMIAK, R. Q. MARMORSTEIN, B. F. LUISI, P. B. SIGLER, Nature 1988, 335, 321. 32 A. JELTSCH, C. WENZ, W. WENDE, U. SELENT, A. PINGOUD, Trends Biotechnol. 1996, 14, 235. 33 J. ALVES, T. RTITER, R. GEIGER, A. FLIESS, G. MAASS, A. PINGOUD, Biochemistry 1989, 28, 2678. 34 R. GEIGER, T. RTITER, J. ALVES, A. FLIESS, H. WOLFES, V. PINGOUD, C. URBANKE, G. MAASS, A. PINGOUD, A. DUSTERHOFT, M. KROGER, Biochemistry 1989, 28, 2667. 35 H. FLORES, J. OSUNA, J. HEITMAN, X. SOBERON, Gene 1995, 157, 295. 36 R. B. LOFTFIELD, D. VANDERJAGT, Biochem. J. 1972, 128, 1353.

37 P. EDELMANN, J. GALLANT, Cell 1977, 10, 131. 38 J. PARKER, T. C. JOHNSTON, P. T. BORGIA, G. HOLTZ, E. REMAUT, W. FIERS J. Biol. Chem. 1983, 258, 10007. 39 A. JELTSCH, J. ALVES, T. OELGESCHLAGER, H. WOLFES, G. MAASS, A. PINGOUD J. Mol. Biol. 1993, 229, 221. 40 C. WENZ, U. SELENT, W. WENDE, A. JELTSCH, H. WOLFES, A. PINGOUD, Biochim. Biophys. Acta 1994, 1219, 73. 41 T. LANIO, U. SELENT, C. WENZ, W. WENDE, A. SCHULZ, M. ADIRAJ, S. B. KATTI, A. PINGOUD, Protein Eng. 1996, 9, 1005. 42 D. A. WAH, J. A. HIRSCH, L. F. DORNER, I. SCHILDKRAUT, A. K. AGGARWAL, Nature 1997, 388, 97. 43 S. C. SCHULTZ, G. C. SHIELDS, T. A. STEITZ, Science 1991, 253, 1001. 44 S. CHANDRASEGARAN, J. SMITH, Biol. Chem. 1999, 380, 841. 45 J. SMITH, M. BIBIKOVA, F. G. WHITBY, A. R. REDDY, S. CHANDRASEGARAN, D. CARROLL, Nucleic Acids Res. 2000, 28, 3361. 46 Y. KIM, S. CHANDRASEGARAN, Proc Natl. Acad. Sci. U. S. A. 1994, 91, 883. 47 Y.-G. KIM, J. SMITH, M. DURGESHA, S. CHANDRASEGARAN, Biol. Chem. 1998, 379, 489. 48 J. BITINAITE, D. A. WAH, A. K. AGGARWAL, I. SCHILDKRAUT, Proc Natl. Acad. Sci. U. S. A. 1998, 95, 10570. 49 M. BIBIKOVA, D. CARROLL, D. J. SEGAL, J. K. TRAUTMAN, J. SMITH, Y. G. KIM, S. CHANDRASEGARAN, Mol. Cell. Biol. 2001, 21, 289. 50 S. CAL, K. L. TAN, A. MCGREGOR, B. A. CONNOLLY, EMBO J. 1998, 17, 7128. 51 S. SCHOTTLER, C. WENZ, T. LANIO, A. JELTSCH, A. PINGOUD, Eur. J. Biochem. 1998, 258, 184. 52 T. LANIO, A. JELTSCH, A. PINGOUD, Protein Eng. 2000, 13, 275. 53 W. P. STEMMER, Nature 1994, 370, 389. 54 A. CRAMERI, S. A. RAILLARD, E. BERMUDEZ, W. P. STEMMER, Nature 1998, 391, 288. 55 F. H. ARNOLD, P. L. WINTRODE, K. MIYAZAKI, A. GERSHENSON, Trends Biochem. Sci. 2001, 26, 100. 56 T. CLACKSON, H. R. HOOGENBOOM, A. D. GRIFFITHS, G. WINTER, Nature 1991, 352, 624.

17

18 Design of Restriction Endonucleases with Novel Speciﬁcity

57 J. HANES, A. PLU¨ CKTHUN, Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 4937. 58 D. S. TAWFIK, A. D. GRIFFITHS, Nat. Biotechnol. 1998, 16, 652. 59 T. LANIO, A. JELTSCH, Biotechniques 1998, 25, 958. 60 T. LANIO, A. JELTSCH, A. PINGOUD, J. Mol. Biol. 1998, 283, 59.

61 T. LANIO, A. JELTSCH, A. PINGOUD, Biotechniques 2000, 29, 338. 62 K. EISENSCHMIDT, Diploma Thesis, Justus-Liebig Universitat (Giessen), 2000. 63 M. ZACCOLO, E. GHERARDI J. Mol. Biol. 1999, 285, 775.

1

S. Dedeblatt

Stefan Lutz, and Stephen J. Benkovic* The Pennsylvania State University, University Park, USA

Orignally published in: Directed Molecular Evolution of Proteins. Edited by Susanne Brakmann, Kai Johnsson. Copyright ľ 2002 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30423-1

1 Introduction

Proteins are involved in virtually every aspect of a living organism, yet they are composed of only twenty natural amino acids. The protein’s tremendous diversity originates from its structural and functional versatility, achieved through polymerization of the chemically distinct amino acid building blocks into sequences of varying composition and length. Upon folding into complex three-dimensional structures, these macromolecules turn into molecular machines with the ability to accelerate even the most complex chemical reactions by many orders of magnitudes with unsurpassed selectivity and speciﬁcity. The superior performance of proteins in respect to functional diversity, speciﬁcity, and efﬁciency has attracted scientists and engineers alike to create tailormade catalysts by protein engineering. Customizing a protein towards practical applications such as industrial catalysts, biosensors, and therapeutics offers many economic incentives. At the same time, protein engineering can address fundamental questions of protein design in respect to their structure and function. This can then lead to the development of new technologies facilitating the more directed exploration of protein sequence space. A readily available and abundant resource for examples on how to engineer proteins is Nature itself. The mechanisms of natural protein evolution have repeatedly succeeded in adapting protein function to ever changing environments. Understanding the principles and motives whereby new protein structures and functions emerge can provide useful guidelines for protein engineering. This chapter is directed towards the mechanisms of natural protein evolution, their implications on structure and function, and their experimental implementation in vitro.

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 S. Dedeblatt

2 Mechanisms of Protein Evolution in Nature

Two fundamental approaches can be employed to generate novel proteins; i) the de novo arrangement of small, structured peptides or ii) the remodeling of an existing protein framework. In computational simulations, the de novo assembly pathway demonstrated its effectiveness for the creation of novel functionality in proteins [1]. However, successful in vitro experiments are lacking and evidence for the occurrence of such a pathway in Nature has been limited to a family of antifreeze glycoproteins, found in arctic and antarctic ﬁsh ([2] and references therein). Having an Ala-Ala-Thr tripeptide (with minor sequence variations) as the fundamental building blocks, the polymeric structures presumably evolved by repeated duplication of a 9-basepair segment. Although antifreeze proteins are believed to have emerged during the geologically recent glaciation period, de novo assembly per se lost its signiﬁcance after the early days of protein evolution. The sparse distribution of functionality in sequence space makes the repeated de novo creation of complex structures very inefﬁcient and extremely difﬁcult if not impossible within the given time constrains (the last 4 billion years). Instead, remodeling of existing protein scaffolds represents a more efﬁcient way to evolve novel functions and properties. Minor changes to explore the local sequence space by mutagenesis of individual positions within an established structure have proven highly successful in accommodating small environmental changes and in optimizing existing functions. The efﬁciency of such an approach is exempliﬁed by the ﬁne-tuning of antibody binding speciﬁcity by hyper-mutation of a set of precursors. Similarly, rationally designed single-amino acid changes in isocitrate dehydrogenase [3], lactate dehydrogenase from Trichomonas vaginalis [4], and T4 lysozyme [5] each resulted in a speciﬁc alteration in substrate speciﬁcity. Finally, a several-fold improvement of thermostability of subtilisin was observed upon introduction of two point mutations [6]. On the other hand, major rearrangements of proteins through duplication, circular permutation, fusion, as well as deletion and insertion of large fragments create entirely novel structural combinations with potentially novel function [7] (Fig. ??). Such radical reorganization of genetic material results in the exploration of extended sequence space and accelerated evolution. 2.1 Gene duplication

Reorganization, optimization, and development of novel function by duplication of genetic material is one of the fundamental mechanisms of protein evolution. Evidence for gene duplication can be found in small repetitive structures such as the previously discussed antifreeze proteins up to entire chromosomes and genomes. The unexpectedly high estimates of duplication frequency in eukaryotes, based on the analysis of several genomic databases, underline its signiﬁcance in evolutionary terms [8]. Mechanistically, gene duplication can originate during chromosomal DNA replication or as a result of unequal crossing-over during

2 Mechanisms of Protein Evolution in Nature

Fig. 1 Schematic overview of mechanisms for major structural reorganization during protein evolution. (Adapted from [49])

meiosis [9]. Depending on the mechanism, either a tandem repeat or two isolated copies of the parental sequences are generated (Fig. ??). In both cases, the copies are confronted with three possible outcomes: one of the two genes is i) inactivated by accumulation of mutation, ii) released from functional constrains which facilitates the accumulation of local mutations and leads to functional differentiation, or iii) both gene products maintain wild-type activity [8]. While gene inactivation has no obvious beneﬁcial effect for the host organism, the presence of multiple functional copies of the same gene can amplify its expression level which may be advantageous by accelerating metabolic ﬂux or increasing production of defensive compounds [10]. Finally, negotiating the ﬁne line between complete inactivation and conservation of wild-type activity, duplication followed by functional differentiation can lead towards the evolution of novel function. However, considering the sensitivity of protein structure and function to mutagenesis, divergence of the duplicate by random drift most likely results in enzyme inactivation. The acquisition of a different function in contrast would be a relatively rare event. A recent modeling study, however, suggested that the accumulation of complementary degenerative mutations in both gene duplicates could actually beneﬁt the host by facilitating the long-term preservation of the duplicated genes. The extended conservation time of both copies increases the probability of evolving novel function [11]. An alternative view of the evolution of functionally novel proteins includes generecruitment [12] or gene sharing [13]. In this model, a period of bifunctionality of a single gene product precedes the duplication event upon which one copy maintains the ancestral function while the other adopts the new function. The

3

4 S. Dedeblatt

various aspects of bifunctionality can hereby range from conservation of structural properties as demonstrated for crystallins [14] to what O’Brian and Herschlag [15] call promiscuous activity in ancestral structures. Independent of the actual mechanism, functional divergence between two proteins with a common ancestor is exempliﬁed in the βα-barrel structures of N-(phosphoribosyl-formimino)-aminoimidazole-carboxamide ribonucleotide isomerase (HisA) and imidazole glycerol phosphate synthase (HisF) of the histidine biosynthetic pathway [16]. The hypothesis of a common origin of the two enzymes was formulated based on sequence comparison [17] and has recently found support by the identiﬁcation of extensive structural similarities [18]. 2.2 Tandem duplication

In contrast to locally separated duplication products, tandem duplication of a gene results in the expression of two or more copies of the gene product within a single polypeptide chain (Fig. ??). Such multiplicity has, as reﬂected by the high frequency of its natural occurrence, clear advantages [7]: Ĺ Ĺ Ĺ Ĺ Ĺ

increased stability, new cooperative functions, formation of binding sites in a newly-formed cleft growth of long repetitive structures such as ﬁbrous proteins, and production of multiple binding sites in series that produce either more efﬁcient or more speciﬁc binding effects.

Furthermore, tandem repeats can undergo circular permutation and thereby reorganize the structural and functional integrity of the parental sequence. 2.2.1 Proteases Often-cited examples for tandem duplications are pepsin, a member of the aspartate protease family [19], and chymotrypsin, representative of the trypsin-like protease family [20]. Structurally, both enzymes consist of two homologous, stacked domains, β-sheets for pepsin and β-barrels for chymotrypsin, with the active site residues located at the domain interface. Despite extensive sequence divergence between the individual domain pairs, their overall backbone structures are mutually superimposable, strongly suggesting a single-domain homodimeric ancestor [21]. Direct evidence for the feasibility of such a primordial enzyme is provided by homodimeric aspartic proteases, found in retroviruses [22]. Similarly, the homodimeric β-barrel structure of the serine protease from cytomegalovirus, a member of the herpes virus family, may resemble an evolutionary precursor of modern serine proteases. Although crystallographic analysis clearly indicates no direct relation to the latter as revealed by signiﬁcant differences in structural arrangement and the catalytic triad [23], the underlying design concept of using a two-barrel structure to form a functional enzyme is identical. Interestingly, more thorough analysis of the

2 Mechanisms of Protein Evolution in Nature

individual domains for both model structures have recently led to the suggestion that they themselves originated from multiple duplication events traceable back to ﬁve- and six-residue peptides [20]. 2.2.2 βα-barrels Another important structure that presumably arose by tandem duplication is the (βα)8 -barrel. Being the most commonly used structural motif in nature, it is found in approximately 10% of all known enzyme structures [24]. The prototypical (βα)8 barrel consists of eight serial strand-turn-helix-turn repeats, the β-strands forming an interior parallel β-sheet through a continuous hydrogen bonding network, and the helices forming the surface. Studying the symmetric nature of the barrel itself, multiple sequence alignments of the previously introduced gene products of hisA and hisF suggested the presence of an internal repetition within the fold [17]. Based on this data, the authors proposed that the (βα)8 -domain arose by tandem duplication of a (βα)4 -unit (Fig. ??). The validity of such a gene duplication hypothesis was recently examined from a structural perspective [18] and experimentally on HisF from Thermotoga maritima [25]. After bisection of the enzyme into two symmetrical (βα)4 -domains and expression of these in E. coli, the two subunits folded independently into their correct conformations. Compelling evidence in support of the hypothesis was found when coexpression of the two (βα)4 -units resulted in self-association into a functional heterodimer with activities similar to wild type. Although speculative, it has been suggest that the (βα)4 -barrel itself may have originated from even smaller elements such as individual (βα)-units ([26] and references therein). Experimentally, the rearrangement and substitution of (βα)-units in the context of the barrel structure are tolerated [27, 28].

Fig. 2 Schematic illustration of the ($")8 barrel structure. The highly symmetrical structure presumably arose by multiple gene duplication events from a $"-fragment. This hypothesis was tested using reverse engi-

neering. Bisection of the barrel into two halves yielded correctly folded fragments which, upon heterodimerization, regained parental function. However, further fragmentation awaits experimental exploration.

5

6 S. Dedeblatt

2.3 Circular permutation

Circular permutation of a protein results in the relocation of its N- and C-termini within the existing structural framework. Initiated by a tandem duplication of a precursor gene, one mechanistic model proposes an in-frame fusion of the original termini, followed by the generation of a new start codon in the ﬁrst repeat and a termination site in the second. In support of the model, tandem duplications are observed in prosaposins [29] and DNA methyltransferases [30], both genes for which circular permutated variants are also known. Initial evidence for the structural diversiﬁcation of the βα-barrel fold by circular permutation was found in sequence alignments of members of the α-amylase super-family [31]. More recently, analysis of crystallographic data of transaldolase B from E. coli suggested that the enzyme was derived from circular permutation of a class I aldolase [32]. In either case, the shift of the two N-terminal βα-repeats (plus the β-strand of the third subunit for amylases) onto the C-terminus resulted in no apparent functional changes. In general, the evolutionary advantages of circular permutation are not clear. However, in vitro experiments have revealed that although the ﬁnal structure of a circular permuted protein is by and large the same, the rearrangement of the subunits changes the folding pathway and can affect the protein’s conformational stability [33]. The potential beneﬁt of such structural rearrangements from an engineering perspective was demonstrated by the increased anti-tumor activity of a circular permuted interleukin 4-Pseudomonas exotoxin fusion construct, resulting from decreased steric interference between the toxin and the interleukin-binding site [34]. 2.4 Oligomerization

Noncovalent association of homo- and heterologous proteins plays an important role in enzyme function [35]. Oligomerization can modulate a protein’s activity as demonstrated in allosteric enzyme regulation of hemoglobins [36] or by the dimer activation of transcription factors [37]. In addition, the agglomeration of structures at exposed hydrophobic surfaces increases the thermodynamic stability of the participating elements and can create cavities and grooves at the protein-protein interface that can accommodate substrate or regulator binding sites. Along the same line, oligomerization can be viewed as a primitive way of substrate shielding in an enzyme’s active site. A special case of oligomerization is the formation of homodimers by 3D-domain swapping, a term easily confused with “domain swapping”. As shown in Fig. ??, the latter is a synonym for domain recruitment and describes the engineering of proteins by substitution of either gene fragments that encode for individual domains or, in the case of inteins, discrete protein fragments. This involves the breakage and reformation of covalent bonds in the peptide backbone. In contrast, 3D-domain swapping describes the structural reorganization between identical

2 Mechanisms of Protein Evolution in Nature

Fig. 3 Graphical distinction between domain swapping/recruitment and 3D-domain swapping.

multi-domain proteins without breakage of covalent bonds. In the process, a portion of one protein is swapped with the same unit of an identical protein, forming an intertwined dimer or higher order oligomers. Several examples of naturally occurring 3D-domain swapping have been reviewed by Schlunegger et al. [38]. The exchanged structure elements range from secondary structures to entire domains and, although the advantages of 3D-domain swapping are not fully understood, one can envision regulatory purposes as well as increased structural stability [39].

2.5 Gene fusion

Fusion of entire genes, which translates into covalently linked multifunctional enzyme complexes, is a more conclusive commitment towards cooperativity between individual enzymes than non-covalent oligomerization. Often found for enzymes that catalyze subsequent steps in biosynthetic pathways, gene fusions can provide stability, advanced regulatory function, substrate channeling, the ability to direct concerted expression, and entropic advantages, giving the host organism a selective advantage. Multifunctional gene fusion constructs in both the pyrimidine and purine biosynthetic pathways have been discussed in detail elsewhere [40]. Additional examples have been identiﬁed in the biosynthetic pathways for histidine [16] and the aromatic amino acids. In the latter, chorismic acid is the key intermediate from which tryptophan, as well as tyrosine and phenylalanine biosynthesis diverges. In E. coli, two bifunctional complexes catalyze the conversion of chorismic acid via the prephenate intermediate into the respective α-ketoacid precursor of tyrosine and phenylalanine. While the tyrosine precursor is synthesized by a chorismate mutase (CM) fused to prephenate dehydrogenase (CM-PDG), phenylpyruvate is produced by a CM-prephenate dehydratase complex (CM-PDT). Either complex is reportedly susceptible to feedback inhibition by its ﬁnal product [41, 42]. After bisection of the fusion construct, the individual monomers remain functional, excluding the necessity of gene fusion for the purpose of substrate channeling. Instead, allosteric control seems the driving force behind dimer formation [41, 43]. These

7

8 S. Dedeblatt

ﬁndings agree with experimental data from the bisection of N-acetylglucosamino1-phosphate uridyltransferase, an enzyme complex that catalyzes the consecutive acetylation and uridylation of glucosamino-1-phosphate. While both enzymes are essential in vivo, their cleavage shows no apparent effect on kinetics of the individual enzymes, the pathway’s overall ﬂux, nor the organism’s phenotype [44]. In contrast, enhanced substrate processing was observed for rationally designed β-galactosidase – galactose dehydrogenase and galactose dehydrogenase-bacterial luciferase fusion constructs [45, 46]. 2.6 Domain recruitment

The generation of functional versatility in proteins by reorganization of individual protein subunits or domains is summarized under the term “domain recruitment”. The idea behind domain recruitment has been vividly depicted by Ostermeier and Benkovic’s adaptation of the clich´e “a thousand monkeys typing at a thousand typewriters would eventually reproduce the works of Shakespeare” [40]. In reference to domain recruitment, these authors suggest that the writing would obviously work much more efﬁciently if the monkeys, once managing to write entire words or sentences, could duplicate these with a single keystroke. Evidence for domain recruitment has been identiﬁed in a wide variety of proteins [47], mechanistically ranging from simple N- or C-terminal fusion to multiple internal insertions and possibly circular permutations [48]. A recent analysis of proteins in the protein structure database (PDB) has further indicated that structural rearrangements as a result of domain shufﬂing have signiﬁcantly contributed to today’s functional diversity [49]. A brief overview of the various modes of domain recruitment and their effects on function, is presented on examples of βα-barrel structures. Substrate speciﬁcity The family of glycosyl hydrolases utilizes a variety of oligo- and polysaccharides. Preserved across the family, the active site is located in a wide-open groove at the C-terminus of the βα-barrel core structure. Substrate speciﬁcity between the family members is modulated through additional domains, which effectively close down the active site, adjusting its size and shape towards the individual substrates. In lacZ from E. coli, four “added” domains were identiﬁed, orchestrated around the central βα-barrel [50]. These add-ons all represent individual folding units with distinct structures; jelly-roll barrel (domain 1), ﬁbronectin III fold (domain 2 & 4), and β-sandwich (domain 5). Functionally, domain 1 was found to directly participate in shaping the active site pocket while domain 2 & 4 act as domain linker to correctly position domain 1 and 5. Although the role of domain 5 is less clear, it seems to contribute to substrate binding and may be relevant to oligomerization. In contrast to the above examples, substrate speciﬁcity can also be acquired by deleterious modiﬁcation of the protein framework. The absence of two consecutive α-helices in the barrel structure of endo-β-N-acetylglucosaminidase F1 (Endo F1 )

2 Mechanisms of Protein Evolution in Nature

was ﬁrst reported by Van Roey et al. [51]. While not affecting the structural or functional integrity of the protein, the unusual feature was interpreted as important to binding of the bulky glycoprotein substrates and their subsequent deglycosylation. Similar alterations of the βα-barrel core structure, although to a lesser extent, were observed in family relatives of Endo F1 and could also be attributed to substrate binding [52].

Regulation The catalytic activity of porcine pancreatic α-amylase, another member of the glucosyl hydrolase family, is metal cofactor-dependent [53]. A calcium ion-binding site is located at the interface of an antiparallel β-sheet, inserted in one of the loop regions of the βα-barrel, and the core structure. In the calcium-bound state, the insertion stabilizes the substrate-binding site, and indirectly constrains part of the active site in a catalytically competent conformation. In phosphoenolpyruvate carboxylase, structural analysis indicates the arrangement of the total 40 helices around the active site in the βα-barrel core structure [54]. The additional domains create a binding site for aspartate, an allosteric inhibitor of the enzyme. Furthermore, the highly conserved hydrophobic C-terminal extension of the enzyme presumably is critical for oligomerization.

Chemistry Flavocytochrome b2 from Saccharomyces cerevisiae, a member of the FMNdependent oxidoreductase superfamily, catalyzes the two-electron oxidation of lactate to pyruvate with subsequent electron-transfer to cytochrome c via the bound ﬂavin [55]. What distinguishes the enzyme from other family members is the N-terminal fusion of a heme-binding domain to the βα-barrel structure, which hosts the primary active site. Rather than dumping the electrons from the reduced ﬂavin hydroquinone onto molecular oxygen, they are transferred intramolecularly to the heme-binding domain and from there in a second intermolecular step to cytochrome c.

2.7 Exon shufﬂing

Structure analysis of several proteases involved in blood coagulation and ﬁbrinolysis reveals a diverse, sometimes repetitive, assembly of discrete protein modules (Fig. ??) [56]. While these modules represent independent structural units with individual folding pathways, their concerted action contributes to function and speciﬁcity in the ﬁnal protein product. On the genetic level, these individual modules are encoded in separate exons. Over the course of modular protein evolution, new genes are created by duplication, deletion, and rearrangement of these exons. Mechanistically, the exon shufﬂing actually takes place in the intervening intron sequences (intronic recombination – for further details see [10]).

9

10 S. Dedeblatt

Fig. 4 Schematic illustration of modular protein evolution, exempliﬁed on proteases, involved in blood coagulation and ﬁbrinolysis. Varying type and number of modules, fused onto a common protease unit, generates a family of highly speciﬁc hydrolytic enzymes. (Adapted from [56])

The signiﬁcance of exon shufﬂing to protein evolution, in particular in respect to the development of multicellularity, is signiﬁed by a short inventory of processes involving proteins created by modular assembly. Exon shufﬂing facilitates the construction of proteins involved in regulation of blood coagulation, ﬁbrinolysis, and complement activation, plus most constituents of the extracellular matrix, cell adhesion proteins, and receptor proteins [10, 57].

3 Engineering Genes and Gene Fragments

In recent years, the protein engineering ﬁeld has developed a series of in vitro methods that mimic natural processes in an attempt to improve existing enzymes and develop novel functions and properties. Particularly attractive in that respect have been modular approaches reminiscent of domain recruitment, gene fusion, and exon shufﬂing. By identifying and characterizing these individual modules or building blocks, protein dissection through fragmentation has substantially contributed to the fundamental understanding of protein design. Combined with rational and

3 Engineering Genes and Gene Fragments

combinatorial methods to facilitate the recombination of these building blocks, the exploration of structural and functional properties of proteins has not only produced many improved biocatalysts of industrial importance but also has provided insight to protein function and evolution. The following sections introduce a variety of protein engineering techniques and discuss their applications. 3.1 Protein fragmentation

Methods for protein fragmentation were initially developed to study protein folding mechanisms and pathways [58, 59]. Since, their application has been extended towards the elucidation of structural organization in proteins and their evolutionary origins, as well as towards the design of novel two-hybrid systems. The analysis of a protein’s structural organization by protein fragment complementation facilitates the identiﬁcation of locations in the peptide backbone, permissible for cleavage without loss in function. Knowledge of such sites is valuable since protein engineering frequently requires the substitution and insertion of modules such as secondary structure elements and domains into an existing protein scaffold. Intuitively, one would assume that the manipulation of a scaffold in regions of lower organization such as loops and linkers is better tolerated by the system. Indeed, the analysis of laboratory data and the Protein Data Bank indicated that the majority of insertions and deletions are actually located in loops [60, 61]. Nevertheless some experimental systems have demonstrated surprising ﬂexibility in the accommodation of insertion in deﬁned structural elements [62] and tolerance towards bisection of the peptide backbone in functionally critical regions [63–65]. Experimentally, the search for permissive sites by protein fragmentation can be attempted by various methods. In the absence of structural data, random fragmentation patterns of proteins can be generated by limited proteolysis. Although biased towards surface loop regions, the partial digestion by chemicals, as well as serine and cysteine proteases has been applied to a variety of proteins [66]. More recently, combinatorial approaches that enable a comprehensive mapping of the entire protein structure were described in the literature. Manoli and Bailey reported the transposon-mediated random insertion of a 31 amino acid fragment into a protein scaffold [67]. Alternatively, a protein scaffold can be sampled for permissive sites by circular permutation [68, 69]. Finally, N- and C-terminal truncation of overlapping DNA regions, using Bal31 exonuclease [70, 71] or exonuclease III [72] can be used to search for bisection points. Furthermore, protein fragmentation has successfully been employed to investigate the evolutionary origin of protein structures and folds. Alternatively to evolutionary studies that focus on the formation of complex protein structures, the disassembly of an existing structural framework may provide valuable insight into the nature of evolutionary signiﬁcant building blocks (Fig. ??). Termed reverse engineering of proteins, the method calls for the fragmentation of an existing monomeric protein into pieces that ideally possess distinct structural properties, meaning that they fold independently, and exhibit afﬁnity towards oligomerization.

11

12 S. Dedeblatt

Probably one of the most extensively studied folds in that respect is the (βα)8 -barrel. Earlier introduced in the context of tandem duplication, protein fragmentation experiments have provided data in support of a (βα)4 -domain precursor [25]. More extensive digestion experiments of the (βα)8 -barrel were reported by Vogel et al. [73] and Ray et al. [66]. In both cases, digestion of triosephosphate isomerases with subtilisin resulted in multiple fragments which maintained proper folding and could be religated by the protease under nonaqueous conditions, reconstituting the catalytic activity. While providing valuable insight towards a better understanding of protein evolution, the bisection of proteins into functional heterodimers has also found practical applications to study protein-protein interactions. Heterodimeric variants of dihydrofolate reductase (DHFR), green ﬂuorescent protein (GFP), GAR transformylase, and phosphotransferases were constructed to work in two-hybrid systems [64]. 3.2 Rational swapping of secondary structure elements and domains

Rational protein engineering aims to exchange and fuse secondary structure elements and domains at precise locations, based on predictions through rational means, to form a functional hybrid structure. Despite the great complexity involved in rational protein engineering, two outstanding experimental successes were recently reported for β-barrel and β/α-barrel structures. Engineering members of the serine protease family which structurally consist of two homologous, stacked β-barrels, Hopfner and coworkers designed a hybrid protease by fusing the N-terminal β-barrel from trypsin to the C-terminal β-barrel of factor Xa [74]. The location of the fusion point in the linker region between the two β-barrels was chosen following close inspection of the X-ray structures of trypsin and factor Xa. The resulting hybrid was shown to be fully functional as it hydrolyzed a broader but distinct spectrum of peptide substrates in comparison to the parental enzymes. Using rational protein engineering in combination with DNA shufﬂing on the βα-barrel scaffold, Altamirano et al. [75] demonstrated the successful conversion of indole-3-glycerol phosphate synthase (IGPS) to N-(5 -phosphoribosyl)anthranilate isomerase (PRAI). The two enzymes, catalyzing consecutive reactions in the tryptophan biosynthetic pathway, are thought to have evolved by gene duplication of a common ancestor. Although they share binding afﬁnity towards a common intermediate, their sequence identity (22%) and chemistry is signiﬁcantly different and no cross-reactivity is detectable. Based on initial modeling studies of the structural differences between the two enzymes, the authors deleted, substituted, and inserted gene fragments corresponding to secondary structure elements into the IGPS scaffold. While the resulting rational hybrid enzymes were virtually inactive, two subsequent rounds of ﬁne-tuning” the structural arrangements by DNA shufﬂing resulted in hybrid enzymes with 90% sequence identity to IGPS, but PRAI-activity ten-fold higher than wild-type PRAI. The work is truly remarkable since neither of the two approaches alone could have generated the ﬁnal

3 Engineering Genes and Gene Fragments

hybrid protein. Instead, the authors successfully combined two methods into a new design strategy, rationally rough-drafting the protein scaffold and combinatorially ﬁne-tuning it. Generally, the rational construction and reorganization of hybrid enzymes relies on a comprehensive knowledge of structure and function of the proteins involved. The availability of such information for only a small percentage of proteins has so far limited the effective rational design on a wider range of targets. 3.3 Combinatorial gene fragment shufﬂing

Alternatively to rational design, combinatorial approaches resembling Darwinian protein evolution through random mutagenesis, recombination, and selection for the desired properties can be employed for protein engineering. These approaches do not require extensive structural and biochemical knowledge and their implementation has resulted in successful modiﬁcations and improvements of a wide diversity of targeted properties. In the last decade, a rapidly increasing number of genetic methods for generating combinatorial libraries of proteins have been reported. The early in vivo systems, using chemical and UV-induced random mutagenesis and E. coli mutator strains, have largely been replaced by in vitro methods. With the introduction of errorprone PCR [76], the ﬁrst technique for rapidly generating large families of related sequences of a speciﬁc target gene became available. The next milestone was the successful demonstration of in vitro evolution of proteins by sexual PCR or DNA shufﬂing [77, 78]. DNA shufﬂing The directed evolution of proteins by random mutagenesis and DNA shufﬂing has proven extremely powerful to modify and optimize functional and physicochemical properties of existing proteins [6, 79–85] (for an up-dated list on targets see: www.che.caltech.edu/groups/fha/Enzyme/directed.html). Experimentally, the method starts off with the random fragmentation of a target gene(s) by DNasel treatment. The resulting mixture of double-stranded oligonucleotides, on average 20–100 basepairs in length, is then taken through several rounds of heat-denaturing and reannealing in the presence of a DNA polymerase. This iterative process generates a library of polynucleotides, consisting of fragments from various parental sequences. Today, multiple variations of Stemmer’s original protocol have been reported in the literature [86–90]. Despite the many successful applications of DNA shufﬂing, the methodology has one serious limitation. Sequence-dependent reassembly, the fundamental mechanism behind DNA shufﬂing, only works efﬁciently between sequences with >70 % sequence identity [91]. Below this threshold, DNA shufﬂing generates libraries biased towards regions of locally higher identity and reassembly of parental sequences becomes a dominant factor.

13

14 S. Dedeblatt

Fig. 5 Principle of family DNA shufﬂing, demonstrated on the identity network of three sequences. The thickness of the arrows corresponds to the pairwise sequence identities. The similarity of sequence A with C is low, disqualifying them from regu-

lar DNA shufﬂing. Addition of B acts as a bridging intermediate, having sufﬁcient identity with either A or C to accommodate successful recombination of fragments from all three parents.

Family DNA shufﬂing or “molecular breeding” of multiple parental sequences lowers the approach’s boundaries of sequence-dependence [82, 83, 92]. In contrast to traditional DNA shufﬂing, this strategy is based on homologous recombination of multiple parental sequences, some with pairwise DNA sequence identities as low as 50%. The fundamental idea behind the approach is outlined in Fig. ?? on three sequences. Each member of the initial gene pool has a sufﬁciently high sequence identity to at least one other member. The homology between two speciﬁc sequences may be low, yet the remaining members of the pool, each of them with higher identities to either of the two, act as bridging elements and allocate the shufﬂing of elements from all parents. The experimental implementation of family DNA shufﬂing on the four amp C lactamases from Citrobacter freudii, Klebsiella pneumoniae, Enterobacter cloacea, and Yersinia enterocolitica was highly successful, resulting in the identiﬁcation of hybrid proteins with up to 540-fold increased activity [82]. Interestingly, a detailed sequence analysis of the functional hybrids revealed the absence of amp C fragments from the Yersinia enterocolitica. Contrary to the argument that its presence may not be required under the applied selection conditions, a recent computational analysis of the system suggested that a lack of homology in critical regions of amp C from Yersinia prevented the sequence from participating in the shufﬂing in the ﬁrst place [93]. This alternative explanation highlights the importance of careful design of family DNA shufﬂing experiments with particular emphasis not just on global but also local DNA sequence homologies of all participants. In recent years, a series of alternative methods to shufﬂe sequences of lowhomology has been proposed. Kikuchi et al. [89] took advantage of multiple restriction sites distributed along the gene sequence, which in combination with DNA shufﬂing added crossovers independent of sequence identity to these region. Others suggested the spiking of the DNase-treated fragment mixture with artiﬁcial, designed intermediates; chemically synthesized oligonucleotides that introduce mutations or crossovers at speciﬁc location in a sequence [94]. Homology-independent fragment swapping The limitations of DNA shufﬂing, arising from its dependency on sequence identity, have motivated the development of new technologies to facilitate an unbiased sampling of sequence space. At present, the reported approaches for creating

3 Engineering Genes and Gene Fragments

sequence-independent hybrid libraries between two parental sequences include a family of methods collectively known as Incremental Truncation for the Creation of Hybrid enzYmes (ITCHY) [72, 95, 96] and Sequence-Homology Independent Protein RECom-bination (SHIPREC) [91]. Although the ﬁnal products of all methods are the same, comprehensive singlecrossover hybrid libraries that includes insertion and deletion sequences, the approaches employ different strategies of library generation (Fig. ??). The original ITCHY method [72] involves separate exonuclease III-treatment of the two parental genes (Fig. ??). The reaction conditions are chosen whereby the nuclease has a sufﬁciently low processivity (∼10 nucleotides/min), allowing the generation of libraries of every single-base pair deletion by quenching of aliquots over time. Following removal of the single-stranded portion of the DNA fragments, the N-terminal and the C-terminal fragment libraries are mixed and ligated to N

C C

N

a N

b

C

C N

N

C

C

N

C N C N C N N

C N

C

N N

C

N

N

C C C C N

N

C N

C N

C N

C N

C N

C N

C

parental size

Fig. 6 Fundamental mechanisms of homology-independent fragment swapping by a) ITCHY and b) SHIPREC. In the ITCHY protocol, the two parental genes are truncated independently, creating two separate libraries that correspond to the N- and Cterminal fragments. Ligation of the mixed

libraries then generates a comprehensive pool of hybrid sequences. SHIPREC on the other hand starts by limited DNasel digestion of the fused genes. Intramolecular fusion, followed by restriction digest at the original fusion point, results in sequence inversion and generates the hybrid library.

15

16 S. Dedeblatt

generate the single-crossover library. Following the same fundamental idea but being experimentally signiﬁcantly faster and less laborious is the use of nucleotide analogs such as α-phosphothioate nucleotides to create ITCHY libraries [96]. Furthermore, a variation of ITCHY to bias truncation libraries towards parental sizes has been developed [95]. SHIPREC, an alternative approach to create sequence-independent singlecrossover libraries has been reported by Arnold and coworkers [91]. Utilizing an inverted fusion construct of the parental genes, partial digestion by DNase I cleaves N- and C-terminal fragments at random. Following a size selection of parental size fragments, the original termini are restored by sequence inversion (Fig. ??b). The potential value of these techniques becomes apparent upon analysis of structure diversity and functional selection of ITCHY and SHIPREC libraries. For the model system of glycinamide ribonucleotide transformylase from E. coli (PurN) and human (hGART), comparison of the diversity of the naive libraries obtained by ITCHY with DNA shufﬂing show the expected even distribution of crossovers over the sampled sequence for the former but a clustering in the central, highly homologous region for the latter [72]. Functional analysis of the ITCHY and SHIPREC libraries proves that interesting and catalytically active solutions for protein engineering can be found outside regions of high sequence identity. The main drawback of these described methods is their limitation to a single crossover per hybrid sequence. In contrast, DNA shufﬂing and related methods have produced up to 14 intersects per nucleotide sequence [90]. Increasing numbers of crossovers are desirable because they have been shown to improve the evolution of proteins with desirable characteristics. 3.3.1 Homology-independent fragment shufﬂing The creation of hybrid libraries with multiple sequence-independent crossovers through a combination of ITCHY and DNA shufﬂing (Fig. ??) was ﬁrst proposed by Ostermeier et al. [97] and has recently been implemented experimentally [98]. Using the PurN/hGART model system, two complementary incremental truncation libraries (purN-hGART and hGART-purN) were generated (Fig. ??). In preparation for the subsequent DNA shufﬂing, these libraries were then pre-selected for in-frame and parental-size hybrid sequences (Fig. ??). Without in-frame selection, the probability of SCRATCHY library members encoding for nonsense sequences would increase exponentionally with the number of introduced crossovers. Furthermore, hybrid enzymes with large insertions are prone to protein folding problems while deletions of extended portions of a protein are likely to terminate enzymatic activity. The resulting libraries resemble large artiﬁcial “families” of intermediates, each carrying a particular crossover between the two parents. DNase I treatment of these families and subsequent reassembly by PCR results in the creation of naive hybrid libraries comprising crossovers from multiple “family” members (Fig. ??c). In experiments, analysis of the naive libraries revealed about 20 % of the members to have two and three crossovers. Their locations were randomly distributed over the entire expected sequence range and functional complementation of an auxotrophic

3 Engineering Genes and Gene Fragments N

C C

N

N

N

C

N N

C

N

N

C

N

C

N

C

N

a

C

C

N

C C

C

N

C N

N

C N

C N

C N

C N

C

C N

parental size

C

parental size

b N

C

N

C

N

C

N

C

N

C

N

C

N

C

N

C

c N C

C

N

N

C

N

C

N

C

N

C

Fig. 7 Homology-independent fragment shufﬂing (SCRATCHY) is a combination of ITCHY and DNA shufﬂing, a) The approach starts by creating two complementary ITCHY libraries. b) To maximize the functional competence of the SCRATCHY library, hybrid sequences of approximately

parental size that are in the correct reading frame are isolated, c) Finally the two ITCHY libraries are mixed, fragmented, and reassembled. The signiﬁcant portion of the resulting hybrid libraries consist of members with multiple crossovers at locations independent of sequence identity.

17

18 S. Dedeblatt

E. coli strain provided evidence for the catalytic competence of multi-crossover hybrid constructs. In respect to catalytic performance of these hybrid enzymes, the method lags behind DNA shufﬂing approaches. This however can mostly be attributed to issues of protein folding rather than chemistry itself. In the following section, the aspects of protein folding and its increasing importance to protein engineering at low sequence identities will be discussed. 3.4 Modular recombination and protein folding

The biological activity of proteins generally depends on a unique three-dimensional conformation, which in turn is inherently linked to its primary sequence. Protein folding, the conversion of the translated polypeptide chain into the native state of a protein, is the critical link between gene sequence and three-dimensional structure. Mechanistically, folding is believed to proceed through a predetermined and ordered pathway, either via kinetic intermediates or by direct transition from the unfolded to the native state [99]. In both cases, local and non-local interactions alike stabilize transient structures along the pathway and funnel the intermediates towards the native state. In the context of protein engineering, these folding and stability issues are of major importance. Modular protein engineering in general and homologyindependent methods in particular can introduce signiﬁcant variation into a system’s primary sequence, impacting the network of interactions that ultimately controls the folding trajectory. Proteins have generally demonstrated amazing ﬂexibility and tolerance towards changes of individual amino acids and small peptide fragments, accommodating them by reorganization of the neighboring environment [62]. In the context of domain swapping and modular engineering, the situation is often more complex. Affecting mostly non-local interactions, this remodeling can disturb long-range interactions crucial to folding and structural integrity. Furthermore, possible steric and physicochemical interferences are expanded to entire surfaces of a protein module. Although the disruption of a network of stabilizing interactions between parental structures is possible on all levels of homology, the probability for structural inconsistencies that lead to misfolding rapidly increases with decreasing sequence similarity. When exploring distant regions in sequence space by homology-independent methods as discussed earlier, deleterious folding effects on hybrid protein libraries upon exchange of subunits of low homology can quickly nullify the potential beneﬁts of structure-based protein engineering. The problem of incorrect folding of hybrid proteins is illustrated by a system investigated in our laboratory. Attempting the substitution of domains from structures with very low sequence homology, incremental truncation libraries between the E. coli proteins GAR transformylase (PurN) and tRNA(fMet)-formyltransferase (FMT) were generated (Marc Ostermeier and Steve Tizio – unpublished data). The N-terminal portions of the two proteins share only 33 % sequence identity, yet their backbone tertiary structures are almost perfectly superimposable (Fig. ??). In addition, they both catalyze the same chemistries; the formyl group-transfer

3 Engineering Genes and Gene Fragments

Fig. 8 a) Structural overlay of glycinamide ribonucleotide transformylase (PurN) [155] and the N-terminal domain of tRNA(fMet)formyltransferase (FMT) [156] from E. coli. Despite the low sequence identity (33 %), the two structures are almost perfectly superimposable. b) Structure-based multiple sequence alignment of PurN and FMT with

human glycinamide ribonucleotide transformylase (hGART) and formyltetrahydrofolate hydrolase (PurU). Conserved residues are labeled with an asterisk and homologous positions are marked with a dot. The three active site residues are highlighted in shaded boxes.

19

20 S. Dedeblatt

from N10 -formyl-tetrahydrofolate onto their respective substrates. In previous experiments, analogous substitution of fragments between the functionally related PurN and N10 -formyl-tetrahydrofolate hydrolase (PurU) from E. coli (60 % DNA sequence identity), as well as PurN and hGART (50% DNA sequence identity) had yielded functional hybrid enzymes [63, 100]. A potential indicator for protein folding problems was the identiﬁcation of several thermosensitive constructs of PurN/hGART, which failed to complement in an auxotrophic host under regular growth conditions at 37◦ C, yet showed sufﬁcient activity when incubated at ambient temperatures [95, 96]. Presumably, the perturbance of the network of non-local interactions at or near the fusion constructs intraface destabilizes the global structure and prevents proper folding at higher temperatures. In contrast, expression of the PurN/FMT hybrid library at either temperature did not yield any functional chimeras. While subsequent analysis of the libraries on the genetic level indicated the expected diversity, Western blots revealed that the expressed protein was found exclusively in the insoluble fraction of the cells (Steve Tizio – unpublished data). The same difﬁculties were reported for a series of rationally designed hybrid constructs between ornithine and aspartate transcarbamoylases, two structure homologs that share only 32 % sequence identity [101]. We hypothesize that the observed instability of hybrid constructs is caused by two mechanisms: i) steric and functional clashes at the module surfaces and ii) disruption of folding pathways. For the former, interference at the module’s intraface is illustrated by the “English mufﬁn-experiment”. Tearing two mufﬁns in half and trying to ﬁt the bottom of the ﬁrst and the top of the second back together is not likely to match up precisely. Instead, the contact surface will have friction points and empty spaces. A similar picture can be drawn for the two fused protein fragments, causing the observed destabilization of the constructs. Furthermore, dissimilarities of module surfaces can result in the exposure of hydrophobic patches, increasing the tendency to agglomerate or misfold. While future studies will have to address these issues in detail, a practical solution to circumvent the problems may be site-directed or random mutagenesis. Mutagenesis of only few positions signiﬁcantly improved protein solubility and as a result functionality in a variety of systems [102–106]. While the rationale for most changes is unclear, the majority of mutations were located on the protein surface and involved the substitution of hydrophobic amino acids with charged constituents, minimizing hydrophobic patches that would otherwise be prone to aggregation. Furthermore, fundamental differences in the folding pathway between the two parental sequences could affect proper structural alignment in a chimera. This hypothesis is supported by recent results from two studies of evolutionary distant structural homologs. The ﬁrst study focused on two members of the globin family, soybean leghemoglobin and myoglobin, two proteins with similar threedimensional structure but only 13 % sequence identity [107]. Extensive analysis and comparison of the folding process revealed overall similar, compact helical folding intermediates. These intermediates however differ signiﬁcantly in detail, consisting of helix A, G, H, and parts of helix B for myoglobin but helix E, G, and H for leghemoglobin. In the second study, the folding mechanisms of two

3 Engineering Genes and Gene Fragments

proteins in the family of intracellular lipid binding proteins were investigated [108]. As in the previous report, the two proteins share very similar structure yet their sequences have diverged to only 23 % sequence identity. The spectroscopic analysis of the folding intermediates indicates very distinct folding pathways leading to the same ﬁnal structures. Upon substitution of individual parts of two such structures, it seems reasonable to assume that neither of the two original folding pathways will be suitable to orchestrate the conformational organization of the structure as a whole. Instead, the system will likely follow a third, unknown trajectory with uncertain outcome. From an experimental standpoint, the issue can be addressed from two sides. As discussed earlier, mutagenesis can provide the means to make the necessary adjustments to restore solubility and functionality. In a recent report, protein-folding problems due to a C-terminal truncation in chloramphenicol acetyltransferase (CAT) could be corrected by the introduction of additional secondary site mutations [109]. Random mutagenesis of a deleterious, insoluble CAT variant followed by selection for chloramphenicol resistance in E. coli yielded a functional mutant carrying a single point mutation. The beneﬁcial effect of the conservative leucine to phenylalanine substitution, located in the hydrophobic core of the protein, clearly could be attributed to an increase in folding efﬁciency. An alternative concept can be the fusion of domains in the context of independent folding units, exempliﬁed by the modular assembly of proteases involved in blood coagulation and ﬁbrinolysis. Modules are commonly associated with domains and, on a genetic level, with exons [110–112] although the correlation is not dominant [113]. Recent work with zinc ﬁnger binding domains in conjunction with nucleases or regulatory elements demonstrates the successful design of hybrids enzymes based on modular engineering.

3.5 Rational domain assembly – engineering zinc ﬁngers

A very popular target for exploring and exploiting the modular design of proteins is the zinc ﬁnger family. Zinc ﬁngers consist of a ββα domain, stabilized by a zinc metal ion with tetrahedral planar coordination to two cysteine and two histidine side-chains. Interacting with the major groove in DNA, the zinc ﬁnger domain speciﬁcally recognizes three-nucleotide stretches. Serial assembly of individual ﬁnger domains can therefore create protein complexes with an extended DNA recognition pattern, a highly desirable property in the post genomic area. Repeatedly, studies have indicated that fusion of ﬁnger modules through their natural linker sequence is, however, limited to three domains. Each insertion of a zinc ﬁnger into the major groove of DNA causes a slight bending of the latter, putting any module past the third domain out of reach for its DNA recognition sequence. To speciﬁcally target a unique site in even a relatively small genome such as E. coli, the 9 nucleotide-recognition pattern of the triple ﬁnger is not sufﬁcient. Furthermore, the repertoire of natural zinc ﬁnger recognition sequences, the diversity of the “codons”, is limited to a few combinations, leaving only few sites

21

22 S. Dedeblatt

that comprise the correct sequence and length. Recent advances in the ﬁeld have addressed and provided insight to these questions and are discussed below. Focusing on the limitations of codon diversity, random mutagenesis of existing zinc ﬁnger domains followed by screening for variations in the recognition sequence by phage display technology has led to the identiﬁcation of domains recognizing all 16 possible 5 -GNN-3 combinations [114]. The ability to speciﬁcally target desired sites in vivo, using the expanded set of ﬁnger domains, was demonstrated. Fusing either three or six modules (two three-ﬁnger constructs, linked by a pentapeptide) to transcription factor domains, the expression of a luciferase reporter gene could be up or down-regulated [115]. An alternative approach to overcome the limitations of triplet diversity has been reported by Klug and coworkers [116, 117]. By introducing structured linkers between the individual binding domains, stretches of up to 10 nucleotides could be bridged without compromising binding afﬁnities. Rather than trying to overcome the limitation of the number of naturally occurring codons in zinc ﬁngers, their tactic is based on extending the length of the target sequence that can be covered by the DNA recognition domains. Having a set of linkers with ﬂexible length allows the custom design of highly speciﬁc binding complexes using only the modules at hand. Combining the two approaches may even further increase the versatility of the zinc ﬁnger domains. Finally, the fusion of such engineered multi-module zinc ﬁnger subunits with a variety of catalytic subunits can provide very powerful tools for the engineering of entire genomes as demonstrated with transcriptional regulators [118] or with endonuclease domains [119].

3.6 Combinatorial domain recombination – exon shufﬂing

Advances in molecular biology and high-throughput screening, as well as the projected ﬂexibility and diversity of modular assembly in natural protein evolution has inspired the development of various techniques to implement combinatorial domain recombination, better known as exon shufﬂing, in vitro. Two early proposals focused on the exploitation of self-splicing introns to catalyze the fusion of RNA libraries. Fisch and coworkers [120] generated large 25-residue peptide libraries by constructing two random 30-base pair exon libraries, ﬂanked either on their 5 - or 3 -end by a group I intron sequence with an inserted loxCre recombination site. Following electroporation of the ﬁrst plasmid-based exon library into E. coli and transformation of the second library through phage infection, in vivo recombination of the two libraries by Cre-recombinase produced a repertoire of ∼1.6 × 1011 members. In a ﬁnal step, the lox-Cre recombination site between the two exon-libraries was excised by the adjacent intron sequences, retaining a 15-base pair stretch at the points of fusion. Screening of the resulting peptide library, comprising two hyper variable 10-amino acid regions with an intervening ﬁve-amino acid residue spacer, by phage display identiﬁed several sequences with up to nanomolar binding afﬁnities against various protein targets.

3 Engineering Genes and Gene Fragments

Simultaneously, Mikheeva and Jarrell [121] proposed in vitro exon shufﬂing directly by engineered trans-splicing group II intron ribozymes. The method was implemented on a model system consisting of two fragments from human tissue plasminogen activator. RNA analogs were generated by separate reverse transcription of the two exons, fused to their corresponding intron sequences. A mixture of the two transcripts was then trans-spliced. Although the ligation efﬁciency was low, gel analysis clearly indicated product formation. In contrast to the previous approach, the engineered ribozymes facilitate the seamless fusion of the two exons. An elegant alternative to the intron-catalyzed recombination has been derived from DNA shufﬂing [122]. For initial experiments on a human single-chain Fv framework, six complementarity-determining regions (CDRs) within the structure (which could be viewed as individual domains in a general sense) were identiﬁed. Fragments of the DNase-treated parental gene were then spiked with synthetic oligonucleotides comprising randomly mutated derivatives of these six CDRs and the mixture was reassembled by PCR. The analysis of the resulting naive library indicated a random mix of wild-type and mutant CDRs, evenly distributed over all six regions. As discussed in a recent review [57], a modiﬁed version of the approach now assembles libraries from mixed pools of homologous exons. The proposed method gives great ﬂexibility to customized libraries based on the design of the chimeric oligonucleotide linkers. Presumably, the size and sequence of the linkers, as well as the concentration of individual exons in each pool and the mixing ratio of the different pools can be used to control the reassembly process (Fig. ??). More hypothetically in the context of exon shufﬂing, post-translational proteinfragment recombination by trans-splicing inteins can be employed to recombine protein modules [123]. Trans-splicing inteins were ﬁrst discovered in Synechocystis sp. PCC6803, catalyzing the speciﬁc and directed religation of the

Fig. 9 Exon shufﬂing: combinatorial libraries of multidomain enzymes can be created through recombination of individual modules or domains (I–IV) from multiple parents (A–D) in an ordered fashion, maintaining the parental size (left diagram) or by random reassembly. This allows he

insertion, deletion, substitution, or duplication of modules in a single step. The black and white hexagons, stars, and circles indicate unique complementary elements (complementary DNA strands or pairs of trans-splicing inteins).

23

24 S. Dedeblatt

Fig. 10 Ligation of two protein fragments by trans-splicing inteins: a) on the DNA level, the two fragments can be encoded in separate ORFs and are ﬂanked by either a 3 -extension encoding the N-intein (Nfragment) or 5 -extension encoding the C-

intein (C-fragment). b) Following transcription and translation, the intein domains dimerize and c) excise themselves, fusing the N- and C-fragment by a peptide backbone bond.

genetically separately encoded protein fragments that make up the replicative DNA polymerase III catalytic α-subunit (DnaE) [124]. Consisting of short protein sequences at the C-terminus of the ﬁrst protein fragment and the N-terminus of the second protein fragment, the inteins, following translation, dimerize in vivo and catalyze the peptide-bond formation between the two protein fragments while simultaneously excising themselves (Fig. ??). Using the same basic approach for the successful random fusion of domains and modules of proteins on a post-translational rather than the genetic level would depend on two fundamental requirements. First, multiple pairs of trans-splicing inteins would have to be available and second, the cross-reactivity between the individual pairs of inteins would have to be either minimal or non-existent. So far, a single pair of naturally occurring trans-splicing inteins has been reported in the literature and an additional four pairs have been engineered from related cis-splicing inteins [125]. Assuming that the number of engineered intein pairs will further increase over time, the ﬁrst requirement should not be a major limitation to the approach. Cross-reactivity between N- and C-termini of trans-splicing inteins from different pairs is the other primary concern. Recently, Otomo et al. [126] reported the simultaneous in vitro recombination of three protein fragments, tagged with two separate pairs of trans-splicing inteins. The ligation, followed by in vitro refolding, generated a correctly assembled, functional full-length protein. Similar experiments in our laboratory, using heterodimeric-GFP, indicate that correct intein-mediated in

4 Gene Fusion – From Bi- to Multifunctional Enzymes 25

vivo assembly of functional protein in the presence of multiple trans-inteins is also possible (C.P. Scott – personal communication).

4 Gene Fusion – From Bi- to Multifunctional Enzymes

While the rationale behind gene consolidation in natural systems is still disputed, protein engineering has made extensive use of the idea of linking genes. Some of the applications may be as simple as attaching a reporter module onto the N- or C-terminus of the target. Alternatively, joining parental genes by insertion of one sequence into the other has created bifunctional hybrids with, in some cases, allosteric properties. Finally, the assembly of multiple functional units into entire protein machineries has been directed towards the multi-step synthesis of complex organic molecules. 4.1 End-to-end gene fusions

Rationally designed end-to-end fusions of genes are employed in the puriﬁcation, detection, selection, and localization of targeted gene products. Probably the widest-known applications of covalently connecting two proteins are for puriﬁcation purposes. Fusion constructs between a target protein and a tagging gene such as the maltose-binding protein, chitin-binding domain, thioredoxin, β-galactosidase, or glutathione S-transferases are frequently employed to aid in puriﬁcation and in some cases actually help stabilize and increase solubility of target proteins [127, 128]. In addition, fusion of a reporter gene such as GFP or bacterial luciferase can be used to identify and localize a protein of interest in vivo and in vitro. Issues of protein solubility and folding can also be addressed by gene fusions in vivo. C-terminal extensions of a target gene with GFP, chloramphenicol acetyltransferase, or β-galactosidase have successfully been applied to select for solubility and in-frame constructs [129–131]. Additional applications of fusion constructs include co-expression with transporter proteins that direct and anchor the target protein to the outer membrane, effectively displaying the protein on the host’s surface. Implemented in phages using the pIII and pVIII coat proteins as fusion partners, as well as in bacterial hosts and yeast using surface-exposed loops or outer-membrane proteins [132–134], these systems are particularly useful for the screening of protein libraries. 4.2 Gene insertions

While most of the gene fusions discussed so far focused on N or C-terminal extensions, the insertion of an entire gene into a second gene is possible and may be advantageous. In a rather intriguing experiment, Betton et al. [135] in-

26 S. Dedeblatt

serted an entire β-lactamase gene (bla, 708 nucleotides) into loop regions of the maltodextrin-binding protein (MalE). Inserting bla at position 130 or 303 in malE which were previously identiﬁed as permissive to smaller insertions and dissection [136–138], the authors were able to isolate fully active bifunctional fusion constructs. In addition, MalE was shown to increase the stability and modulate the catalytic activity of the fused lactamase. Similar experiments were reported, exploring the tolerance of the phosphoglycerate kinase scaffold towards insertion of dihydrofolate reductase or β-lactamase and its effect on folding [139], as well as creating bifunctional fusion proteins between 1,3-1,4-β-glucanase and 1,4-β-xylanase [140]. Further expanding the published protocol, Baird et al. [141] circularly permuted an internal fusion construct between the enhanced yellow ﬂuorescent protein (EYFP) and a zinc ﬁnger domain or calmodulin. Conformational changes upon metal binding at the insertion domain are translated into structural rearrangements of EYFP. These structural changes in turn affect the ﬂuorometric properties of EYFP; making these fusion constructs in vivo biosensors to monitor Zn2+ and Ca2+ concentration in cells. An allosterically regulated, GFP-based biosensor, sensitive to antibiotics, was engineered by inserting β-lactamase into one of the surface loops of GFP followed by random mutagenesis and selection for antibioticdependent changes in the ﬂuorescence of the fusion libraries [142]. 4.3 Modular design in multifunctional enzymes

One of the most exciting areas in the ﬁeld of multifunctional enzyme systems is the synthesis of a wide array of organic molecules by polyketide and nonribosomal protein synthetases. These enzymes are generally characterized by multiple subunits which themselves consist of individual domains with distinct enzymatic activities (Fig. ??a). The range of natural products synthesized by these mega-synthetases includes a considerable number of important antibiotics, antifungals, antitumor and cholesterol-lowering compounds, immunosuppressants, and siderophores. Three categories of synthetases are distinguished, based on their substrate speciﬁcity and mode of product synthesis. The two known types of polyketide synthetases (PKSs) (Type I and II) utilize acyl-coenzyme A (CoA) monomers while nonribosomal peptide synthetases (NRPSs) use amino acids and their analogs as substrates. Type I PKS and NRPS oligomerize these building blocks by a modular assemblyline arrangement while type II PKS iteratively assembles monomeric units. The modular design of PKS and NRPS renders them convenient for protein engineering to generate hybrid enzymes that can synthesize a range of natural and unnatural metabolites. By independently manipulating what Cane et al. [143] call the four degrees of freedom in polyketide synthesis: variation of chain length, choice of ACP domains (substrate), degree of β-carbon modiﬁcation, and stereochemistry, product libraries of extremely high diversity can hypothetically be synthesized. Likewise in NPRSs, the variation of chain length, type and stereochemistry of the substrates, and their level of post-synthetic derivatization can potentially yield

4 Gene Fusion – From Bi- to Multifunctional Enzymes 27

Fig. 11 Organizational overview and summary of protein engineering efforts on the polyketide synthetase (PKS) and nonribosomal peptide synthetase (NRPS) framework, a) In both systems, the complete synthetases consist of multiple subunits encoded on individual genes. The subunits themselves are divided in modules which each catalyze the addition of one acyl-building block (PKS) or amino acid (NRPS). The minimal module for PKSs is made up of three domains – a keto synthase (KS), an acyl transferase (AT), and an acyl carrier protein (ACP). In addition, modules can contain up to three modifying domains to derivatize $-carbons on the

polyketide chain; a keto reductase (KR), a dehydratase (DH), and an enoylreductase (ER). Flanking the entire complex is a loading module (LM) and a termination module (TE). The minimal module for NRPSs consists of an adenylation (A), athiolation (T), and a condensation (C) domain. Additional modiﬁcations on the peptide chain can be introduced by epimerization (E) or N-methylation (NM) domains. The loading module in NRPS consists of an adenylation/thiolation (AT) domain while the synthesis is complete by a thioesterase (TS). b) Protein engineering of PKSs and NRPSs, organized by structural objectives.

an almost unlimited number of compounds with potentially novel or improved properties. Initial efforts to rationally engineer PKS and NRPS have focused on the alteration of the biosynthetic pathway of metabolites by deletion, insertion, and substitution of individual domains (summarized in Fig. ??). Although these experiments provided valuable information about the structure and function of the individual domains and modules, formation of functional hybrid enzymes proved difﬁcult at times. In contrast, taking entire modules and rearranging them in the context of an enzyme complex was shown to be a more successful route towards the creation of functional hybrid enzymes. Of particular interest were data indicating the significance of the linker regions between covalently as well as non-covalently attached modules. By restoring the correct inter- or intra-polypeptide linker sequences, Khosla and coworkers swapped module positions in the polyketide synthetase 6deoxyerythronolide B synthase (DEBS) and introduced modules from other PKSs, generating an array of functional hybrid enzymes [144–146]. Beyond the swapping of whole modules, efforts to prepare libraries of compounds have turned to randomization of the proteins within each module. With

28 S. Dedeblatt

respect to these truly combinatorial engineering efforts, modular designed type I PKSs have so far been most amenable. Shortly after the report of the successful but elaborate generation of a combinatorial library by cassette replacement of domains in DEBS [147], a relatively simple and elegant multi-plasmid approach was described [148]. The three genes of DEBS were cloned in separate plasmid vectors, followed by introduction of mutations to diversify each individual gene. Upon triple transformation into a heterologous host, a small library of related polyketides was produced based on the synthesis by random members of the three vectors. Although these results are very exciting, the method is very much limited by the number of different plasmids that can be present in a single host cell. The efﬁciency of triple transformation is low and plasmid stability may be a major problem. The engineering of iterative (type II) PKSs is considerably more difﬁcult due to their repetitive use of a single module in the biosynthesis of natural products. So far, the only combinatorial libraries of polyketides generated with these types of PKS are created not by genetically encoded diversity but by construction of minimal PKS assemblies. Minimal PKSs are created by removal of all accessory proteins such as cyclases, ketoreductases, and aromatases, reducing the functional enzyme complex to three essential domains: a ketosynthase, a chain length factor, and an acyl carrier protein. Studying the minimal version of whiE (spore pigment) PKS, Moore and coworkers found more than two dozen unique polyketides, generated by a single enzyme multiplex [149]. The diversity was generated by premature product dissociation from the enzyme complex at various stages of the oligomerization, resulting in a variable product chain length and regiospeciﬁcity of the cyclizationevent.Similar variations in the natural product palette of PKS from Streptomyces maritimus suggest that the same mechanismsare employed by Nature to explore potentially beneﬁcial effects of these secondary metabolites [150]. In contrast, Nature’s mechanism of directing biosynthesis towards particular metabolites is accomplished by accessory proteins such as the cyclase in the natural whiE system. By non-covalently binding to the minimal PKS complex, the accessory protein stabilizes the product-enzyme intermediates, ensuring formation of full-length product and correct cyclization. While such a system is valuable for the generation of polyketide libraries, problems with reproducibility of library composition due to the unknown factors that inﬂuence the generation of the diversity may cause problems. Furthermore, upon identiﬁcation of interesting or useful products in the library, no direct approach exists for resynthesis of these particular compounds. Instead, additional rational engineering is required to attempt the construction of a hybrid that exclusively produces the desired compound. Finally, nonribosomal peptide synthetases represent the least explored members in this family of multienzyme complexes. They represent an important and versatile class of megasynthetases, involved in the biosynthesis of a wide spectrum of peptidylic secondary metabolites including penicillins, vancomycins, bleomycin, and cyclosporin A. The biosynthesis of these compounds is achieved by utilizing not only natural amino acid but also a wide range of D-conﬁgured and N-methylated amino acids and hydroxy acids. Variations in the peptide backbone structure result in linear, cyclic, and branched cyclic molecules and these structures can be

5 Conclusions 29

modiﬁed by acylation, heterocyclization, reduction, and glycosylate. Very similar to the modular design of type I PKS, nonribosomal peptide synthetases consist of a linear arrangement of multiple modules linked either covalently or noncovalently (Fig. ??a). The individual modules are responsible for the incorporation of a speciﬁc amino acid in the growing peptide chain, therefore dictating the product sequence. The modules themselves are formed by three domains, an adenylation-domain (A-domain; activates incoming amino acid), a thiolation-domain (T-domain; peptidyl carrier protein), and a condensation-domain (C-domain; facilitates the reaction between peptidyl moiety and activated amino acid). As for PKSs, rational engineering experiments have focused on the exchange of individual domains within a module [151]. More recently, Marahiel and coworkers [152] reported the successful rearrangement and fusion of intact modules, generating novel peptide products based on the amino acid speciﬁcity of the individual building blocks. Combinatorial approaches towards the creation of peptidylic libraries by NPRSs have not yet been reported. Considering the very similar overall design of NPRSs to modular PKSs, one can imagine applying similar methods as discussed above. In addition, the recently described site-directed mutagenesis of the substraterecognizing A-domain of NRPSs, resulting in the alteration or relaxation of its substrate speciﬁcity, may be employed to generate a pool of heterogeneous products [153]. In summary, the structural and functional work on PKSs and NRPSs has provided a basic understanding of secondary metabolite biosynthesis. At the same time, these systems have proven extremely valuable to protein engineering efforts. However, the current methodology is not capable of coping with the demands of a large-scale combinatorial approach to generate polyketide- and peptide libraries. Although all the individual elements are readily available, the limitations of an effective shufﬂing and recombination protocol currently prevents the creation of libraries with >100 individual members.

5 Conclusions

Studies of protein evolution and protein engineering are progressing hand-in-hand. Advances in our understanding of protein evolution have unveiled strategies that have led to the development of novel methods in protein engineering. In return, mimicking natural processes in the laboratory has revealed some of the robust but at the same time delicate nature of complex protein structures, furthering our understanding of the fundamentals in protein evolution. The industrial aspects of protein engineering, as well as the exploration of natural protein evolution, can both proﬁt from one another providing further insight into the fascinating world of proteins. From a methodology standpoint, no single approach provides a universal solution to protein engineering. The nature of the individual target, the objective of the engineering project, and downstream factors such as the availability of

30 S. Dedeblatt

a high-throughput screen or a selection procedure dictates the technique or techniques most suitable to address the problem at hand. While the development of fundamentally novel methods will progress, a particularly attractive approach has become the combination of individual techniques as demonstrated by Fersht and coworkers [75] using rational design and DNA shufﬂing, as well as by our laboratory [98] using ITCHY and DNA shufﬂing. Such pairwise approaches not only overcome the limitations of the individual methods but also provide the means for addressing increasingly complex issues of protein engineering. Beyond the advances in experimental design, the development of in silico protein engineering is growing increasingly important. Progress in computational approaches now provide algorithms that simulate the nucleic acid fragment reassembly process during DNA shufﬂing [93] and assess the tolerance of individual amino acid substitution in the context of the global protein structure [154]. While the days of computer-based protein design are in the far future, the predictions from either modeling system can already provide guidance towards more effective experimental design. Acknowledgments

The authors would like to thank their colleagues in the laboratory and in particular Dr. Walter Fast for many helpful discussion and critical review of the manuscript. We would also like to acknowledge Dr. Marc Ostermeier and Steve Tizio for providing the unpublished data on the PurN/FMT hybrid system.

References 1 L. D. BOGARAD, M. W. DEEM, Proc. Natl. Acad. Sci. USA 1999, 96, 2591–2595. 2 C. H. CHENG, L. CHEN, Nature 1999, 401, 443–444. 3 S. A. DOYLE, S. Y. FUNG, D. E. KOSHLAND, Jr., Biochemistry 2000, 39, 14348–14355. 4 G. WU, A. FISER, B. TER KUILE, A. SALI, M. MULLER, Proc. Natl. Acad. Sci. USA 1999, 96, 6285–90. 5 R. KUROKI, L. H. WEAVER, B. W. MATTHEWS, Proc. Natl. Acad. Sci. USA 1999, 96, 8949–54. 6 K. MIYAZAKI, F. H. ARNOLD, J. Mol. Evol. 1999, 49, 716–720. 7 A. D. MCLACHLAN, Cold Spring Harb. Symp. Quant. Biol. 1987, 52, 411– 420. 8 B. M. LYNCH, J. S. CONERY, Science 2000, 290, 1151–55. 9 S. OHNO, Evolution by Gene Duplication, Springer Verlag, New York 1970.

10 L. PATTHY, Protein Evolution, Blackwell Science, Maiden, MA 1999. 11 A. FORCE, M. LYNCH, F. B. PICKETT, A. AMORES, Y. L. YAN, J. POSTLETHWAIT, Genetics 1999, 151, 1531–45. 12 R. A. JENSEN, Annu. Rev. Microbiol. 1976, 30, 409–425. 13 A. L. HUGHES, Proc. R. Soc. Land. B Biol. Sci. 1994, 256, 119–124. 14 J. PIATIGORSKY, G. WISTOW, Science 1991, 252, 1078–1079. 15 P. J. O’BRIEN, D. HERSCHLAG, Chem. Biol. 1999, 6, R91–R105. 16 R. FANI, P. LIO, A. LAZCANO, J. Mol. Evol. 1995, 41, 760–774. 17 R. FANI, P. LIO, I. CHIARELLI, M. BAZZICALUPO, J. Mol. Evol. 1994, 38, 489–495. 18 D. LANG, R. THOMA, M. HENN-SAX, R. STERNER, M. WILMANNS, Science 2000, 289, 1546–50.

References 19 J. TANG, M. N. JAMES, I. N. HSU, J. A. JENKINS, T. L. BLUNDELL, Nature 1978, 271, 618–621. 20 A. M. BAPTISTA, P. H. JONSON, E. HOUGH, S. B. PETERSEN, J. Mol. Evol. 1998, 47, 353–362. 21 A. D. MCLACHLAN, Eur. J. Biochem. 1979, 100, 181–187. 22 J. TANG, in A. J. BARRETT, N. D. RAWLINGS, J. F. WOESSNER (Eds.): Handbook of Proteolytic Enzymes, Vol. 272, Academic Press, San Diego CA, 1998, 805–814. 23 X. Y. QIU, J. S. CULP, A. G. DILELLA, B. HELLMIG, S. S. HOOG, C. A. JANSON, W. W. SMITH, S. S. ABDELMEGUID, Nature 1996, 383, 275–279. 24 D. REARDON, G. K. FARBER, FASEB J. 1995, 9, 497–503. 25 B. HOCKER, S. BEISMANN-DRIEMEYER, S. HETTWER, A. LUSTIG, R. STERNER, Nat. Struct. Biol. 2001, 8, 32–36. 26 S. JANECEK, S. BALAZ, J. Protein Chem. 1993, 12, 509–514. 27 K. LUGER, U. HOMMEL, M. HEROLD, J. HOFSTEENGE, K. KIRSCHNER, Science 1989, 243, 206–210. 28 V. MAINFROID, K. GORAJ, F. RENTIER-DELRUE, A. HOUBRECHTS, A. LOISEAU, A. C. GOHIMONT, M. E. NOBLE, T. V. BORCHERT, R. K. WIERENGA, J. A. MARTIAL, Protein Eng. 1993, 6, 893– 900. 29 C. P. PONTING, R. B. RUSSELL, Trends Biochem. Sci. 1995, 20, 179–180. 30 A. JELTSCH J. Mol. Evol. 1999, 49, 161–164. 31 E. A. MACGREGOR, H. M. JESPERSEN, B. SVENSSON, FEBS Lett. 1996, 378, 263–266. 32 J. JIA, W. HUANG, U. SCHORKEN, H. SAHM, G. A. SPRENGER, Y. LINDQVIST, G. SCHNEIDER, Structure 1996, 4, 715–724. 33 K. WIELIGMANN, B. NORLEDGE, R. JAENICKE, E. M. MAYR. J. Mol. Biol. 1998, 280, 721–729. 34 R. J. KREITMAN, R. K. PURI, I. PASTAN, Cancer Res. 1995, 55, 3357–3363. 35 G. D’ALESSIO, Prog. Biophys. Mol. Biol. 1999, 72, 271–298. 36 M. F. PERUTZ, Nature 1970, 228, 726– 739. 37 K. A. LEE, J. Cell Sci. 1992, 103, 9–14.

38 M. P. SCHLUNEGGER, M. J. BENNETT, D. EISENBERG, Adv. Protein Chem. 1997, 50, 61–122. 39 G. MACBEATH, P. KAST, D. HILVERT, Protein Sci. 1998, 7, 1757–1767. 40 M. OSTERMEIER, S. J. BENKOVIC, Adv. Protein Chem. 2000, 55, 29–77. 41 S. ZHANG, G. POHNERT, P. KONGSAEREE, D. B. WILSON, J. CLARDY, B. GANEM, J. Biol. Chem. 1998, 273, 6248–53. 42 J. TURNBULL, J. F. MORRISON, W. W. CLELAND, Biochemistry 1991, 30, 7783–7788. 43 R. M. ROMERO, M. F. ROBERTS, J. D. PHILLIPSON, Phytochemistry 1995, 39, 263–276. 44 F. POMPEO, Y. BOURNE, J. van HEIJENOORT, F. FASSY, D. MENGIN-LECREULX, J. Biol. Chem. 2001, 276, 3833–3839. 45 P. LJUNGCRANTZ, H. CARLSSON, M. O. MANSSON, P. BUCKEL, K. MOSBACH, L. BUELOW, Biochemistry 1989, 28, 8786–92. 46 C. LINDBLADH, M. PERSSON, L. BUELOW, K. MOSBACH, Eur. J. Biochem. 1992, 204, 241–247. 47 R. B. RUSSELL, Protein Eng. 1994, 7, 1407–1410. 48 M. NARDINI, B. W. DIJKSTRA, Curr. Opin. Struct. Biol. 1999, 9, 732–737. 49 A. E. TODD, C. A. ORENGO, J. M. THORNTON, J. Mol. Biol. 2001, 307, 1113–1143. 50 D. H. JUERS, R. E. HUBER, B. W. MATTHEWS, Protein Sci. 1999, 8, 122–136. 51 P. Van ROEY, V. RAO, Th. PLUMMER, Jr., A. L. TARENTINO, Biochemistry 1994, 33, 13989–96. 52 A. C. TERWISSCHA VAN SCHELTINGA, M. HENNIG, B. W. DIJKSTRA, J. Mol. Biol. 1996, 262, 243–57. 53 G. BUISSON, E. DUEE, R. HASER, F. PAYAN, EMBO J. 1987, 6, 3909–16. 54 Y. KAI, H. MATSUMURA, T. INOUE, K. TERADA, Y. NAGARA, T. YOSHINAGA, A. KIHARA, K. TSUMURA, K. IZUI, Proc. Natl. Acad. Sci. USA 1999, 96, 823–828. 55 Z. X. XIA, N. SHAMALA, P. H. BETHGE, L. W. LIM, H. D. BELLAMY, N. H. XUONG, F. LEDERER, F. S. MATHEWS, Proc. Natl. Acad. Sci. USA 1987, 84, 2629–33. 56 L. PATTHY, Cell 1985, 41, 657– 663.

31

32 S. Dedeblatt

57 J. A. KOLKMAN, W. P. STEMMER, Nat. Biotechnol. 2001, 19, 423–428. 58 M. L. TASAYCO, J. CAREY, Science 1992, 255, 594–597. 59 A. G. LADURNER, L. S. ITZHAKI, G. de PRAT GAY, A. R. FERSHT, J. Mol. Biol. 1997, 273, 317–329. 60 S. PASCARELLA, P. ARGOS, J. Mol. Biol. 1992, 224, 461–471. 61 N. DOI, H. YANAGAWA, Cell. Mol. Life Sci. 1998, 54, 394–404. 62 M. SAGERMANN, W. A. BAASE, B. W. MATTHEWS, Proc. Natl. Acad. Sci. USA 1999, 96, 6078–83. 63 M. OSTERMEIER, A. E. NIXON, J. H. SHIM, S. J. BENKOVIC, Proc. Natl. Acad. Sci. USA 1999, 96, 3562–67. 64 S. W. MICHNICK, I. REMY, F.X. CAMPBELL-VALOIS, A. VALLEE-BELISLE, J. N. PELLETIER, Methods Enzymol. 2000, 328, 208–230. 65 P. T. BEERNINK, Y. R. YANG, R. GRAF, D. S. KING, S. S. SHAH, H. K. SCHACHMAN, Protein Sci. 2001, 10, 528–537. 66 S. S. RAY, H. BALARAM, P. BALARAM, Chem. Biol. 1999, 6, 625–637. 67 C. MANOIL, J. BAILEY, J. Mol. Biol. 1997, 267, 250–263. 68 R. GRAF, H. K. SCHACHMAN, Proc. Natl. Acad. Sci. USA 1996, 93, 11591–96. 69 J. HENNECKE, P. SEBBEL, R. GLOCKSHUBER, J. Mol. Biol. 1999, 286, 1197–1215. 70 R. J. EUSTANCE, S. A. BUSTOS, R. F. SCHLEIF, J. Mol. Biol. 1994, 242, 330– 338. 71 D. M. NGUYEN, R. F. SCHLEIF, J. Mol. Biol. 1998, 282, 751–759. 72 M. OSTERMEIER, J. H. SHIM, S. J. BENKOVIC, Nat. Biotechnol. 1999, 17, 1205–1209. 73 K. VOGEL, J. CHMIELEWSKI, J. Am. Chem. Soc. 1994, 116, 11163–11164. 74 K. P. HOPFNER, E. KOPETZKI, G. B. KRESSE, W. BODE, R. HUBER, R. A. ENGH, Proc. Natl. Acad. Sci. USA 1998, 95, 9813–18. 75 M. M. ALTAMIRANO, J. M. BLACKBURN, C. AGUAYO, A. R. FERSHT, Nature 2000, 403, 617–622. 76 R. C. CADWELL, G. F. JOYCE, PCR Methods Appl. 1994, 3, S136–S140. 77 W. P. STEMMER, Nature 1994, 370, 389–391.

78 W. P. STEMMER, Proc. Natl. Acad. Sci. USA 1994, 91, 10747–10751. 79 K. CHEN, F. H. ARNOLD, Proc. Natl. Acad. Sci. C/SA 1993, 90, 5618–22. 80 J. C. MOORE, F. H. ARNOLD, Nat. Biotechnol. 1996, 14, 458–467. 81 A. CRAMERI, G. DAWES, E. RODRIGUEZ, S. SILVER, W. P. STEMMER, Nat. Biotechnol. 1997, 15, 436–438. 82 A. CRAMERI, S. A. RAILLARD, E. BERMUDEZ, W. P. STEMMER, Nature 1998, 391, 288–291. 83 J. E. NESS, M. WELCH, L. GIVER, M. BUENO, J. R. CHERRY, T. V. BORCHERT, W. P. STEMMER, J. MINSHULL, Nat. Biotechnol. 1999, 17, 893–896. 84 C. SCHMIDT-DANNERT, D. UMENO, F. H. ARNOLD, Nat. Biotechnol. 2000, 18, 750–753. 85 S. J. ALLEN, J. J. HOLBROOK, Protein Eng. 2000, 13, 5–7. 86 Z. SHAO, H. ZHAO, L. GIVER, F. H. ARNOLD, Nucleic Acids Res. 1998, 26, 681–683. 87 H. ZHAO, L. GIVER, Z. SHAO, J. A. AFFHOLTER, F. H. ARNOLD, Nat. Biotechnol. 1998, 16, 258–261. 88 Y. ZHANG, F. BUCHHOLZ, J. P. MUYRERS, A. F. STEWART, Nat. Genet. 1998, 20, 123–128. 89 M. KIKUCHI, K. OHNISHI, S. HARAYAMA, Gene 1999, 236, 159–167. 90 W. M. COCO, W. E. LEVINSON, M. J. CRIST, H. J. HEKTOR, A. DARZINS, P. T. PIENKOS, C. H. SQUIRES, D. J. MONTICELLO, Nat. Biotechnol. 2001, 19, 354–359. 91 V. SIEBER, C. A. MARTINEZ, F. H. ARNOLD, Nat. Biotechnol. 2001, 19, 456–460. 92 N. W. SOONG, L. NOMURA, K. PEKRUN, M. REED, L. SHEPPARD, G. DAWES, W. P. STEMMER, Nat. Genet. 2000, 25, 436–439. 93 G. L. MOORE, C. D. MARANAS, S. LUTZ, S. J. BENKOVIC, Proc. Natl. Acad. Sci. USA 2001, 98, 3226–31. 94 J. MINSHULL, W. P. STEMMER, Curr. Opin. Chem. Biol. 1999, 3, 284–290. 95 M. OSTERMEIER, S. J. BENKOVIC, Biotech. Lett. 2001, 23, 303–310. 96 S. LUTZ, M. OSTERMEIER, S. J. BENKOVIC, Nucleic Acids Res. 2001, 29, E16. 97 M. OSTERMEIER, A. E. NIXON, S. J. BENKOVIC, Bioorg. Med. Chem. 1999, 7, 2139–2144.

References 98 S. LUTZ, M. OSTERMEIER, G. MOORE, C. MARANAS, S. J. BENKOVIC, Proc. Natl. Acad. Sci. USA 2001, 98, 11248–11253. 99 S. E. RADFORD, Trends Biochem. Sci. 2000, 25, 611–618. 100 A. E. NIXON, M. OSTERMEIER, S. J. BENKOVIC, Trends Biotechnol. 1998, 16, 258–264. 101 L. B. MURATA, H. K. SCHACHMAN, Protein Sci. 1996, 5, 719–728. 102 R. WETZEL, L. J. PERRY, C. VEILLEUX, Biotechnology (NY) 1991, 9, 731–737. 103 Z. LIN, T. THORSEN, F. H. ARNOLD, Biotechnol. Prog. 1999, 15, 467– 471. 104 L. NIEBA, A. HONEGGER, C. KREBBER, A. PLUCKTHUN, Protein Eng. 1997, 10, 435–444. 105 G. E. DALE, C. BROGER, H. LANGEN, A. D’ARCY, D. STUBER, Protein Eng. 1994, 7, 933–939. 106 M. MURBY, E. SAMUELSSON, T. N. NGUYEN, L. MIGNARD, U. POWER, H. BINZ, M. UHLEN, S. STAHL, Eur. J. Biochem. 1995, 230, 38–44. 107 C. NISHIMURA, S. PRYTULLA, H. JANE DYSON, P. E. WRIGHT, Nat. Struct. Biol. 2000, 7, 679–686. 108 P. M. DALESSIO, I. J. ROPSON, Biochemistry 2000, 39, 860–871. 109 J. Van der SCHUEREN, J. ROBBEN, G. VOLCKAERT, Protein Eng. 1998, 11, 1211–1217. 110 M. G. ROSSMANN, P. ARGOS, Annu. Rev. Biochem. 1981, 50, 497–532. 111 C. C. BLAKE, Nature 1979, 277, 598. 112 M. GO, Nature 1981, 291, 90–92. 113 T. TSUJI, K. YOSHIDA, A. SATOH, T. KOHNO, K. KOBAYASHI, H. YANAGAWA, J. Mol. Biol. 1999, 286, 1581–96. 114 D. J. SEGAL, B. DREIER, R. R. BEERLI, C. F. BARBAS, 3rd, Proc. Natl. Acad. Sci. USA 1999, 96, 2758–63. 115 R. R. BEERLI, D. J. SEGAL, B. DREIER, C. F. BARBAS, 3rd Proc. Natl. Acad. Sci. USA 1998, 95, 14628–33. 116 M. MOORE, Y. CHOO, A. KLUG, Proc. Natl. Acad. Sci. USA 2001, 98, 1432–1436. 117 M. MOORE, A. MUG, Y. CHOO, Proc. Natl. Acad. Sci. USA 2001, 98, 1437–1441. 118 R. R. BEERLI, B. DREIER, C. F. BARBAS, 3rd Proc. Natl. Acad. Sci. USA 2000, 97, 1495–1500.

119 J. SMITH, J. M. BERG, S. CHANDRASEGARAN, Nucleic Acids Res. 1999, 27, 674–681. 120 I. FISCH, R. E. KONTERMANN, R. FINNERN, O. HARTLEY, A. S. SOLER-GONZALEZ, A. D. GRIFFITHS, G. WINTER, Proc. Natl. Acad. Sci. USA 1996, 93, 7761–7766. 121 S. MIKHEEVA, K. A. JARRELL, Proc. Natl. Acad. Sci. USA 1996, 93, 7486– 7490. 122 A. CRAMERI, S. CWIRLA, W. P. STEMMER, Nat. Med. 1996, 2, 100–102. 123 F. B. PERLER, Trends Biochem. Sci. 1999, 24, 209–211. 124 H. WU, Z. HU, X. Q. LIU, Proc. Natl. Acad. Sci. USA 1998, 95, 9226–31. 125 F. B. PERLER, Nucleic Acids Res. 2000, 28, 344–345. 126 T. OTOMO, N. ITO, Y. KYOGOKU, T. YAMAZAKI, Biochemistry 1999, 38, 16040–44. 127 N. SHEIBANI, Prep. Biochem. Biotechnol. 1999, 29, 77–90. 128 R. B. KAPUST, D. S. WAUGH, Protein Sci. 1999, 8, 1668–74. 129 L. MAXWELL, A. K. MITTERMAIER, J. D. FORMAN-KAY, A. R. DAVIDSON, Protein Sci. 1999, 8, 1908–1911. 130 S. WALDO, B. M. STANDISH, J. BERENDZEN, T. C. TERWILLIGER, Nat. Biotechnol. 1999, 17, 691–95. 131 C. WIGLEY, R. D. STIDHAM, N. M. SMITH, J. F. HUNT, P. J. THOMAS, Nat. Biotechnol. 2001, 19, 131–136. 132 GEORGIOU, C. STATHOPOULOS, P. S. DAUGHERTY, A. R. NAYAK, B. L. IVERSON, R. CURTISS, Nat. Biotechnol. 1997, 15, 29–34. 133 T. LATTEMANN, J. MAURER, E. GERLAND, T. F. MEYER, J. Bacterial. 2000, 182, 3726–3733. 134 T. BODER, K. D. WITTRUP, Methods Enzymol. 2000, 328, 430–444. 135 M. BETTON, J. P. JACOB, M. HOFNUNG, J. K. BROOME-SMITH, Nat. Biotechnol. 1997, 15, 1276–79. 136 MARTINEAU, J. G. GUILLET, C. LECLERC, M. HOFNUNG, Gene 1992, 113, 35–46. 137 J. M. BETTON, P. MARTINEAU, W. SAURIN, M. HOFNUNG, FEBS Lett. 1993, 325, 34–38. 138 J. M. BETTON, M. HOFNUNG, EMBO J. 1994, 13, 1226–1234.

33

34 S. Dedeblatt

139 B. COLLINET, M. HERVE, F. PECORARI, P. MINARD, O. EDER, M. DESMADRIL J. Biol. Chem. 2000, 275, 17428–33. 140 J. AY, F. GOTZ, R. BORRISS, U. HEINEMANN, Proc. Natl. Acad. Sci. USA 1998, 95, 6613–18. 141 G. S. BAIRD, D. A. ZACHARIAS, R. Y. TSIEN, Proc. Natl. Acad. Sci. USA 1999, 96, 11241–46. 142 N. DOI, H. YANAGAWA, FEBS Lett. 1999, 453, 305–307. 143 D. E. CANE, C. T. WALSH, C. KHOSLA, Science 1998, 282, 63–68. 144 R. S. GOKHALE, S. Y. TSUJI, D. E. CANE, C. KHOSLA, Science 1999, 284, 482–485. 145 R. S. GOKHALE, C. KHOSLA, Curr. Opin. Chem. Biol. 2000, 4, 22–27. 146 S. Y. TSUJI, N. WU, C. KHOSLA, Biochemistry 2001, 40, 2317–25. 147 R. MCDANIEL, A. THAMCHAIPENET, C. GUSTAFSSON, H. FU, M. BETLACH, G. ASHLEY, Proc. Natl. Acad. Sci. USA 1999, 96, 1846–1851. 148 Q. XUE, G. ASHLEY, C. R. HUTCHINSON, D. V. SANTI, Proc. Natl. Acad. Sci. USA 1999, 96, 11740–45. 149 Y. SHEN, P. YOON, T. W. YU, H. G. FLOSS, D. HOPWOOD, B. S. MOORE, Proc. Natl. Acad. Sci. USA 1999, 96, 3622–27. 150 J. PIEL, C. HERTWECK, P. R. SHIPLEY, D. M. HUNT, M. S. NEWMAN, B. S. MOORE, Chem. Biol. 2000, 7, 943–955. 151 P. J. BELSHAW, C. T. WALSH, T Stachelhaus, Science 1999, 284, 486– 489. 152 H. D. MOOTZ, D. SCHWARZER, M. A. MARAHIEL, Proc. Natl. Acad. Sci. USA 2000, 97, 5848–5853. 153 T. STACHELHAUS, H. D. MOOTZ, M. A. MARAHIEL, Chem. Biol. 1999, 6, 493– 505. 154 C. A. VOIGT, S. L. MAYO, F. H. ARNOLD, Z. G. WANG, Proc. Natl. Acad. Sci. USA 2001, 98, 3778–83. 155 R. J. ALMASSY, C. A. JANSON, C. C. KAN, Z. HOSTOMSKA, Proc. Natl. Acad. Sci. USA 1992, 89, 6114–18. 156 E. SCHMITT, S. BLANQUET, Y. MECHULAM, EMBO J. 1996, 15, 4749–58. 157 R. MCDANIEL, C. M. KAO, S. J. HWANG, C. KHOSLA, Chem. Biol. 1997, 4, 667– 674.

158 R. MCDANIEL, S. EBERT-KHOSLA, D. A. HOPWOOD, C. KHOSLA, Nature 1995, 375, 549–554. 159 D. BEDFORD, J. R. JACOBSEN, G. LUO, D. E. CANE, C. KHOSLA, Chem. Biol. 1996, 3, 827–831. 160 OLIYNYK, M. J. BROWN, J. CORTES, J. STAUNTON, P. F. LEADLAY, Chem. Biol. 1996, 3, 833–839. 161 H. SYMMANK, W. SAENGER, F. BERNHARD, J. Biol. Chem. 1999, 274, 21581–21588. 162 S. DOEKEL, M. A. MARAHIEL, Chem. Biol. 2000, 7, 373–384. 163 C. W. CARRERAS, D. V. SANTI, Curr. Opin. Biotech. 1998, 9, 403–411. 164 L. TANG, H. FU, R. MCDANIEL, Chem. Biol. 2000, 7, 77–84. 165 A. RANGANATHAN, M. TIMONEY, M. BYCROFT, J. CORTES, I. P. THOMAS, B. WILKINSON, L. KELLENBERGER, U. HANEFELD, I. S. GALLOWAY, J. STAUNTON, P. F. LEADLAY, Chem. Biol. 1999, 6, 731–741. 166 C. M. KAO, G. L. LUO, L. KATZ, D. E. CANE, C. KHOSLA, J. Am. Chem. Soc. 1995, 117, 9105–06. 167 C. M. KAO, G. L. LUO, L. KATZ, D. E. CANE, C. KHOSLA, J. Am. Chem. Soc. 1996, 118, 9184–85. 168 J. CORTES, K. E. WIESMANN, G. A. ROBERTS, M. J. BROWN, J. STAUNTON, P. F. LEADLAY, Science 1995, 268, 1487–89. 169 A. M. GEHRING, E. DEMOLL, J. D. FETHERSTON, I. MORI, G. F. MAYHEW, F. R. BLATTNER, C. T. WALSH, R. D. PERRY, Chem. Biol. 1998, 5, 573–586. 170 D. TILLETT, E. DITTMANN, M. ERHARD, H. VON DOHREN, T. BORNER, B. A. NEILAN, Chem. Biol. 2000, 7, 753–764. 171 Y. PAITAN, G. ALON, E. ORR, E. Z. RON, E. ROSENBERG, J. Mol. Biol. 1999, 286, 465–474. 172 J. F. APARICIO, I. MOLNAR, T. SCHWECKE, A. KONIG, S. F. HAYDOCK, L. E. KHAW, J. STAUNTON, P. F. LEADLAY, Gene 1996, 169, 9–16. 173 A. F. MARSDEN, B. WILKINSON, J. CORTES, N. J. DUNSTER, J. STAUNTON, P. F. LEADLAY, Science 1998, 279, 199–202. 174 S. KUHSTOSS, M. HUBER, J. R. TURNER, J. W. PASCHAL, R. N. RAO, Gene 1996, 183, 231–236.

1

Engineered Nanopores Hagan Bayley, and Orit Braha

University of Oxford, Oxford, United Kingdom Stephen Cheley, and Li-Qun Gu Texas A&M University, College Station, USA

Originally published in: Nanobiotechnology. Edited by Christof M. Niemeyer and Chad A. Mirkin. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30658-9

1 Overview

Engineered nanopores have potential applications in many areas of technology, including separation science, sensing, and drug delivery. In this chapter, we focus on recent developments regarding pores that span membranes and have diameters of 10 nm or less. We emphasize pores into which new properties have been engineered, highlighting our own recent investigations into α-hemolysin. 1.1 What is a Nanopore?

Nanopores occur in nature, and in the biological literature they are simply known as pores when they are more than 1 nm or so in diameter, or channels when they are narrower. For certain applications in technology, biological pores are being engineered directly. In other cases, new types of pores are being constructed that are based on biological structures [1]. The biological pores that concern us here are transmembrane proteins. Cells also contain soluble pores, such as chaperonins and enzymes that handle nucleic acids, which may be of interest in the future in the context of engineering catalysis into nanopores. Transmembrane proteins fall into two main classes: helix bundles, and β barrels [2]. In general, helix bundles constitute various classes of receptors and channels. Natural examples of these proteins are relatively difﬁcult to engineer: (i) because the helices are usually in close contact and alterations to the side chains disrupt the structure of the protein; and (ii) because they have evolved highly specialized functions, which tend to be difﬁcult to Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Engineered Nanopores

Fig. 1 Examples of various classes of nanopore. (a) Helix bundle in the form of a template-assembled synthetic protein (TASP) [116]. (b) A single subunit of the trimeric porin OmpF (inner diameter ∼2 nm) [117]. (c) The "-hemolysin pore (inner diameter of barrel ∼2 nm) [13]. (d) Nanotube formed from stacked cyclic peptides (inner diameter ∼0.75 nm) [16]. (e) Syn-

thetic barrel based on octiphenyl staves with attached peptides (inner diameter ∼1.5 nm) [20]. (f) Electron micrograph of silica nanotubes made in a porous alumina template (inner diameter, e.g., ∼20 nm). The template was subsequently dissolved [1]. (g) Electron micrograph of a nanopore in silicon nitride sculpted with an Ar ion beam (inner diameter ∼2 nm) [27].

change. Nevertheless, some of the earliest attempts to build membrane proteins de novo involved transmembrane helices. For example, bundles of individual synthetic helices were conceived by the groups of DeGrado and Lear [3], and templateassembled synthetic proteins (TASP) were used initially by Montal, Vogel and colleagues (Figure 1a) [4]. Some progress has been made on de novo single-chain bundles [5, 6], but as yet this work lacks a solid structural basis. By contrast, β-barrels are open structures, permitting dramatic alterations of natural scaffolds [7]. So far, the β-barrel fold has been found in all the outer membrane proteins of Gram-negative bacteria that have been examined, as well as in certain poreforming toxins. Importantly, the side chains that project into a β barrel do not interact strongly with one another, and can be altered to manipulate the properties of the pores. Recently, progress has been made on barrel-like structures formed from helices [8, 9], and we can expect more work in this area in the future. Among the natural channels that have been engineered as nanopores, the most prominent have been porins, proteins that control the permeability of the bacterial outer membrane, and α-hemolysin (αHL), a pore-forming toxin secreted by Staphylococcus aureus. Work on both these systems has been aided by highresolution crystal structures (Figure 1b,c). At this point, the engineering of porins

1 Overview

has been limited, but they do constitute a promising system [10–12]. The αHL pore is a heptameric structure (Figure 1c) [13]. The transmembrane β barrel is topped by a cap domain that contains a large internal cavity. Both the cap and barrel have been engineered by various means (see below). Our laboratory has developed methods for producing pure heteromeric pores, most usefully of the form WT6 ENG1 with just one altered subunit. The engineered subunit (ENG) is chemically or genetically tagged to confer an altered charge. The wild-type (WT) and engineered subunits are then co-assembled and the desired heteromer isolated by preparative sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) [14]. αHL belongs to a family of toxins, which includes the leukocidins. The latter form pores comprising two different, but related, polypeptide chains. We have shown that one leukocidin pore is an octamer, rather than a heptamer, containing four subunits of each class [15]. The fact that both heptamers and octamers exist suggests that it may be possible to control diameter in this family of pores by engineering the subunit–subunit interface. Designed pores containing β structure have been developed by the groups of Ghadiri and Matile. The Ghadiri group pioneered this area by introducing stacked cyclic peptides containing alternating D- and L-amino acids, which form nanotubes that span lipid bilayers (Figure 1d) [16]. The structures resemble gramicidin, a naturally occurring peptide antibiotic that forms transmembrane pores from two head-to-head β spirals. The nanotubes can be stabilized by stapling them together in a metathesis reaction between oleﬁnic amino acid side chains [17]. The diameter of the nanotubes can be adjusted by altering the ring size of the peptides [18], and β-amino acids can form similar structures [19]. Matile’s group has developed synthetic barrels from octiphenyl staves to which all L-amino acid peptides are attached (Figure 1e) [20]. In principle, the structure of the octiphenyl barrels is highly manipulatable. The diameter can be changed by altering the length of the attached peptides, and the number of staves (which depends upon the steric and electrostatic interactions between the peptides) [21]. The length of the barrel can be controlled by adjusting the length of the stave [22]. Finally, heteromeric pores containing two types of staves have been assembled [21]. While limited structural information has been obtained on Ghadiri’s nanotubes [23], the structures of the octiphenyl barrels are based on model building and a substantial body of indirect evidence. Several groups have made nanopores from materials with no connection to biology [1]. For example, commercial track-etched polymer membranes are made by bombarding thin ﬁlms with nuclear ﬁssion particles. The damage tracks are then etched, with base for example in the case of polycarbonate, to form uniform pores as small as 10 nm in diameter, which may be several µm in length. To make smaller pores of far shorter effective length, asymmetric etching techniques have been developed to generate cone-shaped structures [24]. For example, poly(ethylene terephthalate) through which a single high-energy Au ion had passed was treated with concentrated sodium hydroxide from one side, with a stop solution of formic acid on the other side. The membrane was in an electrical cell so that breakthrough could be registered and etching stopped at that point. Further, the applied potential was such that hydroxide ion moved away from the etch site after

3

4 Engineered Nanopores

breakthrough (electro-stopping). Individual pores formed in this way exhibit many of the properties of biological pores, such as ion selectivity, and open and closed states [25]. The chemically etched pores will probably be difﬁcult to make and derivatize in a reproducible way. Fortunately, Martin’s group has developed sophisticated approaches for generating nanotubules with modiﬁable internal surfaces within the pores of track-etched polycarbonate ﬁlters. For example, a gold ﬁlm is deposited on the surface of the pores by electroless plating [26]. This serves both to control the diameter of the nanotubules, which can be made as small as 1 nm, and to provide a reactive surface (see below). Individual nanopores have also been made by using a low-energy beam of Ar ions to “sculpt” a hole in a thin layer of silicon nitride [27]. Surprisingly, it was demonstrated that the beam can close as well as open a hole, depending on the beam ﬂux and duty cycle and the sample temperature. Because the size of the hole could be monitored during sculpting by counting the transmitted Ar ions, it was possible to produce nanopores of known diameter by feedback control of the beam, and in this way diameters below 1.8 nm were obtained (Figure 1g). 1.2 Engineering Nanopores

Protein pores are engineered by using mutagenesis and targeted chemical modiﬁcation. The primary interest lies in engineering the interior (lumen) of the pores. Mutagenesis can be used to introduce any of the twenty natural amino acids, which provide a variety of side chains of differing size, shape, polarity, and reactivity. Nonnatural amino acid mutagenesis can also be used to provide side chains that do not occur in nature, such as ketones, alkenes, and azides. Originally, non-natural amino acids were introduced by using preloaded suppressor tRNAs in an in-vitro translation system, a technology that is sufﬁciently demanding to prevent its routine use. Currently, Escherichia coli strains are under development that provide the mutant suppressor tRNAs and the engineered synthetases required to load them. Indeed, it is even possible to have the bacteria make an amino acid [28]! In targeted chemical modiﬁcation, an amino acid side chain (usually cysteine) is selectively modiﬁed with a reagent. This again adds to the diversity of functionalities that can be incorporated into a protein. Given a high-resolution structure, these approaches to protein engineering allow modiﬁcations to be placed at well-deﬁned locations. If more than one modiﬁcation is made, the geometrical relationship between the new side chains is known. Our group recently implemented a new concept in protein engineering – that of non-covalent modiﬁcation. Speciﬁcally, cyclodextrins and other host molecules (adapters) were allowed to bind to positions within the lumen of the transmembrane barrel of the αHL pore that could be deﬁned by mutagenesis (Figure 2) [29–32]. The adapters remain capable of binding guests and therefore noncovalent modiﬁcation can introduce new binding sites within the pore. The vast literature on host– guest interactions supports the enormous potential of this approach.

1 Overview

Fig. 2 "-Hemolysin pore modiﬁed with noncovalent adapters. The illustration is based on the superposition of structures of "HL and cyclodextrins. The positions of the cyclodextrins were surmised from

mutagenesis experiments. In the structure shown, sites for two different cyclodextrins were engineered. An organic molecule can be trapped in the cavity between the two sites [31].

Designed pores containing β structure, such as those of Ghadiri and Matile, can be engineered using chemical synthesis with a variety of amino acids, and of course there is no requirement for using the natural side chains. The side chains in Ghadiri’s nanotubes all project outwards, which makes it difﬁcult to engineer function, although progress has been made by using functionalized peptides to cap the structures [33]. In Matile’s synthetic barrels, just as in a natural β barrel, the side chains of the amino acids project alternately outwards and into the lumen, permitting the introduction of functional groups within the pore. However, although it is in principle possible, the exquisite precision of protein engineering, in which individual changes are made at speciﬁc positions, has not yet been achieved with the synthetic barrels. At present, for example, 32 or 64 histidine residues are introduced at one time. As expected, the internal surfaces of Martin’s gold-plated nanotubules are amenable to derivatization with a variety of thiols that vary in size, shape, and polarity [34]. Interestingly, the surfaces of the gold-plated nanotubules can also be charged electrostatically by using an applied potential [35]. Recently, the idea of electromodulation has been demonstrated in which electrostatic charging is used to attract an amphiphilic surfactant to the wall of the nanotubule and thereby render the surface hydrophobic [36]. By using silane chemistry, proteins have been attached to the internal surface of another class of nanotubules made of silica in an alumina template (see Figure 1f) [37]. Asymmetrically etched poly(ethylene terephthalate) can also be chemically modiﬁed. For example, the free carboxyl groups can be neutralized by methylation with diazomethane [25] and additional chemistry that would alter surface properties can readily be conceived. Nevertheless, in these

5

6 Engineered Nanopores

cases, the inability to control the number of modiﬁed sites or organize them in space remains a severe drawback. 1.3 What Can a Nanopore Do?

In Arthur Karlin’s succinct words: “Ion channels open, conduct ions selectively, and close” (Figure 3) [38]. The nanopores we are considering also conduct ions, and even though the diameters of the pores are larger than the typical ion channel, they do exhibit at least modest ion selectivity. Importantly, because they are large they can also transport large molecules across a membrane or bilayer. Natural pores, such as gap junctions, can gate (open and close in response to a stimulus), but so far little attention has been given to this issue with engineered nanopores. More important has been the generation of sites within the pores at which blockers can bind (Figure 3d), by analogy with drugs such as verapamil, a Ca-channel blocker. These properties of pores can be elaborated and enhanced by various engineering techniques, and we describe selected examples here. Emphasis is given to investigations on the αHL pore, for which such studies are relatively advanced. The ion selectivity of porins [39] and the αHL pore [32] can be altered by the replacement of residues within the lumen. In general, positively charged residues produce anion selectivity, and negatively charged residues cation selectivity. Similarly, in the case of Matile’s artiﬁcial barrels, the amino acid side chains that project into the lumen can be used to control selectivity [40]. Ghadiri’s nanotubes remain cationselective in the presence of positively charged cyclic peptide caps, but the selectivity is substantially reduced [33]. When the neutral adapter molecule β-cyclodextrin is lodged within the lumen of the αHL pore, the weak anion selectivity of the pore is enhanced. By contrast, when the anionic hepta-6-sulfato-β-cyclodextrin is used, the pore becomes cation-selective [41]. Ion selectivity can also be controlled with a tightly adsorbed but noncovalently bound internal surface layer. Matile and colleagues have

Fig. 3 Basic properties of channels and pores. (a) Ion conduction; (b) ion selectivity; (c) gating; (d) channel block.

1 Overview

shown that phosphate tightly bound to positively charged residues in the lumen of artiﬁcial barrels renders them cation-selective, while Mg2+ ions bound to internal carboxylates render the barrels anion-selective [40]. Underivatized Au nanotubes are unselective in KF solutions, but cation-selective in KCl and KBr because of adsorption of anions to the Au surface [35]. The selectivity of the Au nanotubes can also be controlled by direct electrostatic charging of the metal surface [35]. There is still a great deal to learn about the transport of larger molecules through nanopores. For rigid molecules there is molecular mass cut-off, although transport will be reduced before the cut-off is reached because of an increased barrier to entry (hard-sphere model) and interactions with the walls during transport. In one example, glucose was shown to pass through nanotubes made from cyclic peptides of ten amino acids, but not through those of eight amino acids [18]. Au nanotubules can also mediate size separation [26]. Several examples of facilitated transport have been demonstrated with various derivatized inorganic nanotubules. In the case of small organic molecules, a high degree of selectivity has been obtained after reacting the walls of Au nanotubules with various alkane thiols. For example, nanotubules with a low diameter of 1.5 nm that had been reacted with hexadecanethiol transported toluene more than 100 times faster than pyridine in aqueous solution [34]. It seems likely that the derivatized nanotubules do not ﬁll with water, and that the toluene diffuses through an internal hydrophobic phase. Electromodulation of the hydrophobicity of the interior of Au nanotubules is a recent innovation [36]. The enantiomers of chiral molecules have been separated in nanotubules [37, 42]. For example, a racemate can be passed though silica nanotubules within which a stereospeciﬁc antibody has been attached [37]. It is likely that this separation works by a facilitated transport mechanism in which the permeant molecules partition strongly into nanotubules containing a high density of cognate binding sites. For facilitated transport to occur, these sites must exchange the permeant molecules rapidly. In the case of the αHL pore, the focus has been on the transport of macromolecules. The cut-off for a rigid spherical molecule based on the width of the narrowest constriction in the lumen is less than 1000 Da, but this has not been tested in rigorous experiments. Elongated polymers of much higher mass such as single-stranded DNA [43] and poly(ethylene glycol) (PEG) [44] can pass through the αHL pore. For a highly ﬂexible molecule such as PEG there is a high entropic cost to entry into the pore, which greatly reduces the rate of transport [45]. A great deal of experimental [46, 47] and theoretical effort ([48] and references therein) continues to be devoted to this aspect of nanopore research because of the interest in making sensors for various polymers. α-Helical ion channels are gated (opened and closed) by voltage, ligands, or mechanical force. β-Barrel pores gate naturally by mechanisms that are poorly understood and depend upon variables such as the pH and applied potential. Attempts are being made to engineer voltage gating into porins. For example, positively charged peptide loops have been inserted into turns of a porin. The applied potential pushes the charged insert into the barrel at one polarity and removes it at the other [11], which is akin to the so-called ball-and-chain inactivation of K+ channels [49]. Although weakly asymmetric conduction has been achieved with the porin, more

7

8 Engineered Nanopores

work is required to obtain clear-cut gating. Certain derivatives of gramicidin also appear to gate in a ball-and-chain fashion [50]. There are several other attractive possibilities for controlling the activity of pores. For example, attempts have been made to open and close pores with light, and in such an attempt Woolley and colleagues attached an azobenzene near the mouth of the gramicidin pore [51]. But again, allor-none gating has not yet been achieved. Interestingly, control of the access of organic molecules to cavities within mesoporous silica was recently achieved through reversible photochemical dimerization of covalently attached coumarins [52]. A variety of blocker sites have been engineered into the αHL pore (Figure 4), in this case with a view towards sensor technology (see below). The simplest sites involve mutagenesis with natural amino acids. For example, the introduction of histidine residues at sites within the lumen of the αHL pore allows block by various divalent metal cations [14, 53], while the introduction of arginine residues permits the binding of anionic phosphate esters [54] (Figure 4a). Sites for organic molecules can be formed from noncovalent adapters (Figure 4b) [29–31, 55], although in principle other forms of engineering might be used. Interestingly, the cap domain of the αHL pore, which contains a 45 Å-diameter cavity, is large enough to contain a short covalently attached oligonucleotide, an 8-mer for example (Figure 4c). The tethered oligonucleotide can act as a binding site for complementary strands [56–58]. Most proteins are too large to be accommodated within the lumen of the αHL pore. Nevertheless, the attachment of a protein to an external ligand can be transmitted to the interior and hence affect the conductance of the pore. This has been achieved by attaching the ligand to the end of a molecular ﬁshing line, namely a long PEG chain [59] (Figure 4d). When the protein is bound, the mobility of the part of the chain that lies within the pore is altered, which produces a characteristic change in the electrical signal during bilayer recording. In addition to the fundamental properties of pores discussed above, there are additional features that might be engineered. For example, many membrane proteins form two-dimensional crystals, which have contributed to structural studies. Engineered arrays of protein nanopores might be exploited in the same way as bacterial S-layers. S-layers are porous, two-dimensional arrays of a single protein species that envelope a variety of bacteria and are being explored for various applications

Fig. 4 Blocker and related sites in the "HL pore. (a) Genetically engineered site for phosphate anions [54]. (b) Site for organic molecules formed with a noncovalent adapter [29]. (c) Covalently attached oligonucleotide for duplex formation [57]. (d) Ligand attached to a long PEG chain for the capture of protein analytes [59].

1 Overview

including ultraﬁltration, the geometrically deﬁned immobilization of macromolecules, and acting as templates for the formation of arrays of nanocrystals [60]. 1.4 What are the Potential Applications of Nanopores?

There are many exciting prospects for the utilization of engineered nanopores, and in some cases practical applications are emerging. For example, as described above, Martin and colleagues have effected separations with various modiﬁed inorganic nanopores. The separations have been based on several properties including molecular mass [26], charge [35], hydrophobicity [34, 36], and even stereochemistry [37, 42]. In principle, similar separations might be carried out with engineered protein nanopores, but with the exception of S-layers with which size separation has been accomplished [60], proteins have not yet been exploited for this purpose. Another area of application of protein nanopores is in cell permeabilization. Because of its importance, the utility of reversible plasma membrane permeabilization in cell and tissue preservation has received the most attention. A mutant αHL pore, H5, which can be closed with divalent metal ions, can be used to introduce small molecules into cells [61]. Once the cells are loaded, efﬂux can be prevented by Zn(II) ions at the micromolar concentrations normally found in plasma. This procedure has been used to introduce sugars such as sucrose and trehalose into cells (Figure 5) [62]. Cells treated in this way have far higher survival rates after cryopreservation [63] and desiccation [64, 65] than those that have not been treated. Various therapeutic approaches might also be mediated by nanopores. First, nanopores might be used in direct attacks on infectious or malignant cells. For example, Ghadiri’s group has demonstrated that selected cyclic peptides have a rapid bactericidal action on both Gram-negative and Gram-positive species [66]. They suggest that the nanotubes formed from the peptides act by assembling at target membranes and forming “carpets”, thereby rendering them leaky, rather than by acting as individual pores. Malignant mammalian cells might repair themselves

Fig. 5 Loading mammalian cells with trehalose by using "HL-H5 pores, which can be blocked completely by Zn(II) ions [63].

9

10 Engineered Nanopores

after permeabilization, and it has been suggested that the effect of pore formation might be enhanced in the presence of cytotoxic molecules that would not normally enter target cells. Selectivity might be achieved by directing poreforming proteins to target cells with antibodies or lectins. Additional selectivity might be achieved by activation of the poreforming protein at the target cell surface. Along these lines, αHL polypeptides that can be activated by a tumor protease, cathepsin B, have been obtained by screening a library of inactive two-chain αHL mutants containing a combinatorial cassette encoding thousands of potential protease recognition sites [67]. Nanopores have excellent prospects in sensor technology. For example, the ionic current ﬂowing through a pore in a transmembrane potential and the modulation of the current by analytes that act, for instance, as blockers can be monitored. In this way, sensors based on either many pores (macroscopic currents) or single pores are being devised. Here, we focus on single pores and the reader is referred to reviews, where key papers on macroscopic approaches can be located [68–70]. Single pores are used in stochastic sensing where individual interactions between the pores and analyte molecules are detected [47, 70, 71]. In stochastic sensing, the frequency of occurrence of individual binding events reveals the concentration of an analyte, while the analyte can be identiﬁed from the signature of the binding events (the extent and duration of current block, for example) (Figure 6). As indicated in the previous section, a variety of blocker sites have been engineered into the αHL pore, and they can be used to detect a wide variety of analytes ranging from small cations and anions, to organic molecules, to macromolecules including DNA and proteins. Polymers can be detected by nanopores in a second mode in which their transit through the pores is registered [46, 47]. This was ﬁrst demonstrated with single-stranded DNA [43], but there is no reason why the procedure cannot be adapted to other polymers. Here, the nanopore is acting as more than a simple Coulter counter, because limited information about the structure of the polymer is

Fig. 6 Basis of stochastic sensing [71]. Individual interactions between single pores and analyte molecules are detected. (a) Engineered nanopore in the absence of analyte. A ﬂat current trace is seen. (b) Engineered nanopore in the presence of analytes. In-

dividual binding events are manifested as transient reductions in current. The frequency of occurrence of binding events reveals the concentration of an analyte. Further, an analyte can be identiﬁed from the signatures of the events.

1 Overview

revealed by the changes in current during transit and the transit velocity. It has been proposed that DNA might be sequenced by this approach, but this seems to be an unlikely proposition in its present formulation [47]. However, protein engineering has not been applied to this idea in earnest, and it might be of help. Finally, engineered nanopores have applications in basic science. For example, the ability to introduce molecules into cells is an important tool in cell biology, and pore-forming proteins are one of the devices that have been used for this purpose [72]. Obviously, the size of the pore determines what goes in and, just as importantly, what comes out. For example, αHL has often been used to bring small molecules (up to ∼1000 Da) into cells, while streptolysin O (which forms pores ∼35 nm in diameter) can be used to transfer macromolecules of up to 100 kDa. Prolonged permeabilization of cells is lethal, and so a means of closing the pores is required. In some cases, wild-type lesions can be eliminated. For example, resealing of the pores formed by streptolysin O occurs by an unknown mechanism that is promoted by Ca2+ ions [73]. Engineered pores can also be used and αHL-H5, which can be closed with divalent metal ions [61], has found several application in the laboratory. For example, Palczewski and colleagues used αHL-H5 to load nucleotides into retinal rod cells [74]. In our own laboratory, it is becoming clear that the αHL pore can be used as a nano-reactor for the investigation of both noncovalent and, most recently, covalent interactions at the single molecule level. Single molecule studies are capable of revealing information that is obscured in ensemble measurements [75]. These studies stem from our attempts to make sensor elements from the αHL pore (see above). In the case of sensors, it does not matter whether the kinetics seen in bulk solution are maintained inside or close to the pore. However, our studies suggest that several binary noncovalent interactions occur with similar kinetics and thus possess similar Kd values to those observed in bulk solvent. These include host–guest interactions [29], DNA duplex formation [56], and protein-ligand interactions [59]. While we can say that association and dissociation rate constants are within a factor of ten of those in bulk solvent, more detailed comparisons will be required to ascertain just how close they are. One partner in each of these interactions was a small molecule. Where a macromolecule must assume a compact state to enter the pore, overall Kd values are increased [45, 76]. In recent studies, we have investigated a variety of covalent chemistries within the αHL pore at the single molecule level, and again the observed kinetic constants are close to those seen in bulk solvent [77, 78]. 1.5 Keeping Nanopores Happy

One issue of great importance in sensor technology is the search for more robust platforms for existing nanopores (Figure 7), or simply more robust nanopores [71]. This is an area that is gaining from several energing aspects of nanotechnology. The lipid bilayers used in the laboratory are usually formed across apertures in polymer ﬁlms, such as Teﬂon. These bilayers are mechanically unstable, and a great deal of work is aimed at obtaining a storable and shippable alternative [79–82]. One opportunity lies with improved apertures, and several based on silicon

11

12 Engineered Nanopores

Fig. 7 Different classes of robust bilayer. (a) Bilayer across a small aperture formed in silicon nitride [83]. (b) Bilayer on a porous support comprising a bacterial S-layer [84]. (c) Tethered supported bilayer on a smooth gold surface [86].

substrates have been reported. For example, Peterman and colleagues recently showed that bilayers can be formed across 25-µm apertures in silicon nitride made by established fabrication technology [83] (Figure 7a). The capacitance of the device was reduced by oxidation of exposed silicon substrate and the use of a polyimide surface coating. The bilayers were more robust than those on Teﬂon apertures, as judged by their ability to withstand high potentials. Another promising area is the use of porous supports, such as nanoporous glass, agarose, etc. One interesting example, with which steady improvements are being made, is the use of S-layers [84], the porous two-dimensional protein crystals from bacterial envelopes (Figure 7b). Recent investigations have demonstrated that S-layer-supported bilayers have an increased resistance to hydrostatic pressure [85]. The exploration of bilayers on solid supports continues. In general, these bilayers have been too leaky for single-channel current recording. Recently, tethered supported bilayers on ultrasmooth Au surfaces have been obtained with speciﬁc capacitance and impedance values rivaling those of planar lipid bilayers [86] (Figure 7c). Therefore, if sufﬁciently small surface areas are used, it should be possible to examine single nanopores in such systems [87]. However, a second serious issue is the high resistance of the volume between the bilayer and the support, which is a consequence of the modest reservoir of mobile ions, at least with tethered bilayers [88, 89]. It may also be possible to replace lipid bilayers completely. For example, polymerizable block copolymers appear to be a promising substitute [90]. In short, progress has been made, but we do not yet have a bilayer that can be slipped into a briefcase.

2 Methods

In this article we have covered a wide variety of types of nanopores, and it would be presumptuous of us to discuss the methodology involved in areas where we do not work directly. Instead, we present a summary of the methods that have been crucial for the progress that has been made with the αHL pore. 2.1 Protein Production

The monomeric form of αHL and its mutants are obtained in abundance and high purity by expression in Staphylococcus [91] or E. coli [92]. When necessary, the

2 Methods 13

puriﬁcations can be aided with C-terminal His tags. Conveniently, small amounts of αHL are available by in-vitro transcription and translation, which produces around 50 µg mL−1 [92]. For single-channel recording in the laboratory, we use only nanograms of protein. Other applications likewise require modest amounts; for example, a 1 m2 , two-dimensional crystal of αHL pores would contain only 3.5 mg of protein. The in-vitro production technology is readily scaled up to provide milligrams and even grams [93]. While assembly into heptamers can be facilitated by receptors on target cells, such as rabbit red blood cells [94], the αHL pore self-assembles on artiﬁcial lipid bilayers [95]. The heptameric pore is robust and remains stable in the denaturing detergent SDS at up to 65◦ C [96]. This stability is the basis of the separation by SDS-PAGE of heteroheptamers, pores containing various combinations of engineered and WT subunits. The availability of pure heteroheptameric species has been a key aspect of our work on the αHL pore. To achieve the electrophoretic separation, the mutant subunits are tagged; originally, the tag was a targeted chemical modiﬁcation with a disulfonate near the C terminus [14], but more recently we have used a genetically encoded oligoaspartate tail [57]. 2.2 Protein Engineering

With the beneﬁt of high-resolution structural information, protein engineering [97] allows substitute amino acid side chains or chemical modiﬁcations to be placed in a protein at deﬁned sites. Engineering studies on membrane proteins have been limited compared to other systems, largely because the proteins can be tricky to handle and because, until quite recently, there has been a lack of structural information. As noted earlier, de-novo design is a difﬁcult task, and very few studies have been conducted on genetically encoded designed membrane proteins. “Redesign” is more readily accomplished, and since 1996 it has been aided in the case of αHL by the availability of a 1.9 Å crystal structure of the heptameric pore. More recently, structural studies of LukF, a related toxin, have provided information about the monomeric form of αHL [98, 99]. Therefore, the impact of structural changes on both the pore and its ability to assemble can be appraised. The αHL pore is a “blank slate” in terms of engineering, and the goal has been to introduce function rather than change function – which has been a difﬁcult task when attempted in proteins more specialized than αHL. In αHL, most desired changes are made on the internal surface of the protein, which is an unusual challenge. Nevertheless, the open structure of the αHL lumen is more readily manipulated than a narrow channel. For example, we have replaced the transmembrane β barrel with a retro (reversed) amino acid sequence and obtained a functional pore [7]. In practice, redesign is done both by rational approaches (e. g., the examination of molecular models) and hit-and-miss methods (e.g., the screening of libraries of mutants). The approaches are often combined. For example, structural studies may suggest a region for intensive mutagenesis. This approach has been applied to the transmembrane region of αHL by using a semi-synthetic gene in which selected

14 Engineered Nanopores

residues are encoded by a replaceable cassette [7]. Nowadays, most forms of sitedirected mutagenesis are greatly accelerated by tools such as the polymerase chain reaction and ligation-free recombination of DNA fragments. Targeted chemical modiﬁcation is an invaluable adjunct to direct mutagenesis. In general, modiﬁcations are carried out at speciﬁc introduced cysteine residues, and in the case of αHL all manner of additions have been made, including the attachment of small molecules with and without poly(ethylene glycol) linkers, peptides, and oligonucleotides [56, 59]. Finally, as noted above, it is possible to make heteromeric αHL pores, which are most commonly molecules with six unaltered subunits and one engineered subunit. 2.3 Electrical Recording

While ensembles of pores can be useful in many circumstances, one notable capability is the means to examine function in intricate detail by single-channel recording. Single-channel recording is also the basis of stochastic sensing. Both monomeric and heptameric αHL pores can assemble or insert into lipid bilayers in a single orientation. Unlike many other pores, the αHL pore remains open indeﬁnitely at moderate transmembrane potentials. Under typical conditions (in 1 M NaCl), the pore has a high conductance of 700 pS, and changes of a few percent in the current carried by a single pore can be measured under optimal conditions. Many measurements can be carried out on a timescale of tens of microseconds. Single-channel recording allows the determination of the basic properties of a nanopore with conﬁdence, including conductance values, ion selectivity, stability, and uniformity. Further, the noncovalent and covalent interactions of exogenous molecules with nanopores can be examined with exquisite sensitivity. 2.4 Other Systems

Analogous methods to those described above will have to be developed for other nanopore systems, if they are to reach the state of sophistication that, in the case of the αHL pore, has been aided by structural studies, protein engineering, and singlechannel recording. It is not at all clear how the structures of many of the alternative systems might be examined at atomic resolution. Nor is it clear how they might be modiﬁed at discrete sites with deﬁned stoichiometry. If the current passing through single nanopores is to be recorded, they will have to be assembled into systems that do not suffer from current leaks and are of low intrinsic capacitance.

3 Outlook

As suggested by the preceding discussion, many improvements might be made to existing nanopore technology.

3 Outlook 15

3.1 Rugged Pores

There are limits to the stability of proteins, and therefore several alternative approaches have been taken to the preparation of nanopores, as described earlier. In the long term, whichever techniques are used must be amenable to mass production and compatible with array technology (see below). Of course, a major advantage of protein nanopores is the ability to carry out functionalization with precision by replacing amino acid side chains or carrying out targeted chemical modiﬁcation. While alternative nanopores, made from polymers, carbon nanotubes, inorganic materials, etc., can be randomly derivatized, it is not yet clear how deﬁned patterns of functionalization or single-site alterations might be made. One compromise would be to immobilize a single adapter molecule (as we have used with the αHL pore) at the entrance of a mechanically and thermally stable nanopore, such as an Au nanotubule. 3.2 Supported Bilayers

To accommodate protein nanopores, it is clear that additional work is needed on supported bilayers and related areas. The formation of defect-free bilayers appears to require ultrasmooth surfaces [86]. “Edge effects” – that is, leaks at the perimeter of the bilayers – must also be tackled. In addition, improved electrical properties demand a substantial reservoir of electrolyte between the bilayer and the support surface [89]. Finally, a reasonable fraction of the incorporated nanopores should be active. Pores with an appreciable extramembranous domain that is located between the bilayer and the underlying surface, might be denatured by the support unless precautions such as polymer cushions are used. In this area, a near-term goal is to observe single channels in supported bilayers [87]. 3.3 Membrane Arrays

Arrays of lipid bilayers would be useful for screening the properties of membrane proteins and for monitoring their interactions with other molecules (Figure 8). The same technology could be used for array-based sensors. Several assays for poreforming molecules have been developed in which liposomes are used in microtiter formats [100–102]. When they are more fully developed, addressable bilayer arrays containing pores will be far more versatile. Several procedures for generating bilayer arrays have been developed, including the formation of corrals by scratching off or blotting the bilayer, or by patterning the support in various ways, for example by depositing the bilayer with a poly-dimethylsiloxane (PDMS) stamp or by making corrals with various barriers including metals, metal oxides, and proteins [103]. Recently, barriers have been formed by the polymerization of diacetylenic lipids [104]. All corrals can be ﬁlled with a single type of bilayer by exposure to a vesicle

16 Engineered Nanopores

Fig. 8 Membrane array on a glass surface. The pattern was formed by photolithography of a glass slide coated with photoresist. Each corral contains a bilayer doped with either 1% NBD-phosphatidyl ethanolamine or 1% Texas Red-phosphatidyl ethanolamine.

A false-color image was generated from epiﬂuorescence images recorded through a home-built macroscope. (E. T. Castellana and P.S. Cremer, Texas AM University, unpublished data.)

suspension, or individual corrals can be ﬁlled with a micropipette [105, 106] or by ﬂow patterning [103]. Two goals of further development are to address each corral electrically and to wash each one individually by using a microﬂuidic system. It should also become possible to arrange other classes of pores, such as those fabricated with ion beams, in addressable formats. 3.4 Alternative Protein Pores

Advances in protein folding and structural biology will drive the search for alternative protein pores. While research on αHL has been extremely fruitful, the pore is a heptamer and therefore intricate to handle. A single-chain, monomeric pore would be ideal, and we have begun to explore the monomeric porin OmpG [107, 108]. It is likely that yet more sophisticated engineering techniques will be applied to protein nanopores than have been in the past. For example, multistep syntheses on the lumen wall might be used to produce complex host molecules within a pore. Because highly stable bilayers with desirable properties have been difﬁcult to obtain, the possibility of directly housing a protein nanopore inside a mechanically stable pore of slightly greater diameter should be considered. The surfaces of both the protein nanopore and the recipient pore (e.g., an Au nanotubule) could be tailored for compatibility (Figure 9).

3 Outlook 17

Fig. 9 Possible means for stabilizing a single protein pore. The nanopore (here the "HL pore) is placed inside a mechanically stable pore (here a Au nanotubule). The surface of the nanopore and the "HL pore are tailored for compatibility. (C. R. Martin, University of Florida.)

3.5 Pores with New Attributes and Applications

Several advanced applications of nanopore engineering are being initiated. For example, the αHL pore can be used as a “nanoreactor” for investigating both noncovalent and covalent interactions. The study of noncovalent interactions elaborates upon the work on sensors. Single-channel current traces contain kinetic information. For example, for a simple binary system, such as a cyclodextrin-guest interaction, kon , koff and Kd can be readily obtained. However, while the values are close to those in bulk solution, further work is required to determine whether there are signiﬁcant differences. Considerations include steric effects, transmembrane and local potentials, solvent conﬁnement, and electroosmosis. In pursuit of ideas for sensing reactive molecules, we have recently begun to explore covalent interactions at the single molecule level with αHL as a nanoreactor, and related considerations come into play here. In addition, it might soon be possible to build various catalytic activities into the pores. This has a precedent in nature, as several enzymes contain a series of active sites located inside an internal channel [109]. Substrates enter the tunnel at one end and products leave at the other. These channels span up to 100 Å, which is the length of the αHL pore. Esterase and ribonuclease activity in a synthetic barrel lined with histidine residues has been observed by Matile and colleagues [110, 111]. By using the variety of substitutions available to protein engineering, and the ability to place them with known geometry, it may be possible to improve turnover and couple catalysis to transmembrane transport.

18 Engineered Nanopores

3.6 Theory

With a continually expanding output of experimental data, much of which is purely descriptive, there is a need for theory and computation in studies of nanopores. Of course, the applications of calculations and simulations to membrane channels and pores is well developed, though far from perfected. By combining aspects of results from structural studies, molecular dynamics simulations, Brownian dynamics and electrodiffusion theory, estimates of fundamental properties of channels and pores have been obtained, such as conductance values, the shape of current-voltage curves, and the magnitude of ion selectivity [112, 113]. The topics covered in this chapter suggest additional areas for exploration such as the properties of water [114] and polymers [48] under conﬁnement, and electroosmosis in pores of nanometer dimensions [115, 118]. It should go without saying that all of this must be coupled to experimentation. Acknowledgments

The authors thank several colleagues for help with the ﬁgures, and acknowledge grants from the U.S. Department of Energy, the Multidisciplinary University Research Initiative (ONR 1999), the National Institutes of Health, the Ofﬁce of Naval Research and DARPA (MOLDICE program). H.B. is the holder of a Royal SocietyWolfson Research Merit Award.

References 1 C. R. MARTIN, P. KOHLI, Nature Rev. Drug Disc. 2003, 2, 29–37. 2 G. E. SCHULZ, Biochim. Biophys. Acta 2002, 1565, 308–317. 3 K. S. ÅKERFELDT, J. D. LEAR, Z. R. WASSERMAN, L. A. CHUNG, W. F. DEGRADO, Acc. Chem. Res. 1993, 26, 191–197. 4 H. BAYLEY, Curr. Opin. Biotechnol. 1999, 10, 94–103. 5 S. LEE, T. KIYOTA, T. KUNITAKE, E. MATSUMOTO, S. YAMASHITA, K. ANZAI, G. SUGIHARA, Biochemistry 1997, 36, 3782–3791. 6 E. MATSUMOTO, T. KIYOTA, S. LEE, G. SUGIHARA, S. YAMASHITA, H. MENO, Y. ASO, H. SAKAMOTO, H. M. ELLERBY, Biopolymers 2001, 56, 96–108. 7 S. CHELEY, O. BRAHA, X. LU, S. CONLAN, H. BAYLEY, Protein Sci. 1999, 8, 1257–1267.

8 A. J. WALLACE, T. J. STILLMAN, A. ATKINS, D. J. JAMIESON, P. A. BULLOUGH, J. GREEN, P. J. ARTYMIUK, Cell 2000, 100, 265–276. 9 B. NORTH, C. M. SUMMA, G. GHIRLANDA, W. F. DEGRADO, J. Mol. Biol. 2001, 311, 1081–1090. 10 G. E. SCHULZ, Curr. Opin. Struct. Biol. 1996, 6, 485–490. 11 M. BANNWARTH, G. E. SCHULZ, Prot. Eng. 2002, 15, 799–804. 12 S. TERRETTAZ, W.-P. ULRICH, H. VOGEL, Q. HONG, L. G. DOVER, J. H. LAKEY, Prot. Sci. 2002, 11, 1917–1925. 13 L. SONG, M. R. HOBAUGH, C. SHUSTAK, S. CHELEY, H. BAYLEY, J. E. GOUAUX, Science 1996, 274, 1859–1865. 14 O. BRAHA, B. WALKER, S. CHELEY, J. J. KASIANOWICZ, L. SONG, J. E. GOUAUX, H. BAYLEY, Chem. Biol. 1997, 4, 497–505. 15 G. MILES, L. MOVILEANU, H. BAYLEY, Protein Sci. 2002, 11, 894–902.

References 16 J. D. HARTGERINK, T. D. CLARK, M. R. GHADIRI, Chem. Eur. J. 1998, 4, 1367–1372. 17 T. D. CLARK, M. R. GHADIRI, J. Am. Chem. Soc. 1995, 117, 12364–12365. 18 J. R. GRANJA, M. R. GHADIRI, J. Am. Chem. Soc. 1994, 116, 10785–10786. 19 T. D. CLARK, L. K. BUEHLER, M. R. GHADIRI, J. Am. Chem. Soc. 1998, 120, 651–656. 20 S. MATILE, Chem. Rec. 2001, 1, 162–172. 21 G. DAS, N. SAKAI, S. MATILE, Chirality 2002, 14, 18–24. 22 G. DAS, S. MATILE, Chirality 2001, 13, 170–176. 23 M. R. GHADIRI, J. R. GRANJA, R. A. MILLIGAN, D. E. MCREE, N. KHAZANOVICH, Nature 1993, 366, 324–327. 24 P. Y. APEL, Y. E. KORCHEV, Z. SIWY, R. SPOHR, M. YOSHIDA, Nucl. Intr. Meth. Physics Res. B 2001, 184, 337–346. 25 C. A. PASTERNAK, G. M. ALDER, P. Y. APEL, C. L. BASHFORD, D. T. EDMONDS, Y. E. KORCHEV, A. A. LEV, G. LOWE, M. MILOVANOVICH, C. W. PITT, T. K. ROSTOVTSEVA, N. I. ZHITARIUK, Radiation Measurements 1995, 25, 675–683. 26 K. B. JIRAGE, J. C. HULTEEN, C. R. MARTIN, Science 1997, 278, 655–658. 27 J. LI, D. STEIN, C. MCMULLAN, D. BRANTON, M. J. AZIZ, J. A. GOLOVCHENKO, Nature 2001, 412, 166–169. 28 R. A. MEHL, J. C. ANDERSON, S. W. SANTORO, L. WANG, A. B. MARTIN, D. S. KING, D. M. HORN, P. G. SCHULTZ, J. Am. Chem. Soc. 2003, 125, 935–939. 29 L.-Q. GU, O. BRAHA, S. CONLAN, S. CHELEY, H. BAYLEY, Nature 1999, 398, 686–690. 30 J. SANCHEZ-QUESADA, M. R. GHADIRI, H. BAYLEY, O. BRAHA, J. Am. Chem. Soc. 2000, 122, 11758–11766. 31 L.-Q. GU, S. CHELEY, H. BAYLEY, Science 2001, 291, 636–640. 32 L.-Q. GU, S. CHELEY, H. BAYLEY, J. Gen. Physiol. 2001, 118, 481–494. 33 J. S´ANCHEZ-QUESADA, M. P. ISLER, M. R. GHADIRI, J. Am. Chem. Soc. 2002, 124, 10004–10005. 34 J. C. HULTEEN, K. B. JIRAGE, C. R. MARTIN, J. Am. Chem. Soc. 1998, 120, 6603– 6604.

35 M. NISHIZAWA, V. P. MENON, C. R. MARTIN, Science 1995, 268, 700–702. 36 S. B. LEE, C. R. MARTIN, J. Am. Chem. Soc. 2002, 124, 11850–11851. 37 S. B. LEE, D. T. MITCHELL, L. TROFIN, T. K. NEVANEN, H. S¨ODERLUND, C. R. MARTIN, Science 2002, 296, 2198–2200. 38 J. M. PASCUAL, A. KARLIN, J. Gen. Physiol. 1998, 111, 717–739. 39 K. SAXENA, V. DROSOU, E. MAIER, R. BENZ, B. LUDWIG, Biochemistry 1999, 38, 2206–2212. 40 N. SAKAI, N. SORDE´ , G. DAS, P. PERROTTET, D. GERARD, S. MATILE, Org. Biomol. Chem. 2003, 1, 1226–1231. 41 L.-Q. GU, M. DALLA SERRA, J. B. VINCENT, G. VIGH, S. CHELEY, O. BRAHA, H. BAYLEY, Proc. Natl. Acad. Sci. USA 2000, 97, 3959–3964. 42 B. B. LAKSHMI, C. R. MARTIN, Nature 1997, 388, 758–760. 43 J. J. KASIANOWICZ, E. BRANDIN, D. BRANTON, D. W. DEAMER, Proc. Natl. Acad. Sci. USA 1996, 93, 13770–13773. 44 L. MOVILEANU, S. CHELEY, S. HOWORKA, O. BRAHA, H. BAYLEY, J. Gen. Physiol. 2001, 117, 239–251. 45 L. MOVILEANU, H. BAYLEY, Proc. Natl. Acad. Sci. USA 2001, 98, 10137– 10141. 46 S. M. BEZRUKOV, J. Membr. Biol. 2000, 174, 1–13. 47 H. BAYLEY, C. R. MARTIN, Chem. Rev. 2000, 100, 2575–2594. 48 M. MUTHUKUMAR, J. Chem. Phys. 2003, 118, 5174–5184. 49 M. ZHOU, J. H. MORAIS-CABRAL, S. MANN, R. MACKINNON, Nature 2001, 411, 643–644. 50 G. A. WOOLLEY, V. ZUNIC, J. KARANICOLAS, A. S. I. JAIKARAN, A. V. STAROSTIN, Biophys. J. 1997, 73, 2465–2475. 51 L. LIEN, D. C. J. JAIKARAN, Z. H. ZHANG, G. A. WOOLLEY, J. Am. Chem. Soc. 1996, 118, 12222–12223. 52 N. K. MAL, M. FUJIWARA, Y. TANAKA, Nature 2003, 421, 350–353. 53 O. BRAHA, L.-Q. GU, L. ZHOU, X. LU, S. CHELEY, H. BAYLEY, Nature Biotechnol. 2000, 17, 1005–1007. 54 S. CHELEY, L.-Q. GU, H. BAYLEY, Chem. Biol. 2002, 9, 829–838.

19

20 Engineered Nanopores

55 L.-Q. GU, H. BAYLEY, Biophys. J. 2000, 79, 1967–1975. 56 S. HOWORKA, L. MOVILEANU, O. BRAHA, H. BAYLEY, Proc. Natl. Acad. Sci. USA 2001, 98, 12996–13001. 57 S. HOWORKA, S. CHELEY, H. BAYLEY, Nature Biotechnol. 2001, 19, 636–639. 58 S. HOWORKA, H. BAYLEY, Biophys. J. 2002, 83, 3202–3210. 59 L. MOVILEANU, S. HOWORKA, O. BRAHA, H. BAYLEY, Nature Biotechnol. 2000, 18, 1091–1095. 60 M. S´ARA, U. SLEYTR, J. Bacteriol. 2000, 182, 859–868. 61 M. J. RUSSO, H. BAYLEY, M. TONER, Nature Biotechnol. 1997, 15, 278–282. 62 J. P. ACKER, X. LU, V. YOUNG, H. CHELEY, H. BAYLEY, A. FOWLER, M. TONER, Biotechnol. Bioeng. 2003, 82, 525–532. 63 A. EROGLU, M. J. RUSSO, R. BIEGANSKI, A. FOWLER, S. CHELEY, H. BAYLEY, M. TONER, Nature Biotechnol. 2000, 18, 163–167. 64 T. CHEN, J. P. ACKER, A. EROGLU, S. CHELEY, H. BAYLEY, A. FOWLER, M. TONER, Cryobiology 2001, 43, 168–181. 65 J. P. ACKER, A. FOWLER, B. LAUMAN, S. CHELEY, M. TONER, Cell Preserv. Technol. 2002, 1, 129–140. 66 S. FERNANDEZ-LOPEZ, H.-S. KIM, E. C. CHOI, M. DELGADO, J. R. GRANJA, A. KHASANOV, A. KRAEHENBUEHL, G. LONG, D. A. WEINBERGER, K. M. WILCOXEN, M. R. GHADIRI, Nature 2001, 412, 452– 455. 67 R. G. PANCHAL, E. CUSACK, S. CHELEY, H. BAYLEY, Nature Biotechnol. 1996, 14, 852–856. 68 H. A. FISHMAN, D. R. GREENWALD, R. N. ZARE, Annu. Rev. Biophys. Biomol. Struct. 1998, 27, 165–198. 69 C. ZIEGLER, W. G¨OPEL, Curr. Opin. Chem. Biol. 1998, 2, 585–591. 70 H. BAYLEY, O. BRAHA, L.-Q. GU, Adv. Mater. 2000, 12, 139–142. 71 H. BAYLEY, P. S. CREMER, Nature 2001, 413, 226–230. 72 S. BHAKDI, U. WELLER, I. WALEV, E. MARTIN, D. JONAS, M. PALMER, Med. Microbiol. Immunol. 1993, 182, 167–175. 73 I. WALEV, S. C. BHAKDI, F. HOFMANN, N. DJONDER, A. VALEVA, K. AKTORIES, S. BHAKDI, Proc. Natl. Acad. Sci. USA 2001, 98, 3185–3190.

74 A. E. OTTO-BRUC, R. N. FARISS, J. P. Van HOOSER, K. PALCZEWSKI, Proc. Natl. Acad. Sci. USA 1998, 95, 15014–15019. 75 X. S. XIE, H. P. LU, J. Biol. Chem. 1999, 274, 15967–15970. 76 L. MOVILEANU, S. CHELEY, H. BAYLEY, Biophys. J. 2003, 85, 897–910. 77 T. LUCHIAN, S.-H. SHIN, H. BAYLEY, Angew. Chem. Int. Ed. 2003, 42, 1926–1929. 78 S.-H. SHIN, T. LUCHIAN, S. CHELEY, O. BRAHA, H. BAYLEY, Angew. Chem. Int. Ed. 2002, 41, 3707–3709. 79 F. S. LIGLER, T. L. FARE, K. D. SEIB, B. SEE, J. W. SMUDA, A. SINGH, P. AHL, M. E. AYERS, A. DALZIEL, P. YAGER, Medical Instrumentation 1988, 22, 247– 256. 80 E. SACKMANN, M. TANAKA, Trends Biotechnol. 2000, 18, 58–64. 81 E.-K. SINNER, W. KNOLL, Curr. Opin. Chem. Biol. 2001, 5, 705–711. 82 J. T. GROVES, Curr. Opin. Drug Discov. Devel. 2002, 5, 606–612. 83 M. C. PETERMAN, J. M. ZIEBARTH, O. BRAHA, H. BAYLEY, H. A. FISHMAN, D. A. BLOOM, Biomed. Microdevices 2002, 4, 231–236. 84 B. SCHUSTER, D. PUM, O. BRAHA, H. BAYLEY, U. B. SLEYTR, Biochim. Biophys. Acta 1998, 1370, 280–288. 85 B. SCHUSTER, U. B. SLEYTR, Biochim. Biophys. Acta 2002, 1563, 29–34. 86 S. M. SCHILLER, R. NAUMANN, K. LOVEJOY, H. KUNZ, W. KNOLL, Angew. Chem. Int. Ed. Engl. 2003, 42, 208–211. 87 G. WIEGAND, K. R. NEUMAIER, E. SACKMANN, Rev. Sci. Instrum. 2000, 71, 2309–2320. 88 B. A. CORNELL, V. L. B. BRAACH-MAKSVYTIS, L. G. KING, P. D. J. OSMAN, B. RAGUSE, L. WIECZOREK, R. J. PACE, Nature 1997, 387, 580–583. 89 G. KRISHNA, J. SCHULTE, B. A. CORNELL, R. J. PACE, P. D. OSMAN, Langmuir 2003, 19, 2294–2305. 90 W. MEIER, C. NARDIN, M. WINTERHALTER, Angew. Chem. Int. Ed. Engl. 2000, 39, 4599–4602. 91 A. VALEVA, A. WEISSER, B. WALKER, M. KEHOE, H. BAYLEY, S. BHAKDI, M. PALMER, EMBO J. 1996, 15, 1857– 1864.

References 92 B. J. WALKER, M. KRISHNASASTRY, L. ZORN, J. J. KASIANOWICZ, H. BAYLEY, J. Biol. Chem. 1992, 267, 10902–10909. 93 T. LAMLA, W. STIEGE, V. A. ERDMANN, Mol. Cell. Proteomics 2002, 1.6, 466–472. 94 A. HILDEBRAND, M. POHL, S. BHAKDI, J. Biol. Chem. 1991, 266, 17195–17200. 95 G. MENESTRINA, J. Membr. Biol. 1986, 90, 177–190. 96 B. WALKER, H. BAYLEY, Protein Eng. 1995, 8, 491–495. 97 W. F. Ed. DEGRADO, Chem. Rev. 2001, 101, special issue. 98 R. OLSON, H. NARIYA, K. YOKOTA, Y. KAMIO, E. GOUAUX, Nature Struct. Biol. 1999, 6, 134–140. 99 J.-D. P´EDELACQ, L. MAVEYRAUD, G. PRE´ VOST, L. BABA-MOUSSA, A. GONZA´ LEZ, E. COURCELLE, W. SHEPARD, H. MONTEIL, J.-P. SAMAMA, L. MOUREY, Structure 1999, 7, 277–288. 100 S. KOLUSHEVA, L. BOYER, R. JELINEK, Nature Biotechnol. 2000, 18, 225–227. 101 J. M. RAUSCH, W. C. WIMLEY, Anal. Biochem. 2001, 293, 258–263. 102 G. DAS, P. TALUKDAR, S. MATILE, Science 2002, 298, 1600–1602. 103 J. T. GROVES, S. G. BOXER, Acc. Chem. Res. 2002, 35, 149–157. 104 K. MORIGAKI, T. BAUMGART, A. OFFENHA¨ USSER, W. KNOLL, Angew. Chem. Int. Ed. Engl. 2001, 40, 172–174. 105 P. S. CREMER, T. YANG, J. Am. Chem. Soc. 1999, 121, 8130–8131. 106 T. YANG, E. E. SIMANEK, P. S. CREMER, Anal. Chem. 2000, 72, 2587–2589.

107 S. CONLAN, Y. ZHANG, S. CHELEY, H. BAYLEY, Biochemistry 2000, 39, 11845–11854. 108 M. BEHLAU, D. J. MILLS, H. QUADER, W. K¨UHLBRANDT, J. VONCK, J. Mol. Biol. 2001, 305, 71–77. 109 X. HUANG, H. M. HOLDEN, F. M. RAUSHEL, Annu. Rev. Biochem. 2001, 70, 149–180. 110 B. BAUMEISTER, N. SAKAI, S. MATILE, Org. Lett. 2001, 3, 4229–4232. 111 B. BAUMEISTER, S. MATILE, Macromolecules 2002, 35, 1549–1555. 112 M. S. P. SANSOM, I. H. SHRIVASTAVA, K. M. RANATUNGA, G. R. SMITH, Trends Biochem. Sci. 2000, 25, 368– 374. 113 B. ROUX, Curr. Opin. Struct. Biol. 2002, 12, 182–189. 114 U. RAVIV, P. LAURAT, J. KLEIN, Nature 2001, 413, 51–54. 115 A. T. CONLISK, J. MCFERRAN, Z. ZHENG, D. HANSFORD, Anal. Chem. 2002, 74, 2139–2150. 116 G. G. KOCHENDOERFER, J. M. TACK, S. CRESSMAN, Bioconjug. Chem. 2002, 13, 474–480. 117 S. W. COWAN, R. M. GARAVITO, J. N. JANSONIUS, J. A. JENKINS, R. KARLSSON, N. K¨ONIG, E. F. PAI, R. A. PAUPTIT, P. J. RIZKALLAH, J. P. ROSENBUSCH, G. RUMMEL, T. SCHIRMER, Structure 1995, 3, 1041–1050 118 L.-Q. GU, S. CHELEY, H. BAYLEY, Proc. Natl. Acad. Sci. USA, 2003, 100, 15498–15503

21

1

Design and Stability of α-Helices

Andrew J. Doig, Neil Errington, and Teuku M. Iqbalsyah UMIST, Manchester, United Kingdom

Originally published in: Protein Folding Handbook. Part I. Edited by Johannes Buchner and Thomas Kiefhaber. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30784-2

1 Introduction

Proteins are built of regular local folds of the polypeptide chain called secondary structure. The α-helix was ﬁrst described by Pauling, Corey, and Branson in 1950 [1], and their model was quickly supported by X-ray analysis of hemoglobin [2]. Irrefutable proof of the existence of the α-helix came with the ﬁrst protein crystal structure of myoglobin, where most secondary structure is helical [3]. α-Helices were subsequently found in nearly all globular proteins. It is the most abundant secondary structure, with ≈30% of residues found in α-helices [4]. In this review, we discuss structural features of the helix and their study in peptides. Some earlier reviews on this ﬁeld are in Refs [5–11].

2 Structure of the α-Helix

A helix combines a linear translation with an orthogonal circular rotation. In the α-helix the linear translation is a rise of 5.4 Å per turn of the helix and a circular rotation is 3.6 residues per turn. Side chains spaced i, i + 3, i, i + 4, and i, i + 7 are therefore close in space and interactions between them can affect helix stability. Spacings of i, i + 2, i, i + 5, and i, i + 6 place the side chain pairs on opposite faces of the helix, avoiding any interaction. The helix is primarily stabilized by i, i + 4 hydrogen bonds between backbone amide groups. The conformation of a polypeptide can be described by the backbone dihedral angles φ and ψ. Most φ, ψ combinations are sterically excluded, leaving only the broad β region and narrower α region. One reason why the α-helix is so stable Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Design and Stability of "-Helices

is that a succession of the sterically allowed α, φ, ψ angles naturally position the backbone NH and CO groups towards each other for hydrogen bond formation. It is possible that a succession of the most stable conformation of an isolated residue in a polymer with alternative functional groups could point two hydrogen bond donors or acceptors towards each other, making secondary structure formation unfavourable. One reason why polypeptides may have been selected as the polymer of choice for building functional molecules is that the sterically most stable conformations also give strong hydrogen bonds. The residues at the N-terminus of the α-helix are called N -N-cap-N1-N2-N3N4 etc., where the N-cap is the residue with nonhelical φ, ψ angles immediately preceding the N-terminus of an α-helix and N1 is the ﬁrst residue with helical , ψ angles [12]. The C-terminal residues are similarly called C4-C3-C2-C1-C-cap-C etc. The N1, N2, N3, C1, C2, and C3 residues are unique because their amide groups participate in i, i + 4 backbone-backbone hydrogen bonds using either only their CO (at the N-terminus) or NH (at the C-terminus) groups. The need for these groups to form hydrogen bonds has powerful effects on helix structure and stability [13]. The following rules are generally observed for N-capping in α-helices: Thr and Ser N-cap side chains adopt the gauche− rotamer, hydrogen bond to the N3 NH and have ψ restricted to 164 ± 8◦ . Asp and Asn N-cap side chains either adopt the gauche− rotamer and hydrogen bond to the N3 NH with ψ = 172 ± 10◦ , or adopt the trans rotamer and hydrogen bond to both the N2 and N3 NH groups with ψ = 107 ± 19◦ . With all other N-caps, the side chain is found in the gauche+ rotamer so that the side chain does not interact unfavorably with the N-terminus by blocking solvation and ψ is unrestricted. An i, i + 3 hydrogen bond from N3 NH to the N-cap backbone C=O is more likely to form at the N-terminus when an unfavorable N-cap is present [14]. Bonds between sp3 hybridized atoms show preferences for dihedral angles of +60◦ (gauche− ), −60◦ (gauche+ ) or 180◦ (trans). Within amino acid side chains the preferences for these rotamers varies between secondary structures [15–18]. Within the α-helix, rotamer preferences vary greatly between different positions of N-cap, N1, N2, N3, and interior [14, 19]. At helix interior positions, the gauche+ rotamer is usually most abundant (64%), followed by trans (33%) with gauche− very rare (3%), though considerable variations are seen. For example, Ser and Thr are gauche− 19% and 14% of the time, respectively, as their hydroxyl groups can hydrogen bond to the helix backbone in this conformation. The β-branched side chains Val, Ile, and Thr are the most restricted with gauche+ very common. 2.1 Capping Motifs

The amide NH groups at the helix N-terminus are satisﬁed predominantly by side chain hydrogen bond acceptors. In contrast, carbonyl CO groups at the C-terminus are satisﬁed primarily by backbone NH groups from the sequence following the helix [13]. The presence of such interactions would therefore stabilize helices. A well-known interaction is helix capping, deﬁned as speciﬁc patterns found at or

2 Structure of the "-Helix Table 1 The most common capping motifs at α-helix termini

Sequence Related position Designation N-capping p-XXp

N-cap → N3

hp-XXph

N → N4

Expanded capping box/Hydrophobic staple

hP-XXVh

N → N4

Pro-box motif

C-capping hXp-XGh

C3 → C

Schellman motif

hXp-XGp

C2 → C

α L motifs

X-Pro

C-cap → C

Capping box

Pro-capping motif

Interactions

References

[24, 25] Reciprocal H-bonds between residues at N-cap and N3, where S/T at N-cap and E at N3 are the most frequently observed residues Reciprocal H-bonds between [26, 250] residues at N-cap and N3 accompanied by hydrophobic interactions between residues at N and N4 [28] N-cap Pro residues are usually associated with Ile and Leu at N , Val at N3 and a hydrophobic residue at N4 H-bonds between CNH and the C3 CO at and between C NH and C2 CO, respectively accompanied by hydrophobic interaction between C3 and C As The Schellman motif but H-bond between C NH and C3 CO where Cpolar A stabilizing electrostatic interaction of the residues at positions C-cap and Pro at C with the helix macrodipole

[31, 223]

[32]

[34]

p, polar amino acids; h,hydrophobic amino acids;X,any amino acids; P, proline; G, glycine.

near the ends of helices [12, 20–24]. The patterns involve hydrogen bonding and hydrophobic interactions as summarized in Table 1. A common pattern of capping at the helix N-terminus is the capping box. Here, the side chain of the N-cap forms a hydrogen bond with the backbone of N3 and, reciprocally, the side chain of N3 forms a hydrogen bond with the backbone of the N-cap [25]. The deﬁnition of the capping box was expanded by Seale et al. [26] to include an associated hydrophobic interaction between residues N and N4 and is also known as a “hydrophobic staple” [27]. A variant of the capping box motif, termed the “big” box with an observed hydrophobic interaction between nonpolar side chain groups in residues N4 and N (not N ) [26]. The Pro-box motif involves three hydrophobic residues and a Pro residue at the N-cap [28]. The N-cap Pro residue is usually associated with Ile or Leu at position N, Val at position N3, and a hydrophobic residue at position N4. This motif is arguably very destabilizing since it discourages helix elongation due to the low

3

4 Design and Stability of "-Helices

N-cap preference of Pro [29] and the poor helix propagation parameter value of Val [30]. This suggests that the Pro-box motif will not specially contribute to protein stability but to the speciﬁcity of its fold. The two primary capping motifs found at helix C-termini are the Schellman and the α L motifs [31–33]. The Schellman motif is deﬁned by a doubly hydrogenbonded pattern between backbone partners, consisting of hydrogen bonds between the amide NH at C and the carbonyl CO at C3 and between the amide NH at C and the carbonyl CO at C2, respectively. The associated hydrophobic interaction is between C3 and C. In a Schellman motif, polar residues are highly favored at the C1 position and the C residue is typically glycine. If C is polar, the alternative α L motif is observed. The α L motif is deﬁned by a hydrogen bond between the amide NH at C and the carbonyl CO at C3. As in the Schellman motif, the C residue is typically glycine, which adopts a positive value of dihedral angle. However, the hydrophobic interaction in an α L is heterogeneous, occurring between C3 and any of several residues external to the helix (C3 , C4 , or C5 ) [32]. A statistical analysis of the protein crystal structures database detected another possible C-cap local motif, designated Pro C-capping [34]. Certain combinations of X-Pro pairs, in which residue X is the C-cap and the Pro is at position C , are more abundant than expected. The aliphatic (Ile, Val, and Leu) and aromatic residues (Phe, Tyr, and Trp), together with Asn, His, and Cys are the most favored residues accompanying Pro at position C . A notable difference between the N- and C-terminal motifs is that at the Nterminus, helix geometry favors side chain-to-backbone hydrogen bonding and selects for compatible polar residues [14, 19]. Accordingly, the N-terminus promotes selectivity in all polar positions, especially N-cap and N3 in the capping box. In contrast, at the C-terminus, side chain-to-backbone hydrogen bonding is disfavored. Backbone hydrogen bonds are satisﬁed instead by posthelical backbone groups. The C-terminus need only select for C residues that can adopt positive values of the backbone dihedral angle , most notably Gly [32]. 2.2 Metal Binding

In general, the stabilization of folded forms can be achieved by reducing the entropy by cross-linking, such as via metal ions. Another way to stabilize helix conformations, especially in short peptides, is to introduce an artiﬁcial nucleation site composed of a few residues ﬁxed in a helical conformation. For example, the calcium-binding loop from EF-hand proteins saturated with a lanthanide ion promotes a rigid short helical conformation at its C-terminus region [35]. Metal ions play a key structural and functional role in many proteins. There is therefore interest in using peptide models containing metal complexes. In stabilizing proteins, the rule of thumb in metal–ligand binding is that “hard metals” prefer “hard ligands”. For example Ca and Mg prefer ligands with oxygen as the coordinating atoms (Asp, Glu) [36]. In contrast, soft metals, such as Cu and Zn,

2 Structure of the "-Helix

bind mostly to His, Cys, and Trp ligands and sometimes indirectly via water molecules [37]. In studies of helical peptide models, soft ligands have been mostly used. In the presence of Cd ions, a synthetic peptide containing Cys-His ligands i, i + 4 apart at the C-terminal region promoted helicity from 54% to 90%. The helicity of a similar peptide containing His-His ligands increased by up to 90% as a result of Cu and Zn binding [38]. The addition of a cis-Ru(III) ion to a 6-mer peptide, Ac-AHAAAHANH2 , changed the peptide conformation from random coil to 37% helix [39]. An 11-residue peptide was converted from random coil to 80% helix content by the addition of Cd ions, although the ligands used were not natural amino acids but aminodiacetic acids [40]. As(III) stabilizes helices when bound to Cys side chains spaced i, i + 4 by −0.7 to −1.0 kcal mol−1 [41]. 2.3 The 310 -Helix

310 -Helices are stabilized by i, i + 3 hydrogen bonds instead of the i, i + 4 bonds found in α-helices, making the cylinder of the 310 -helix narrower than α and the hydrogen bonds nonlinear. About 3–4% of residues in crystal structures are in 310 -helices [4, 42], making the 310 -helix the fourth most common type of secondary structure in proteins after α-helices, β-sheets, and reverse turns. Most 310 -helices are short, only 3 or 4 residues long, compared with a mean of 10 residues in α-helices [4]. 310 -Helices are commonly found as N- or C-terminal extensions to an α-helix [4, 43, 44]: Strong amino acid preferences have been observed for different locations within the interiors [42] and N- and C-caps [14] of 310 -helices in crystal structures. The 310 -helix is being recognized as of increasing importance in isolated peptides and as a possible intermediate in α-helix formation [45, 46]. It has been proposed that peptides made of the standard 20 L-amino acids can form 310 -helices [47], or at least populate a signiﬁcant amount of 310 -helix at their N- and C-termini. The fraction of 310 -helix present in a helical peptide is strongly sequence dependent [48, 49]. 310 -helix formation can be induced by the introduction of a Cα,α -disubstituted α-amino acid, of which α-aminoisobutyric acid (AIB) is the prototype. For most amino acids, the α-helical geometry (φ = −57◦ , ψ −70◦ ) is of lower energy than the 310 geometry (φ = −49◦ , ψ = −26◦ ). There is no disallowed region between the α and 310 conformations in the Ramachandran plot, and a peptide can therefore be gradually transformed from one helix to the other [50]. 2.4 The p -Helix

In contrast to the widely occurring α- and 310 -helices, the π-helix is extremely rare. The π-helix is unfavorable for three reasons: its dihedral angles are energetically unfavorable relative to the α-helix [51, 52], its three-dimensional structure has a 1Å hole down the center that is too narrow for access by a water molecule, resulting in the loss of van der Waals interactions, and a higher number of residues (four) must

5

6 Design and Stability of "-Helices

be correctly oriented before the ﬁrst i, i + 5 hydrogen bond is formed, making helix initiation more entropically unfavorable than for α- or 310 -helices. The π-helix may be of more than theoretical interest [53–57].

3 Design of Peptide Helices

The ﬁrst protein crystal structures showed an abundance of α-helices, leading to speculation whether peptide fragments of the helical sequences could be stable in isolation. Since isolated helices lack hydrophobic cores, this question could be rephrased to ask whether the amide-amide hydrogen bond is strong enough to oppose the loss of conformational entropy arising from restricting the peptide into a helical structure [58]. Early estimates of these terms suggested that hydrogen bonds were not strong enough and this was conﬁrmed by peptide studies of helices from myoglobin [59] and staphylococcal nuclease [60], which found no helix formation. Early estimates of helix/coil parameters from a host–guest system (see Section 3.1) suggested that polypeptides would need to be hundreds of residues long to form stable helices. Hence, the earliest work on peptide helices was on long homopolymers of Glu or Lys, which show coil-to-helix transitions on changing the pH from charged to neutral. The neutral polypeptides are metastable and prone to aggregation, ultimately to β-sheet amyloid [61]. A conﬂicting result was found by Brown and Klee in 1971 [62], who reported that the C-peptide of ribonuclease A, which contains the ﬁrst 13 residues of the protein and which forms a helix in the protein, was helical at 0◦ C. This observation was not followed up for 10 years until extensive work on the sequence features responsible for helix formation in this peptide, and in the larger S-peptide, was performed by Baldwin and coworkers [63–73], while NMR studies by Rico and coworkers precisely deﬁned the helical structure [74–82]. Some important features responsible for the helicity of the C-peptide that emerged from this work included an i, i + 4 Phe-His interaction, an i, i + 3 Glu-His salt bridge, an unusual i, i + 8 Glu-Arg salt bridge that spanned two turns of the helix and a helix termination signal at Met. The stabilizing effects of salt bridges were inferred from pH titrations and circular dichroism (CD), where a decrease in helicity was seen when a residue participating in a salt bridge was neutralized. Quantiﬁcation of these features in terms of a free energy was not possible at that time, as helix/coil theory that included side-chain interactions and termination signals (caps) had not been developed. Perhaps the most interesting result from the C- and S-peptide work was the importance of Ala. The replacement of interior helical residues with Ala was stabilizing, indicating that a major reason why this helix was folded in isolation was the presence of three successive alanines from positions 4–6. This led to the successful design of isolated, monomeric helical peptides in aqueous solution, ﬁrst containing several salt bridges and a high alanine content, based on (EAAAK)n [83, 84] and then a simple sequence with a high alanine content solubilized by several lysines [85]. These “AK peptides” are based on the sequence (AAKAA)n , where n is typically

3 Design of Peptide Helices 7

2–5. The Lys side chains are spaced i, i + 5 so they are on opposite faces of the helix, giving no charge repulsion and may be substituted with Arg or Gln to give a neutral peptide. Hundreds of AK peptides have been studied, giving most of the results on helix stability in peptides (see below). The Alanines in the (EAAAK)n type peptides may be removed entirely; E4 K4 peptides, with sequences based on (EEEEKKKK)n or EAK patterns are also helical, stabilized by large numbers of salt bridges [8, 86–88]. 3.1 Host–Guest Studies

Extensive work from the Scheraga group has obtained helix/coil parameters using a host–guest method. Long random copolymers were synthesized of a water-soluble, nonionic guest (poly[N 5 -(3-hydroxypropyl)-L-glutamine] (PHPG) or poly[N 5 -(4-hydroxybutyl)-L-glutamine] (PHBG)), together with a low (10–50%) content of the guest residue. Using the s and σ Zimm-Bragg helix/coil parameters (see below) for the host homopolymer, it was possible to calculate those for the guest using helix/coil theory as a function of temperature. The results from the host–guest work are in disagreement with most of those from short peptides of ﬁxed sequence (see below). 3.2 Helix Lengths

Helix formation in peptides is cooperative, with a nucleation penalty. Helix stability therefore tends to increase with length, in homopolymers at least. As the length of a homopolymer increases, the mean fraction helix will level off below 100% as long helices tend to break in two. In heteropolymers, observed lengths are highly sequence dependent. As helices are at best marginally stable in monomeric peptides in aqueous solution, they are readily terminated by the introduction of a strong capping residue or a residue with a low intrinsic helical preference. The length distribution of helices in peptides is very different to proteins [4]. Most helices are short, with 5–14 residues most common. There is a general trend for a decrease in frequency as the length increases beyond 13 residues. Helix lengths longer than 25 are rare. This is a consequence of the organization of proteins into domains of similar size, rather than showing different rules for stability; helices do not extend beyond the boundaries of the domain and so terminate. There is also a preference to have close to an integral number of turns so that their N- and C-caps are on the same side of the helix [89]. 3.3 The Helix Dipole

The secondary amide group in a protein backbone is polarized with the oxygen negatively charged and hydrogen positively charged. In a helix the amides are

8 Design and Stability of "-Helices

all oriented in the same direction with the positive hydrogens pointing to the N-terminus and negative oxygens pointing to the C-terminus. This can be regarded as giving a positive charge at the helix N-terminus and a negative charge at the helix C-terminus [90–92]. In general, therefore, negatively charged groups are stabilizing at the N-terminus and positive at the C-terminus, as shown by numerous titrations that measure helix content as a function of pH and amino acid preferences for helix terminal positions. An alternative interpretation of these results is that favored side chains are those that can make hydrogen bonds to the free amide NH groups at N1, N2, and N3 or free CO groups at C1, C2, and C3 [93]. Charged groups can form stronger hydrogen bonds than neutral groups, thus providing an alternative rationalization of the pH titration results. These hypotheses are not mutually exclusive, as a charged side chain can also function as a hydrogen bond acceptor or donor. A free energy simulation of Tidor [94] suggested that helix-stabilizing interactions, as a result of a Tyr → Asp substitution at an N-cap site, arose from hydrogen-bonding interactions from its direct hydrogen-bonding partners, and from more distant electrostatic interactions with groups within the ﬁrst two turns of the helix. Measurements of the amino acid preferences for the N-cap, N1, N2, and N3 positions in the helix allow a comparison to be made of the relative importance of helix-dipole and hydrogen-bonding interactions [29, 95–97]. The helix-dipole model implies that the side chains most favored at the helix N-terminus are those with a negative charge while positive charges are disfavored. A pure hydrogen-bonding model implies that favored side chains are those that can make hydrogen bonds to the free amide NH groups at N1, N2, and N3. In general, the N-cap results suggest that hydrogen bonding is more important than helix-dipole interactions; the best N-caps are Asn, Asp, Ser, and Thr [29], which can accept hydrogen bonds from the N2 and N3 NH groups [14]. Glu has only a moderate N-cap preference despite its negative charge. In contrast, N1, N2, and N3 results suggest that helix-dipole interactions are more important. The contrasting results between the different helix N-terminal positions can be rationalized by considering the geometry of the hydrogen bonds. N-cap hydrogen bonds are close to linear [14] and so are strong, while N1 and N2 hydrogen bonds are close to 90◦ [19], making them much weaker. Helix-dipole effects are likely to be present at all sites, as also shown by every pH titration where a more negative side chain is favored over a more positive side chain, but this can be overwhelmed by strong hydrogen bonds, as at the N-cap. In the absence of strong hydrogen bonds, helix-dipole effects dominate. Hydrogen bonds can therefore make a substantial contribution to protein stability, but only if their geometry is close to linear (as it is for backbone to backbone hydrogen bonds in α-helices and β-sheets). 3.4 Acetylation and Amidation

A simple, yet effective, way to increase the helicity of a peptide is to acetylate its N-terminus [22, 98]. This is readily done with acetic anhydride/pyridine, after completion of a peptide synthesis, but before cleavage of the peptide from the resin

3 Design of Peptide Helices 9

and deprotection of the side chains. Acetylation removes the positive charge that is present at the helix terminus at low or neutral pH; this would interact unfavorably with the positive helix dipole and free N-terminal NH groups. The extra CO group from the acetyl group can form an additional hydrogen bond to the NH group, putting the acetyl at the N-cap position. This has a strong stabilizing effect by approximately 1.0 kcal mol−1 compared with alanine [29, 30, 99]. The acetyl group is one of the best N-caps. We found only Asn and Asp to be more stabilizing [29]. Amidation of the peptide C-terminus is achieved by using different types of resin in solid-phase peptide synthesis, resulting in the replacement of COO− with CONH2 . This is similar structurally to acetylation: the helix is extended by one hydrogen bond and an unfavorable charge–charge repulsion with the helix dipole is removed. The energetic beneﬁt of amidation is rather smaller, however, with the amide group being no better than Ala and in the middle if the C-cap residues are ranked in order of stabilization effect [29]. As most helical peptides studied to date are both acetylated and amidated, and acetylation is more stabilizing than amidation, the helicity of peptides is generally skewed so that residues near the N-terminus are more helical than those near the C-terminus. Acetylation and amidation are often also beneﬁcial in removing a pH titration that can obscure other effects. 3.5 Side Chain Spacings

Side chains in the helix are spaced at 3.6 residues per turn of the helix. Side chains spaced at i, i + 3, i, i + 4, and i, i + 7 are therefore close in space and interactions between them can affect helix stability. Spacings of i, i + 2, i,i + 5, and i, i + 6 place the side chain pairs on opposite faces of the helix, avoiding any interaction. Care must therefore be taken to avoid unwanted interactions when designing helices. For example, we studied these peptides to determine the energetics of the Phe-Met interaction [100]: F7M12 Ac-YGAAKAFAAKAMAAKAA-NH2 F8M12 Ac-YGAAKAAFAKAMAAKAA-NH2 The control peptide F7M12 has Phe and Met spaced i, i + 5 where they cannot interact; F8M12 moves the Phe one place towards the C-terminus so that Phe and Met are spaced i, i + 4 where they can form a noncovalent interaction. The increase in helix content was used to derive the Phe-Met interaction energy. These sequences are not ideal, however. F7M12 contains a Phe-Lys i, i + 3 interaction that is replaced by a Lys-Phe i, i + 3 interaction in F8M12. If either of these are stabilizing or nonstabilizing the Phe-Met i, i + 4 energy will be in error. While this problem is likely to be small, in this case at least, our more recent designs try to minimize such potential problems. For example, the following peptides, designed to measure Ile-Lys i, i + 4 energies, do not introduce or remove an unwanted i, i + 3 spacing [101]: IKc Ac-AKIAAAAKIAAAAKAKAGY-NH2 IKi Ac-AKAIAAAKAIAAAKAKAGY-NH2

10 Design and Stability of "-Helices

3.6 Solubility

Lack of peptide solubility can be a problem. Peptides designed to be helices can even become highly insoluble amyloid with a high β-sheet content. We have occasionally found that alanine-based peptides designed to be helical instead form amyloid, though this may only take place after several years storage in solution. The measurement of interactions in a helix will be compromised by peptide oligomerization so it is generally essential to check that the peptides are monomeric. This can be done rigorously by sedimentation equilibrium, which determines the oligomeric state of a molecule in solution. This is difﬁcult, however, with the short peptides often used, as their molecular weights are at the lower limit for this technique. A simpler method is to check a spectroscopic technique that depends on peptide structure, most obviously CD, as a function of concentration. If the signal depends linearly on peptide concentration across a large range, including that used to study the peptide structure, it is safe to assume that the peptide is monomeric. For example, if helicity measurements are made at 10 µM, CD spectra can be acquired from 5 µM to 100 µM. An oligomer that does not change state, such as a coiled-coil, across the concentration range cannot be excluded, however. Light scattering can detect aggregation. A monomeric peptide should have a ﬂat baseline in a UV spectrum outside the range of any chromophores in the peptide. In stock solutions of a peptide with a single tyrosine isolated from the helix region by Gly should have A300 /A275 < 0.02 and A250 < A275 < 0.2 [102]. Consideration of solubility is essential when designing helical peptides. While Ala has the highest helix propensity and would provide an ideal theoretical background for substitutions, poly(Ala) is insoluble. Solubility can be achieved most easily by including polar side chains spaced i, i + 5 in the sequence where they cannot interact. Lys, Arg, and Gln are used most often for this purpose. Gln may be preferred if unwanted interactions with charged Lys or Arg may be a problem, but some AQ peptides lack sufﬁcient solubility and AQ peptides are less helical. We have found that it is not easy to predict peptide solubility. For example, 20 peptides with sequence Ac-XAAAAQAAAAQAAGY-NH2, where X is all the amino acids encoded by the genetic code, were all soluble [95]. However, when the homologous peptide Ac-AAAAAAQAAAAQAAAAQGY-NH2 was synthesized to study the N2 position it tended to aggregate. The Gln side chains were therefore replaced with Lys, giving no further problems [96]. The spacing of side chains in the helix is best visualized with a helical wheel, to ensure that the designed helix does not have a nonpolar face that may lead to dimerization. The following web pages provide useful resources for this: http://www.site.uottawa.ca/∼turcotte/resources/HelixWheel/ http://marqusee9.berkeley.edu/kael/helical.htm

3 Design of Peptide Helices 11

3.7 Concentration Determination

An accurate measurement of helix content depends on an accurate spectroscopic measurement and, equally importantly, peptide concentration. This is usually achieved by including a Tyr side chain at one end of the peptide. The extinction coefﬁcient of Tyr at 275 nm is 1450 M−1 cm−1 [103]. If Trp is present, measurements at 281 nm can be used where the extinction coefﬁcient of Trp is 5690 M−1 cm−1 and Tyr 1250 M−1 cm−1 [104]. Phe absorbance is negligible at this wavelength. These UV absorbances are ideally made in 6 M GuHCl, pH 7.0, 25 ◦ C, though we have found very little variation with solvent so measurements in water are identical within error. The main source of error is in pipetting small volumes; this is typically around 2%. Pipetting larger volumes with well-maintained ﬁxed volume pipettors can help minimize this error, which may well be the largest when measuring a percentage helix content by CD. Though the inclusion of aromatic residues is required for concentration determination, this can have the unwanted side effect of perturbing a CD spectrum, leading to an inaccurate measure of helix content. A simple solution to this is to separate the terminal Tyr from the rest of the sequence by one or more Gly residues [105]. If the aromatic residues must be included within the helical region, the CD spectrum should be corrected to remove this perturbation (see later). 3.8 Design of Peptides to Measure Helix Parameters

Numerous peptides have been studied to measure the forces responsible for helix formation (see below). In general, one or more peptides are synthesized that contain the interaction of interest, while all other terms that can contribute to helix stability are known. The helix content of the peptide is measured and the statistical weight of the interaction is varied until predictions from helix/coil theory match experiment. The weight of the interaction (and hence its free energy, as -RT ln(weight)) is then known. In practice it is wise to also synthesize a control peptide that lacks the interaction, but is otherwise very similar. The helix content of this peptide should be predictable from helix/coil theory using known parameters. For example, Table 2 gives sequences of control and interaction peptides used to measure Phe-Lys i, i + 4 side-chain interaction energies [106]. The FK control peptide has no i, i + 4 side-chain interactions and all the helix/coil weights for these residues are known. The helicity predicted by SCINT2 [30] for this peptide is in close agreement with experiment. This is an important conﬁrmation that the helix/coil model and the parameters used are accurate. The discrepancy between prediction and experiment for the FK interaction peptide is because this peptide contains two Phe-Lys i, i + 4 side-chain interactions whose weights are assumed to be 1 (i.e., G = 0). If the Phe-Lys weight is changed to 1.3, the prediction agrees with experiment. Hence G (Phe-Lys) = - RT ln 1.3 = −0.14 kcal mol−1 .

12 Design and Stability of "-Helices

Table 2 Peptides designed to measure the energetics of the Phe-Lys interaction

Peptide name

Sequence

FK control Ac-AAAKFAAAAKFAAAAKAKAGY-NH2 FK interaction Ac-AAAKAFAAAKAFAAAKAKAGY-NH2

Predicted nelicity Phe-Lys weight G (Phe-Lys) Experimental helicity (no side-chain interaction) to ﬁt experiment kcal mol−1 35.4% 41.1%

35.6 34.6

N/A 1.3

N/A −0.14

3 Design of Peptide Helices 13

It is important to minimize the error in determining helix/coil parameters. Errors can be calculated by assuming an error in measurement of % helix (±3% is reasonable) and reﬁtting the results across this experimental error range. For the case above, G (Phe-Lys) was calculated for 38.1% and 44.1% helix, giving a range of −0.05 to −0.18 kcal mol−1 . This is a remarkably small error range, resulting from the high sensitivity of helix content to a small change in helix stability. This sensitivity can be maximized by including multiple identical interactions; here the FK interaction peptide has two Phe-Lys interactions, for example. The helix contents of the control and interaction peptides should be close to 50%. This may be difﬁcult to achieve if the residues being used have low helix preferences. The best way to maximize sensitivity is to calculate it in advance using possible sequences, helix/coil theory, an error of ±3% and guessing a sensible value for the interaction energy. The interaction of interest can be placed at various positions in the peptide, its length can be changed by adding further AAKAA sequences, terminal residues such as a Tyr can be moved, and solubilizing side chains can be changed. Peptide design can be lengthy, considering sensitivity and solubility, but is a valuable process. The complex nature of the helix/coil equilibrium with frayed conformations highly populated means that considerable variations can be seen for apparently small changes, such as moving an interaction towards one terminus of a sequence.

3.9 Helix Templates

A major penalty to helix formation is the loss of entropy arising from the requirement to ﬁx three consecutive residues to form the ﬁrst hydrogen bond of the helix. Following this nucleation, propagation is much more favored as only a single residue need be restricted to form each additional hydrogen bond. A way to avoid this barrier is to synthesize a template molecule that facilitates helix initiation, by ﬁxing hydrogen bond acceptors or donors in the correct orientation for a peptide to bond in a helical geometry. The ideal template nucleates a helix with an identical geometry to a real helix. Kemp’s group applied this strategy and synthesized a proline-like template that nucleated helices when a peptide chain was covalently attached to a carboxyl group (Figure 1a) [107–111]. The template is in an equilibrium between cis and trans isomers of the proline-like part of the molecule. Only the trans isomer can bond to the helix so helix content is determined by measuring the cis/trans ratio by NMR. Bartlett et al. reported on a hexahydroindol-4-one template (Figure 1b) [112] that induced 49–77% helicity at 0 ◦ C, depending on the method of determination, in an appended hexameric peptide. Several other templates were less successful and could only induce helicity in organic solvents [113–115]. Their syntheses are often lengthy and difﬁcult, partly due to the challenging requirement of orienting several dipoles to act as hydrogen bond acceptors or donors.

14 Design and Stability of "-Helices

Fig. 1 Helix templates. a) Kemp, b) Bartlett.

3.10 Design of 310 -Helices

Peptides can be induced to form 310 -helix by the incorporation of a signiﬁcant amount of disubstituted Cα amino acids, of which the simplest is α-aminoisobutyric acid (Aib) [116–119]. The presence of steric interactions from the two methyl groups on the α-carbon in AIB results in the 310 geometry being energetically favored over the α. Peptides rich in Cα, α-disubstituted α-amino acids are readily crystallized and many of their structures have been solved (reviewed in Refs [50] and [120]). Aib is conformationally restricted so that shorter Aib-based helices are more stable than Ala-based α-helices [121]. Many examples of helix stabilization or 310 -helix formation by Aib have been reported. The α/310 equilibrium has been studied in peptides. Yokum et al. [122] synthesized a peptide composed of Cα, α-disubstituted α-amino acids that forms mixed 310 -/α-helices in mixed aqueous/organic solvents. Millhauser and coworkers have studied the 310 /α equilibrium in peptides by electron spin resonance (ESR) and nuclear magnetic resonance (NMR) [47, 49, 123, 124] and argued that mixed 310 -/α-helices are common in polyalanine-based helices. Yoder et al. [125] studied a series of L-(αMe)-Val homopolymers and showed that their peptides can form coil, 310 -helix, or α-helix depending on concentration, peptide length, and solvent. Kennedy et al. [126] used Fourier transform infrared (FTIR) spectroscopy to monitor 310 - and α-helix formation in poly(Aib)-based peptides. Hungerford et al. [127] synthesized peptides that showed an α to 310 transition upon heating. 310 -Helices have also been proposed as thermodynamic intermediates where helical peptides of moderate stability exist as a mixture of α- and 310 -structures [45, 128].

4 Helix Coil Theory

Guidelines for the inclusion of the 20 natural amino acids in 310 -helices can be taken from their propensities in proteins, both at interior positions [42] and at N-caps [14]. These are similar, but not identical to α preferences. Side-chain interactions in the 310 -helix are spaced i, i + 3, with i, i + 4 on opposite sides of the helix. This offers scope to preferentially stabilize 310 over α, by including stabilizing i, i + 3 interactions and i, i + 3 repulsions. As the 310 - and α-helix structures are so similar, with only a small change in backbone dihedral angle, peptides designed to form 310 -helix are likely to form a mixture of 310 and α, with a central α segment and 310 at the helix termini common. This complex equilibrium, and the spectro-scopic similarity of α and 310 makes the analysis of these peptides difﬁcult. 3.11 Design of p -helices

To our knowledge, no peptide has yet been made that forms π-helix. There are 4.4 side chains per turn of the π-helix, so i, i + 5 interactions may weakly stabilize πhelix over α. Given the other strongly destabilizing features of the π-helix, however, we doubt whether this effect will ever be strong enough. We expect that no peptide composed of the 20 natural amino acids will ever fold to a monomeric π-helix in aqueous solution. We are willing to back up this prediction, but in view of the out-come of the Paracelsus challenge [129, 130], we are taking the advice of George Rose [131] and offering only an “I Met the π-helix Challenge” T-shirt as a prize.

4 Helix Coil Theory

Peptides that form helices in solution do not show a simple two-state equilibrium between a fully folded and fully unfolded structure. Instead they form a complex mixture of all helix, all coil or, most frequently, central helices with frayed coil ends. In order to interpret experiments on helical peptides and make theoretical predictions on helices it is therefore essential to use a helix/coil theory that considers every possible location of a helix within a sequence. The ﬁrst wave of work on helix/coil theory was in the late 1950s and early 1960s and this has been reviewed in detail by Poland and Scheraga [132]. In 1992, Qian and Schellman [133] reviewed current understanding of helix/coil theories. Our review covered the extensive development of helix/coil theory since this date [134]. The simplest way to analyze the helix/coil equilibrium, still occasionally seen, is the two-state model where the equilibrium is assumed to be between a 100% helix conformation and 100% coil. This is incorrect and its use gives serious errors. This is because helical peptides are generally most often found in partly helical conformations, often with a central helix and frayed, disordered ends, rather than in the fully folded or fully unfolded states.

15

16 Design and Stability of "-Helices

Fig. 2 Zimm-Bragg and Lifson-Roig codes and weights for the "-helix.

4.1 Zimm-Bragg Model

The two major types of helix/coil model are (i) those that count hydrogen bonds, principally Zimm-Bragg (ZB) [135] and (ii) those that consider residue conformations, principally Lifson-Roig [136]. In the ZB theory the units being considered are peptide groups and they are classiﬁed on the basis of whether their NH groups participate in hydrogen bonds within the helix. The ZB coding is shown in Figure 2. A unit is given a code of 1 (e.g., peptide unit 5 in Figure 2) if its NH group forms a hydrogen bond and 0 otherwise. The ﬁrst hydrogen-bonded unit proceeding from the N-terminus has a statistical weight of σ s, successive hydrogen bonded units have weights of s and nonhydrogen-bonded units have weights of 1. The s-value is a propagation parameter and σ is an initiation parameter. The most fundamental feature of the thermodynamics of the helix/coil transition is that the initiation of a new helix is much more difﬁcult than the propagation of an existing helix. This is because three residues need to be ﬁxed in a helical geometry to form the ﬁrst hydrogen bond while adding an additional hydrogen bond to an existing helix require that only one residue is ﬁxed. These properties are thus captured in the ZB model by having σ smaller than s. The statistical weight of a homopolymeric helix of a N hydrogen bonds is σ sN−1 . The cost of initiation, σ , is thus paid only once for each helix while extending the helix simply multiplies its weight by one additional s-value for each extra hydrogen bond. 4.2 Lifson-Roig Model

In the LR model each residue is assigned a conformation of helix (h) or coil (c), depending on whether it has helical , ψ angles. Every conformation of a peptide of N residues can therefore be written as a string of N c’s or h’s, giving 2N conformations in total. Residues are assigned statistical weights depending on their conformations and the conformations of surrounding residues. A residue in an h conformation with an h on either side has a weight of w. This can be thought of as an equilibrium constant between the helix interior and the coil. Coil residues are used as a reference and have a weight of 1. In order to form an i, i + 4 hydrogen bond in a helix three successive residues need to be ﬁxed in a helical

4 Helix Coil Theory

conformation. M consecutive helical residues will therefore have M–2 hydrogen bonds. The two residues at the helix termini (i.e., those in the center of chh or hhc conformations) are therefore assigned weights of v (Figure 2). The ratio of w to v gives the approximately the effect of hydrogen bonding (1.7:0.036 for Ala [30] or - RT ln(1.7/0.036) = −2.1 kcal mol−1 ). A helical homopolymer segment of M residues has a weight of v2 wM-2 and a population in the equilibrium of v2 wM-2 divided by the sum of the weights of every conformation (i.e., the partition function). In this way the population of every conformation is calculated and all properties of the helix/coil equilibrium evaluated. The LR model is easier to handle conceptually for heteropolymers since the w and v parameters are assigned to individual residues. The substitution of one amino acid at a certain position thus changes the w- and vvalues at that position. In the ZB model the initiation parameter σ is associated with several residues and s with a peptide group, rather than a residue. It is therefore easier to use the LR model when making substitutions. Indeed, most recent work has been based on this model. A further difference is that the ZB model assigns weights of zero to all conformations that contain a chc or chhc sequence. This excludes a very large number of conformations that contain a residue with helical φ,ψ angles but with no hydrogen bond. In LR theory, these are all considered. The ZB and LR weights are related by the following formulae [133]: s = w/(1 + v); σ = v2 /(1 + v)4 . The complete helix/coil equilibrium is handled by determining the statistical weight for every possible conformation that contains a helix plus a reference weight of 1 for the coil conformation. Each conformation considered in the helix/coil equilibrium is given a statistical weight. This indicates the stability of that conformation, with the higher the weight, the more probable the conformation. Weights are deﬁned relative to the all coil conformation, which is given a weight of 1. The statistical weight of a conformation can thus be regarded as an equilibrium constant relative to the coil; a weight >1 indicates the conformation is more stable than coil, <1 means less stable and 1 means equally stable. The population of each conformation is given by the statistical weight of that conformation divided by the sum of the statistical weights for every conformation (i.e., the partition function). Thus the greater the statistical weight, the more stable the conformation. The key to using helix/coil theory is the partition function. All the properties of a system at equilibrium are contained within the partition function, which makes it very valuable. Partition functions are extremely powerful concepts in statistical thermodynamics since they include all properties of an equilibrium. Any property of the equilibrium can be extracted from the partition function by applying the appropriate mathematical function. This is analogous to quantum mechanics where any property can be determined from a system by applying an operator function to a wavefunction. In this case the properties could be the mean number of hydrogen bonds, the mean helix length, the probability that each residue is within a helix etc. In particular, the mean number of residues with a weight x is given by ∂ ln Z/∂ ln x. Circular dichroism is commonly used to give the mean helix content of a helical peptide, namely the fraction of residues that have a weight of w. LR-based models can thus be related to experimental data by equating the measured mean

17

18 Design and Stability of "-Helices

helix content to (∂ ln Z/∂ ln x)/N, where N is the number of residues in the peptide. Statistical weights can be regarded as equilibrium constants for the equilibrium between coil and the structure (as the reference coil weight is deﬁned as 1). They can therefore be converted to free energies as - RT ln (weight). Equations for partition functions are determined as follows: Write a table of the conformations and assigned statistical weights for residues. For the Lifson-Roig model, the weights of a residue depend on its conformation and the weights of its two neighboring residues as follows: Conformation Weight of central residue in triplet ccc cch chc chh hcc hch hhc hhh

1 1 v v 1 1 v w

The same information is rewritten in the form of a matrix (M):

h h¯ M = h c¯ c h¯ c c¯

¯ hc ¯ c¯h c¯c hh   wv00 0 0 1 1   v v 0 0 0 011

For each triplet, the state of the left-most residue is shown at the start of each row in the matrix. The state of the right-most residue in each triplet is shown at the end of the top of each column. The state of the center residue of each triplet is shown as the barred residue in both the rows and columns. When the states of the center residues differ (i.e., one is c while the other is h), the entry in the matrix is zero. Otherwise, the matrix gives the weight, taken from the table. This apparently simple change in presentation now (amazingly) allows the generation of the partition function (Z) for a polypeptide of N residues as:   0  1  Z = ( 0 0 1 1 )M N  1 1 The end vectors are present to ensure that the ﬁrst and last residues in the peptide cannot have a w-weighting. A w-weighting indicates that the residue is between two residues that hydrogen bond to each other and this is impossible for terminal residues. Further extensions to helix/coil theory are dealt with in the same way, by deﬁning residue weight assignments, rewriting as a matrix and determining end

4 Helix Coil Theory

vectors. The work may be made easier by combining any identical columns. For example, the Lifson-Roig matrix above has identical third and fourth columns so can be simpliﬁed to a 3 × 3 matrix:

M = h h¯

h c¯ c(h∪c)

¯ hc ¯ c¯(h ∪ c) hh   wv0 0 0 1 v v0

Here, h ∪ c shows the residues being combined. The matrix entries for the combined positions are the entries for the noncombined entries, with zero being discounted if combined with a nonzero entry. New end vectors are now required, as their lengths must be the same as the order of the square matrix. 4.3 The Unfolded State and Polyproline II Helix

The treatment of peptide conformations is based on Flory’s isolated-pair hypothesis [137]. This states that while and ψ for a residue are strongly interdependent, giving preferred areas in a Ramachandran plot, each , ψ pair is independent of the , ψ angles of its neighbors. Pappu et al. examined the isolated-pair hypothesis in detail by exhaustively enumerating the conformations of poly(Ala) chains [138]. Each residue was considered to populate 14 mesostates, deﬁned by ranges of , ψ values. By considering all 14 N mesostate strings, all conformations were considered for up to seven alanines. The number of allowed conformations was found to be considerably fewer than the maximum, thus showing that the isolatedpair hypothesis is invalid. The chains mostly populate extended or helical conformations as many partly helical conformations are sterically disallowed. Such effects are not included in helix/coil theories, thus presenting a considerable challenge for the future. Helix/coil theories assign the same weight (1) to every coil residue; steric exclusion means that these should vary and be lower than 1 in many cases. The polyproline II helix may well be an important conformation for unfolded proteins [139–148]. In particular, denatured alanine-rich peptides may form polyproline II helix [143, 148, 149]. It may therefore be valid to consider residues in helical peptides to be in three possible states (helix, coil, or polyproline II), rather than two (helix or coil). No current helix/coil model takes this into account. A scale of amino acid preferences for the polyproline II helix has recently been published [150]. 4.4 Single Sequence Approximation

Since helix nucleation is difﬁcult, conformations with multiple helical segments are expected to be rare in short peptides. In the one-, or single-, helical sequence approximation, peptide conformations containing more than one helical segment are assumed not to be populated and are excluded from the partition function (i.e.,

19

20 Design and Stability of "-Helices

assigned statistical weights of zero). As peptide length increases, the approximation is no longer valid since multiple helical segments can be long enough to overcome the initiation penalty. The single sequence approximation will also break down when a sequence with a high preference for a helix terminus is within the middle of the chain. The error from using the single sequence approximation will therefore show a wide variation with sequence and could be potentially serious if a sequence has a high preference to populate more than one helix simultaneously. Conformations with two or more helices may also often include helix-helix tertiary interactions that are ignored in all helix/coil models. 4.5 N- and C-Caps

In the original LR model weights are assigned to residues in the center of hhh triplets (a weight of w for propagation) or in the center of chh or hhc triplets (a weight of v for initiation). Residues in all other triplets have weights of 1. LR-based models have been extended by assigning weights to additional conformations. Ncapping can therefore be included in LR theory be assigning a weight of n to the central residue in a cch triplet [99]. Similarly, the C-cap is the ﬁrst residue in a nonhelical conformation (c) at the C-terminus of a helix. C-cap weights (cvalues) are assigned to central residues in hcc triplets. Application of the model to experimental data where the C-terminal amino acid of a helical peptide was varied, allowed the determination of the c-values and hence free energies of C-capping (as -RT ln c) of all the amino acids [29]. A problem with the original deﬁnitions of the capping weights above is that they apply to isolated h or hh conformations that are best regarded as part of the random coil. A helical hydrogen bond can only form when a minimum of three consecutive h residues are present. Andersen and Tong [151] and Rohl et al. [9] therefore changed the deﬁnition of the N-cap to apply only to the c residue in a chhh quartet. The N-cap residues in chc or chhc conformations have weights of zero in the Andersen-Tong model and 1 in the Rohl et al. model. 4.6 Capping Boxes

The N-terminal capping box [25] includes a side chain-backbone hydrogen bond from N3 to the N-cap (i, i−3). This is included in the LR model by assigning a weight of w*r to the chhh conformation, where r is the weight for the Ser backbone to Glu side chain bond [9]. 4.7 Side-chain Interactions

As helices have 3.6 residues per turn, side chains spaced i, i + 3 or i, i + 4 are close in space. Side-chain interactions are thus possible when four or ﬁve consecutive

4 Helix Coil Theory

residues are in a helix. They are included in the LR-based model by giving a weight of w*q to hhhh quartets and w*p to hhhhh quintets. The side-chain interaction is between the ﬁrst and last side chains in these groups; the w weight is maintained to keep the equivalence between the number of residues with a w weighting and the number of backbone helix hydrogen bonds [100]. Scholtz et al. [152] used a model based on the one-helical-sequence approximation of the LR model to quantitatively analyze salt bridge interactions in alanine-based peptides. Only a single interaction between residues of any spacing was considered, though this was appropriate for the sequences they studied. Shalongo and Stellwagen [153] also proposed incorporating side-chain interaction energies into the LR model, using a clever recursive algorithm. 4.8 N1, N2, and N3 Preferences

The helix N-terminus shows signiﬁcantly different residue frequencies for the N-cap, N1, N2, N3, and helix interior positions [12, 19, 154, 155]. A complete theory for the helix should therefore include distinct preferences for the N1, N2, and N3 positions. In the original LR model, the N1 and C1 residues are both assigned the same weight, v. Shalongo and Stellwagen [153] separated these as vN and vC . Andersen and Tong [151] did the same and derived complete scales for these parameters from ﬁtting experimental data, though some values were tentative. The helix initiation penalty is vN *vC and so vN and vC values are all small (≈ 0.04). We added weights for the N1, N2, and N3 (n1, n2, and n3) positions as follows [156]: The n1 value is assigned to a helical residue immediately following a coil residue. The penalty for helix initiation is now n1.v, instead of v2 , as v remains the C1 weight. An N2 helical residue is assigned a weight of n2.w, instead of w. The weight w is maintained in order to keep the useful deﬁnition of the number of residues with a w weighting being equal to the number of residues with an i, i + 4 main chain-main chain hydrogen bond. The n2 value is an adjustment to the weight of an N2 residue that takes into account the structures that can be adopted by side chains uniquely at this position. Similarly, an N3 residue is now assigned the weight n3.w, instead of w. 4.9 Helix Dipole

Helix-dipole effects were added to the LR model by Scholtz et al. [152], though they used the one-sequence approximation so that only one or no dipoles in total are present. In LR models helix-dipole effects are subsumed within other energies. For example, N-cap, N1, N2, and N3 energies will include a contribution from the helixdipole interaction so the energy of interaction of charged groups at this position with the dipole should not be counted in addition.

21

22 Design and Stability of "-Helices

4.10 310 - and p -Helices

The Lifson-Roig formalism can easily be adapted to describe helices of other cooperative lengths [157]. The fundamental difference between a 310 -helix and an α-helix is that the 310 -helix has an i, i + 3 hydrogen-bonding pattern rather than the i, i + 4 pattern characteristic of the α-helix. For a given number of units in helical conformations, a 310 -helix will consequently have one more hydrogen bond than an α-helix. To include this difference in the 3 10 -helix theory, one of the αhelical-initiating residues (i.e., the central unit of either the hα hα c or the chα hα triplet) must become a 310 -helix-propagating residue. We arbitrarily chose to assign the propagating statistical weight, wτ , to the central unit of the hτ hτ c triplet such that helix propagating unit i is associated with the hydrogen bond formed between the CO of peptide i−2 and the NH of peptide i+1. The remainder of the statistical weights applicable to the α-helix/coil theory are maintained. The models described above for the α-helix/coil and 310 -helix/coil transitions can be combined to describe an equilibrium including pure α-helices, pure 310 -helices, and mixed α-/310 -helices [157]. In this model, three conformational states are possible, 310 -helical (hτ ), α-helical (hα ), and coil (c). The pure 310 -helix and the mixed α-/310 -helix models were subsequently extended to include side-chain interactions [158]. Sheinerman and Brooks [128] independently produced a model for the α/310 /coil equilibrium, based on the ZB formalism, rather than the LR model. They similarly extend the classiﬁcation of conformations from α/coil to α/310 /coil. In a π-helix, formation of an i, i + 5 hydrogen bond requires that four units be constrained to the π-helical conformation, hπ . The π subscript designates the conformation and weights describing the π-helix, whose dihedral angles are distinct from α- and 310 -helices. Assigning statistical weights to individual units requires consideration of the conformations of the unit itself and its three nearest neighbors [157]. The initiating statistical weight, vπ , is assigned to a helical unit when one or more of its two N-terminal and nearest C-terminal neighbors are in the coil conformation. The deﬁnition of helix-initiating units as the two N-terminal and one C-terminal units of each helical stretch is again arbitrary. Units in a π-helical conformation with three helical neighbors are assigned the propagating statistical weight, wπ . A π-helix propagating residue, i, is thus associated with the hydrogen bond between the NH of residue i + 2 and the CO of residue i −3. 4.11 AGADIR

AGADIR is an LR-based helix/coil model developed by Serrano, Mu˜ noz and coworkers. The original model [159] included parameters for helix propensities excluding backbone hydrogen bonds (attributed to conformational entropy), backbone hydrogen bond enthalpy, side-chain interactions, and a term for coil weights at the end of helical sequences (i.e., caps). The single-sequence approximation was used. The original partition function assumed that many helical conformations did not exist, as all conformations in which the residue of interest is not part of a helix were

4 Helix Coil Theory

excluded [100, 159]. These were corrected in a later version, AGADIRms, which considers all possible conformations [160]. If AGADIR and LR models are both applied to the same data, to determine a side-chain interaction energy, for example, the results are similar, showing that the models are now not signiﬁcantly different [160, 161]. The treatment of the helix/coil equilibrium differs in a number of respects from the ZB and LR models and these have been discussed in detail in by Mu˜ noz and Serrano [160]. The minimal helix length in AGADIR is four residues in an h conformation, rather than three. The effect of this assumption is to exclude all helices which contain a single hydrogen bond; only helices with two or more hydrogen bonds are allowed. In practice, this probably makes little difference as chhhc conformations are usually unfavorable and hence have low populations. Early versions of AGADIR considered that residues following an acetyl at the N-terminus or preceding an amide at the C-terminus were always in a c conformation; this was changed to allow these to be helical [162]. The latest version of AGADIR, AGADIR1s-2 [162], includes terms for electrostatics [162], the helix dipole [162, 163], pH dependence [163], temperature [163], ionic strength [162], N1, N2, and N3 preferences [164], and capping motifs such as the capping box, hydrophobic staple, Schellman motif, and Pro-capping motif [162]. The free energy of a helical segment, Ghelical-segment , is given by Ghelical-segment + GInt + GHbond + GSD + Gdipole +GnonH + Gelectrost , which are terms for the energy required to ﬁx a residue in helical angles (with separate terms for N1, N2, N3, and N4), backbone hydrogen bonding, side-chain interactions excluding those between charged groups, capping, and helix-dipole interactions, respectively. Electrostatic interactions are calculated with Coulomb’s equation. Helix-dipole interactions were all electrostatic interactions between the helix dipole or free N- and Ctermini and groups in the helix. Interactions of the helix dipole with charged groups located outside the helical segment were also included. pH-dependence calculations considered a different parameter set for charged and uncharged side chains and their pK a values. The single sequence approximation (see above) is used again, unlike in AGADIRms. This means that it must not be used for full protein sequences, though this has been done, even if they do not have any tertiary interactions. AGADIR is at present the only model that can give a prediction of helix content for any peptide sequence, thus making it very useful. It can also predict NMR chemical shifts and coupling constants. In order to do this it must include estimates of all the terms that contribute to helix stability, notably the 400 possible i, i + 4 side-chain interactions. Since only a few of these interactions have been measured accurately, the terms used cannot be precise. Further determination of energetic contributions to helix stability is therefore still needed. 4.12 Lomize-Mosberg Model

Lomize and Mosberg also developed a thermodynamic model for calculating the stability of helices in solution [165]. Interestingly, they extended it to consider helices in micelles or a uniform nonpolar droplet to model a protein core environment. Helix stability in water is calculated as the sum of main chain interactions, which is the

23

24 Design and Stability of "-Helices

free energy change for transferring Ala from coil to helix, the difference in energy when replacing an Ala with another residue, hydrogen bonding and electrostatic interactions between polar side chains and hydrophobic side-chain interactions. An entropic nucleation penalty of two residues per helix is included. Different energies are included for N-cap, N1–N3, C1–C3, C-cap, hydrophobic staples, Schellman motifs, and polar side-chain interactions, based on known empirical data at the time (1996). Hydrophobic interactions were calculated from decreases in nonpolar surface area when they are brought in contact. Helix stability in micelles or nonpolar droplets are found by calculating the stability in water then adding a transfer energy to the nonpolar environment. 4.13 Extension of the Zimm-Bragg Model

Following the discovery of short peptides that form isolated helices in aqueous solution, V´asquez and Scheraga extended the ZB model to include helix-dipole and side-chain interactions [166]. The model is very general as it can include interactions of any spacing within a single helix. It was applied to determine i, i + 4 and i, i + 8 interactions. Long-range interactions, beyond the scope of LR models, can thus be included. Roberts [167] and Gans et al. [88] also reﬁned the ZB model to include side-chain interactions. 4.14 Availability of Helix/Coil Programs

Some Lifson-Roig based helix/coil models are available at http://www.bi. umist.ac.uk/users/mjfajdg/HC.htm. We recommend SCINT2 for peptides with side-chain interactions (freely available from us). If no side-chain interactions are present, CAPHELIX is identical and simpler to run. If N1, N2, or N3 preferences are important in your sequence N1N2N3 can be used. If the peptide contains side-chain interactions that have not been well characterized (which is the case for the great majority of sequences) AGADIR can be run at http://www. embl-heidelberg.de/Services/serrano/agadir/agadir-start.html. In our experience, SCINT2 or CAPHE-LIX are more accurate at predicting helix contents of alaninebased control peptides than AGADIR (see Refs [101] and [106] for examples), though AGADIR can be applied to many more sequences. The Andersen version of an LRbased helix/coil is at http://faculty.washington.edu/nielshan/helix/. 5 Forces Affecting α-helix Stability 5.1 Helix Interior

Since the advent of crystal structures of proteins it had been noticed that some amino acids appeared frequently in α-helices and others less frequently [168, 169].

5 Forces Affecting "-helix Stability 25

For example alanine and leucine are abundant, whereas proline and glycine appear rarely. As more of this kind of information became available a helix propensity scale was derived [170] and eventually allowed prediction of the location of α-helices (and other structures) in folded proteins from their sequence [171]. Different approaches have been used in order to determine the helical propensity or preference of individual amino acids. Scheraga and coworkers used a host–guest strategy (see Section 3.1) to derive values for the helical preference of various amino acid residues [172, 173]. This has been carried out for all 20 naturally occurring amino acids [174]. This work has been criticized as the host side chains can interact with each other [175]. The introduction of a guest residue thus removes host–host interactions and replaces them with PHBG–guest or PHPG– guest side-chain interactions that may obscure the intrinsic helix propensities. We believe that helix propensities are best evaluated in an alanine background, where no side-chain interactions can affect the helix stability. Rohl et al. [30] used many alanine-based peptides with the general sequences Ac(AAKAA)m Y-NH2 (or with Q instead of K) to measure interior helix propensities. Substitutions in the helix interior and subsequent measures of helicity using CD spectroscopy in both water and 40% (v/v) triﬂuoroethanol (TFE) allowed both the calculation of the Lifson-Roig w parameter and stabilization energy for all 20 amino acids (see Table 3). Kallenbach and coworkers also used synthetic peptides of the form succinyl-YSEEEEKAKKAXAEEAEKKKK-NH2 where substitutions at X

Table 3 Helix propagation propensities and free energies of amino acids in water

Amino acid Ala Arg+ Leu Met Lys+ Gln Glu Ile Trp Ser Tyr Phe Val His Asn Thr Cys Asp Gly Pro From Ref. [30].

G◦ (helix) (kcal mol−1 )

w value

−0.27 −0.052 0.095 0.25 0.019 0.28 0.21 0.44 0.69 0.52 0.42 0.73 0.77 0.57 0.69 0.95 0.64 0.52 1.7 >3.8

1.70 1.14 0.87 0.65 1.00 0.62 0.70 0.46 0.29 0.40 0.48 0.27 0.25 0.36 0.29 0.18 0.32 0.40 0.048 <0.001

26 Design and Stability of "-Helices

allowed determination of helix stabilising energies for common amino acids [87]. Stellwagen and coworkers made substitutions in position 9 of Ac-Y(EAAAK)3 ANH2 [84]. These agree well with the alanine-based peptide work described previously [30, 176]. Other groups have investigated helical propensities and stabilization using whole protein methods. Blaber et al. [177–179] used mutagenesis in the helices of phage T4 lysozyme to study the structural effects of substitutions of amino acids. With the exception of substituting proline, they found that no substitutions signiﬁcantly distorted the helix backbone. G values correlated well (71–93%) with model peptide studies and with studies on the frequency of amino acid occurrence in protein structures [171]. Fersht and coworkers used a similar method with barnase to study the effect of replacing Ala32 of the second helix in this protein with the other 19 naturally common amino acids [180]. They used reversible urea denaturation to measure free energies of unfolding. O’Neil and DeGrado [181] used substitutions into an α-helical two-stranded coiled-coil system to deduce helix-forming tendencies of common amino acids, through the design of a peptide that forms a noncovalent α-helical dimer, which is in equilibrium with a randomly coiled monomeric state. The α-helices in the dimer contain a single solvent-exposed site that is surrounded by small, neutral amino acid side chains. Each of the commonly occurring amino acids was substituted into this guest site, and the resulting equilibrium constants for the monomer– dimer equilibrium were determined to provide a list of free energy difference (G◦ ) values. Again these values show good agreement with those of other groups working in different model systems. In 1998 Pace and Scholtz [182] gathered information from many different sources and derived a scale for the propensity of each amino acid in the helix interior. This is summarized in Table 4. The values are in (G) relative to alanine as zero. Alanine was taken as zero as it is generally (though not universally) agreed that this amino acid has the highest helical propensity. Shown for comparison in Table 5 are the s values from the Zimm-Bragg model as derived by the Scheraga group [174]. Both tables are ranked in terms of descending helical propensity. As can be seen from Tables 4 and 5, the two series do not agree on even the relative helical propensities of the amino acid residues. The data summarized by Pace and Scholtz shows alanine having the highest helix propensity and all other residues having lower values (a positive (G) value relative to Ala). Both series show proline and glycine to have the lowest helical propensity (highest (G) value or lowest s value. The most controversial of these differences over the years has been that of alanine. Host–guest analysis showing alanine to be effectively helix-neutral has been supported by data from some other groups, notably the templated helices of Kemp and coworkers [107]. Several efforts have been made to try to explain this discrepancy, including implication of the charged groups used to solubilize the alanine-based peptides [183, 184], but little reconciliation has been achieved. The use of template-nucleated helices has been criticized by Rohl et al. [185] who argued that the low apparent helix propensity of alanine is a consequence of properties of the template–helix junction. However, recent work by Kemp and

5 Forces Affecting "-helix Stability 27 Table 4 Summary of other experimental helix propensities (relative to alanine)

Amino acid Ala Arg+ Leu Met Lys+ Gln Glu Ile Trp Ser Tyr Phe Val His Asn Thr Cys Asp Gly Pro

Helix propensity (G)(kcal mol−1 ) 0.00 0.21 0.21 0.24 0.26 0.39 0.40 0.41 0.49 0.50 0.53 0.54 0.61 0.61 0.65 0.66 0.68 0.69 1.00 3.16

From Ref. [182]. Table 5 Helical propensity values from host–guest studies

Amino acid Glu Met Leu Ile Trp Phe Ala Arg+ Tyr Cys Gln Val Lys+ His Thr Asn Asp Ser Gly Pro From Ref. [174].

s (20◦ C) 1.35 1.2 1.14 1.14 1.11 1.09 1.07 1.03 1.02 0.99 0.98 0.95 0.94 0.85 0.82 0.78 0.78 0.76 0.59 0.19

28 Design and Stability of "-Helices

coworkers [186] may at last settle the controversy. Using templates to investigate the helix-forming tendency of polyalanine, these workers extended the length of the polyalanine beyond the previous limit of six residues. Below six residues, both this group and Scheraga’s had low helix propensities for alanine (see above) but when the limit of six was exceeded, a dramatic increase in helix propensity was observed. For chains with less than six alanines, w = 1:03, in agreement with both Kemp and Scheraga’s earlier experimental results. For chains with 6–9 alanines, w = 1:15 and for more than 10 Alanines, w is 1.26. This indicates that there is a length-dependent term in the helicity of polyalanine and that the charged groups are not having the effect previously ascribed to them. These values for longer polyalanine sequences are also much more consistent with values published by other groups. 5.2 Caps

Serrano and Fersht [20] explored the capping preferences at the N-cap by mutating Thr residues at the N-cap of two helices in barnase. They found that negatively charged residues were favored at the N-cap with a rank order of Asp > Thr > Glu > Ser > Asn > Gly > Gln > Ala > Val. Interestingly, their result conﬂicted with the statistical survey result that Asn is one of the most frequently found N-caps in proteins [12]. Experimentally Asn destabilized the helices by 1.3 kcal mol−1 relative to Thr. The rank order of amino acids N-cap preferences in T4 lysozyme was found to be Thr > Ser > Asn > Asp > Val = Ala > Gly [21]. They suggested that Asn can be inherently as good an N-cap as Ser or Thr, but it requires a change in backbone dihedral angles of N-cap residues, which might be altered in native proteins as the results of tertiary contacts. Indeed Asn is the most stabilizing residue at N-cap in a peptide model in the absence of tertiary contacts and other side-chain interactions (see below). The Kallenbach group [187] substituted several amino acids at the N-cap position in peptide models in the presence of a capping box. They found that Ser and Arg are the most stabilizing residues with G relative to Ala of −0.74 and −0.58 kcal mol−1 , respectively, whilst Gly and Ala are less stabilizing. The results are in agreement with the results of Forood et al. [23], who found that the trend in α-helix inducing ability at the N-cap is Asp > Asn > Ser > Glu > Gln > Ala. A more comprehensive work to determine the preferences for all 20 amino acids at the N-cap position used peptides with a sequence of NH2 -XAKAAAAKAAAAKAAGYCONH2 [22, 29, 99]. N-Capping free energies ranged from Asn (best) to Gln (worst) (Table 6). We have used a similar approach using peptide models to probe the preferences at N1 [95], N2 [96], and N3 [97] using peptides with sequences of CH3 COXAAAA-QAAAAQAAGY-CONH2 , CH3 CO-AXAAAAKAAAAKAAGY-CONH2 and CH3 CO-AAXAAAAKAAAAKAGY-CONH2 , respectively. The results have given N1, N2 and N3 preferences for most amino acids for these positions (Table 6) and these agree well with preferences seen in protein structures, with the interesting exception of Pro at N1. Petukhov et al. similarly obtained N1, N2, and N3

5 Forces Affecting "-helix Stability 29 Table 6 Amino acid propensities at N- and C-terminal positions of the helix

G relative to Ala for transition from coil to the position (kcal mol−1 )

Residue

A C◦ C− D◦ D− E◦ E− F G H◦ H+ I K+ L M N P Q R+ S T V W Y

N-cap

N1

29

95 164;188

0

0

−1.4 1.0 0.5 −1.6 0 1.0 −0.7 0.1 −0.7 1.4 −1.2 1.0 −0.7 0.7 −0.5 0.1 −0.7 −0.3 −1.7 −0.4 2.5 −0.1 −1.2 −0.7 −0.1 −1.3 −0.9

0.5 0.7 0.4 0.5 0.6 0.5 0.7 0.4 0.5 0.6 0.4

N2

0

96

N3

C3

164;188 97 164;188 189 0

0

0.9 0.7 −0.2 −0.2 −0.4 0.9 0.7

0

0

0

C2 190 0

C1 C-cap 189 189 0

0

29

0.6 1.3 0.4

0 0.2

0

0.2

0.3

−0.4 −0.5

0.3

0.8

2.1

0.6 1.0 0.6

0.4

0.5

0.2

0.2 0.4

0.5

0.5

0.1 0.1 0.7 0.4

−0.1 −0.1 −0.3 0.3 0.1

0.1

0.2 0.1 0.6 0.5 0.3 0.4 0.5

0.5 0.8 0.7 0.5 0.5 0.8

0.1 −1.1

2.6 −0.2

0.6 0.9 0.5 0.7 1.7

251

1.1

0.8 0.4

C

0.5 0.5 0.3 0.7

0.7 0.9 0.8 0.7

0.4 0.4 0.7

0.3

1.2

0.2

0.5 0.5 0.4

1.1 1.2

0.6 0.6 0.4

4.0 1.2

0.2 −0.02 0.2 0.05 −0.5 −0.4 0.6 0.5 0.7 0.5 0.8 0.8 0.6 0.5 0.8 0.3 0.4 0.7 0.6 0.9

−0.9 1.5 −0.1 0.1 −0.4 1.2 −0.1 −0.2 0.3 1.1 1.6 0.7

−2.2

NH2 −XAKAAAAKAAAAKAAGY-CONH2 CH3 CO-XAAAAQAAAAQAAGY-CONH2 c CH3 CO-XAAAAAAARAAARGGY-NH2 d CH3 CO-AXAAAAKAAAAKAAGY-CONH2 e CH3 CO-AXAAAAAARAAARGGY-NH2 f CH3 CO-AAXAAAAKAAAAKAGY-CONH2 g CH3 CO-AAXAAAAARAAARGGY-NH2 h NH2 −YGGSAKEAAARAAAAXAA-CONH2 i Substitution of residue 32 (C2 position) of α-helix of ubiquitin. j NH2 −YGGSAKEAAARAAAAAXA-CONH2 k NH2 −YGGSAKEAAARAAAAAAX-CONH2 l CH3 CO-YGAAKAAAAKAAAAKAX-COOH m Substitution of residue 35 (C position) of α-helix of ubiquitin. a b

preferences for nonpolar and uncharged polar residues by applying AGADIR to experimental helical peptide data, and found almost identical results [164, 188]. The complete sequences of peptides used can be seen in the table footnote. In general, at N1, N2, and N3, Asp and Glu as well as Ala are preferred, presumably because negative side chains interact favorably with the helix dipole or NH groups while Ala has the strongest interior helix preference.

30 Design and Stability of "-Helices

Although it is also unique in terms of the presence of unsatisﬁed backbone hydrogen bonds, the C-terminal region is less explored experimentally. The Cterminus of the α-helix tends to fray more than the N-terminus, making C-terminal measurements less accurate. Preferences at the C-cap position differ from those at the N-cap. At the N-terminus, the helix geometry favors side chain-to-backbone hydrogen bonding, so polar residues are preferred [14, 19]. At the C-terminus unsatisﬁed backbone hydrogen bonds are fulﬁlled by interactions with backbone groups upstream of the helix. Zhou et al. [48] found that Asn is the most favored residue at the C-cap followed by Gln > Ser∼Ala > Gly∼Thr. Forood et al. [23] tested a limited number of amino acids at the C-terminus (C1) ﬁnding a rank order of Arg > Lys > Ala. Doig and Baldwin [29] determined the C-capping preferences for all 20 amino acids in α-helical peptides. The thermodynamic propensities of some amino acids at C, C-cap, C1, C2, and C3 are also included in Table 6 [189, 190]. 5.3 Phosphorylation

Phosphoserine is destabilizing compared with serine at interior helix positions [191, 192]. We investigated the effect of placing phosphoserine at the N-cap, N1, N2, N3, and interior position in alanine-based α-helical peptides, studying both the −1 and −2 phosphoserine charge states [193]. Phosphoserine stabilizes at the N-terminal positions by as much as 2.3 kcal mol−1 , while it destabilizes in the helix interior by 1.2 kcal mol−1 , relative to serine. The rank order of free energies relative to serine at each position is N2 > N3 > N1 > N-cap > interior. Moreover, −2 phosphoserine is the most preferred residue known at each of these N-terminal positions. Experimental pK a values for the −1 to −2 phosphoserine transition are in the order N2 < N-Cap < N1 < N3 < interior. Phosphoserine can form stabilizing salt bridges to arginine [192]. 5.4 Noncovalent Side-chain Interactions

Many studies have been performed on the stabilizing effects of interactions between amino acid side chains in α-helices. These studies have identiﬁed a number of types of interaction that stabilize the helix including salt bridges [83, 86, 88, 152, 194–198], hydrogen bonds [152, 198–200], hydrophobic interactions [100, 201–203], basic/aromatic interactions [106, 204], and polar/nonpolar interactions [101]. The stabilizing energies of many pairs in these categories have been measured, though some have only been analyzed qualitatively. As described earlier, residue side chains spaced i, i + 3 and i, i + 4 are on the same face of the α-helix, though it is the i, i + 4 spacing that receives most attention in the literature, as these are stronger. A summary of stabilizing energies for side-chain interactions is given in Table 7. We give only those that have been measured in helical peptides with the side-chain interaction energies determined by applying helix/coil theory. Almost all are attractive, with the sole exception of the Lys-Lys repulsion.

5 Forces Affecting "-helix Stability 31 Table 7 Summary of side-chain interaction energies from literature

Interaction Ile-Lys (i, i + 4) Val-Lys (i, i + 4) Ile-Arg (i, i + 4) Phe-Met (i, i + 4) Met-Phe (i, i + 4) Gln-Asn (i, i + 4) Asn-Gln (i, i + 4) Phe-Lys (i, i + 4) Lys-Phe (i, i + 4) Phe-Arg (i, i + 4) Phe-Orn (i, i + 4) Arg-Phe (i, i + 4) Tyr-Lys (i, i + 4) Glu-Phe (i, i + 4) Asp-Lys (i, i + 3) Asp-Lys (i, i + 4) Asp-His (i, i + 3) Asp-His (i, i + 4) Asp-Arg (i, i + 3) Glu-His (i, i + 3) Glu-His (i, i + 4) Glu-Lys (i, i + 3) Glu-Lys (i, i + 4) Phe-His (i, i + 4) Phe-Met (i, i + 4) His-Asp (i, i + 3) His-Asp (i, i + 4) His-Glu (i, i + 3) His-Glu (i, i + 4) Lys-Asp (i, i + 3) Lys-Asp (i, i + 4) Lys-Glu (i, i + 3) Lys-Glu (i, i + 4) Lys-Lys (i, i + 4) Leu-Tyr (i, i + 3) Leu-Tyr (i, i + 4) Met-Phe (i, i + 4) Gln-Asp (i, i + 4) Gln-Glu (i, i + 4) Trp-Arg (i, i + 4) Trp-His (i, i + 4) Tyr-Leu (i, i + 3) Tyr-Leu (i, i + 4) Tyr- Val (i, i + 3) Tyr- Val (i, i + 4) Arg (i, i + 4) Glu (i, i + 4) Arg Arg (i, i + 3) Glu (i, i + 3) Arg Arg (i, i + 3) Glu (i, i + 4) Arg Arg (i, i + 4) Glu (i, i + 3) Arg Phosphoserine-Arg (i, i + 4)

G (kcal mol−1 )

Reference

−0.22 −0.25 −0.22 −0.8 −0.5 −0.5 −0.1 −0.14 −0.10 −0.18 −0.4 −0.1 −0.22 −0.5 −0.12 −0.24 >−0.63 >−0.63 −0.8 −0.23 −0.10 −0.38 −0.44 −1.27 −0.7 −0.53 −2.38 −0.45 −0.54 −0.4 −0.58 −0.38 −0.46 +0.17 −0.44 −0.65 −0.37 −0.97 −0.31 −0.4 −0.8 −0.02 −0.44 −0.13 −0.31 −1.5 −1.0 −0.3 −0.1 −0.45

[101] [101] [101] [100] [100] [200] [200] [106] [106] [106] [204] [106] [106] [252] [227] [227] [253] [253] [254] [227] [227] [152] [152] [198] [203] [198] [241] [227] [227] [227] [227] [227] [227] [196] [153] [153] [203] [199] [152] [252] [161] [153] [153] [153] [153] [255] [255] [255] [255] [192]

32 Design and Stability of "-Helices

5.5 Covalent Side-chain interactions

Lactam (amide) bonds formed between NH3 + and CO2 − side chains can stabilize a helix, acting in a similar way to disulﬁde bridges in a protein by constraining the side chains to be close, reducing the entropy of nonhelical states [205]. Lactam bridges between Lys-Asp, Lys-Glu, and Glu-Orn spaced i, i + 4 have been introduced into analogs of human growth hormone releasing factor [206], and proved to be stabilizing with Lys-Asp most effective. The same Lys-Asp i, i + 4 lactam was stabilizing in other helical peptide systems [207–210], while Lys-Glu i, i + 4 lactam bridges were less effective [211]. Two overlapping Lys-Asp lactams were even more stabilizing [212]. The effect of the ring size formed by the lactam was investigated by replacing Lys with ornithine or (S)-diaminopropionic acid. A ring size of 21 or 22 atoms was most stabilizing (a Lys-Asp i, i + 4 lactam is 20 atoms) [206]. Lactams between side chains spaced i, i + 7 [213, 214] or i, i + 3 [214]; [215], spanning two or one turns of the helix have also been reported. i, i + 7 disulﬁde bonds have been introduced into alanine-based peptides, using (D)- and (L)-2-amino-6-mercaptohexanoic acid derivatives [216]. Strongly stabilizing effects were observed. Some interesting recent work has shown that helix formation can be reversibly photoregulated. Two cysteine residues are cross-linked by an azobenzene derivative which can be photoisomerized from trans to cis, causing a large increase or decrease in the helix content of the peptide, depending on its spacing [217–219].

5.6 Capping Motifs

Although the N-terminal capping box sequence stabilizes helices by inhibiting N-terminal fraying, it does not necessarily promote elongation unless accompanied by favorable hydrophobic interactions as in a “hydrophobic staple” motif [220, 221]. The nature of the capping box stabilizing effect thus not only arises from reciprocal hydrogen bonds between compatible residues, but also from local interactions between side chains, helix macrodipole-charged residue interactions and solvation [222]. Despite statistical analyses revealing that Schellman motifs are observed more frequently that expected at the helix C-terminus, this motif populates only transiently in aqueous solution but it is formed in 30% TFE [223]. This might be due to the C-terminus being very frayed and the increase of helical content contributed from this motif is small. Energetically this motif is not very favorable due to the entropic cost of ﬁxing a Gly residue at the position C . The Schellman motif is believed to be a consequence of helix formation and does not involve α-helix nucleation [224]. The α L motif seems to be more stable than the alternative Schellman motif [221].

5 Forces Affecting "-helix Stability 33

5.7 Ionic Strength

Electrostatic interactions between charged side chains and the helix macrodipole can stabilize the helix [92, 102, 225]. The interactions are potentially quite strong, but are alleviated by the screening effects of water, ions, and nearby protein atoms. In theory, increasing ionic strength of the solvent (up to 1.0 M) should stabilize the helix through interactions with α-helix dipole moments by shifting the equilibrium between α-helix and random coil, which has a random orientation of the peptide dipoles [226]. The energetics of the interaction between fully charged ion pairs can be diminished by added salt and completely screened at 2.5 M NaCl [197, 227]. In peptides containing side chain-to-side-chain interactions, the effect of ion pairs and charge/helix-dipole interactions cannot be clearly separated. There are, however, indications that the interactions of charged residues with the helix macrodipole are less affected than those between charged side chains [227, 228]. In coiled-coil peptides, salt also affects hydrophobic residues by strengthening their interactions at the coiled-coil interface. This can be explained through alterations of the peptide-water interactions at high salt concentration. However, this requires a strong kosmotropic anion to accompany the screening cation [229]. 5.8 Temperature

Thermal unfolding experiments show that the helix unfolds with increasing temperature [230–232]. There is no sign of cold denaturation, as seen with proteins. Enthalpy and entropy changes for the helix/coil transition are difﬁcult to determine as the helix/coil transition is very broad, precluding accurate determination of highand low-temperature baselines by calorimetry [230]. Nevertheless, isothermal titration calorimetric studies of a series of peptides that form helix when binding a nucleating La3+ , ﬁnd H for helix formation to be −1.0 kcal mol−1 [135, 233], in good agreement with the earlier work. 5.9 Triﬂuoroethanol

Peptides with sequences of helices in proteins usually show low helix contents in water. An answer to this problem is to add TFE (2,2,2-triﬂuoroethanol) to induce helix formation [234–237]. For many peptides, the concentration of TFE used to increase the helix content is only up to 40% [234–236, 238, 239]. TFE may act by shielding CO and NH groups from the water solvent while leading to hydrogen bond formation between them. The conformational equilibrium thus shifts toward more compact structures, such as the α-helical conformation [184]. The mechanism involves interaction between TFE and water with several interpretations. One view suggests that TFE indirectly disrupts the solvent shell on α-helices [240, 241]. Another view proposes that TFE destabilizes the unfolded species and thereby

34 Design and Stability of "-Helices

indirectly enhances the kinetics and thermodynamics of folding of the coiled coil [242]. A more compromising view suggests that TFE forms clusters in water solution, which at lower concentration pulls the water molecules from the surface of proteins. At higher concentration, TFE clusters associate with appropriate hydrophobic side chains reducing their conformational entropy and switch the conformation at TFE concentration > 40% [243]. The propagation propensities of all amino acids increase variably in 40% TFE relative to water. The propagation propensities of the nonpolar amino acids increase greatly in 40% TFE whilst other amino acids propensity increase uniformly. However, glycine and proline are strong helix breakers in both in water and 40% TFE solvents [30]. In addition, 40% TFE dramatically alters electrostatic (and polar) interactions and increases the dependence of helix propensities on the sequence [244]. 5.10 pK a Values

Evaluation of pK a values of titrable amino acids in a peptide sequence can be used to analyze the strength of the possible interaction they form in water. pK a shifts of charged residues at the helix termini are signiﬁcant because they can potentially interact with unsatisﬁed hydrogen bonds of the NH groups and CO group at the N-terminus and C-terminus, respectively, or the helix dipole. The pK a values can be measured accurately from the change in ellipticity across a broad range of pH. The asymptotic values of the ellipticities for the different protonation states are ﬁtted to a Henderson-Hasselbach equation to calculate the pK a . In general, the pK a values of Glu and His at N1–N3 are normal compared with those in model compounds. In contrast, Asp and Cys have shifted pK a to lower values [95, 96, 102, 152, 225, 245–248]. An exception for negatively charged residues at the N-cap is that they have a lower pK a [29]. This may be because side chains at the N-cap can form strong hydrogen bonds to NH groups of N2 and N3, while the bonds formed by side chains at N1, N2, and N3 are much weaker [14, 19]. The negatively charged residues at higher pH destabilize helices when at the Ccap [29]. The increased pK a may result from an unfavorable electrostatic interaction with the C-terminal dipole or partial negative charges on the terminal CO groups. 5.11 Relevance to Proteins

Many of the features studied in peptide helices are also applicable to proteins and can be used to rationally modify protein stability or to design new helical proteins. Helices in proteins are often found on the surface with one face exposed to solvent and the other buried in the protein core. Helix propensities and sidechain interactions measured in peptides are thus directly applicable to the solventexposed face. Substitutions at buried positions are much more complex and tertiary interactions also make major contributions to stability. Tertiary interactions at helix

References

termini are rare; nearly all side-chain interactions are local [14]. Preferences for capping sites and the ﬁrst and last turn of the helix are therefore applicable to most protein helices. The feature of protein helices of amphiphilicity, reﬂected in possession of a hydrophobic moment [249], is irrelevant to monomeric isolated helices. Acknowledgments

We thank all our coworkers in this ﬁeld, namely Avi Chakrabartty, Carol Rohl, Buzz Baldwin, Tod Klingler, Ben Stapley, Jim Andrew, Eleri Hughes, Simon Penel, Duncan Cochran, Nicoleta Kokkoni, Jia Ke Sun, Jim Warwicker, Gareth Jones, and Jonathan Hirst. Current work in our lab on helices is supported by the Wellcome Trust (grant references 057318 and 065106). TMI thanks the Indonesian government for a scholarship.

References 1 PAULING, L., COREY, R. B., and BRANSON, H. R. (1951). The structure of proteins: two hydrogen-bonded helical conﬁgurations of the polypeptide chain. Proc. Natl Acad. Sci. USA 37, 205–211. 2 PERUTZ, M. F. (1951). New X-ray evidence on the conﬁguration of polypeptide chains. Nature 167, 1053–1054. 3 KENDREW, J. C., DICKERSON, R. E., STRANDBERG, B. E. et al. (1960). Structure of myoglobin. Nature 185, 422–427. 4 BARLOW, D. J. and THORNTON, J. M. (1988). Helix geometry in proteins. J. Mol. Biol. 201, 601–619. 5 SCHOLTZ, J. M. and BALDWIN, R. L. (1992). The mechanism of α-helix formation by peptides. Annu. Rev. Biophys. Biomol. Struct. 21, 95–118. 6 BALDWIN, R. L. (1995). α-helix formation by peptides of deﬁned sequence. Biophys. Chem. 55, 127–135. 7 CHAKRABARTTY, A. and BALDWIN, R. L. (1995). Stability of α-Helices. Adv. Protein Chem. 46, 141–176. 8 KALLENBACH, N. R., LYU, P., and ZHOU, H. (1996). CD spectroscopy and the helix-coil transition in peptides and polypeptides. In Circular Dichroism and the Conformational Analysis of Biomolecules G. D. FASMAN, editor, Plenum Press, New York, 201–259.

9 ROHL, C. A. and BALDWIN, R. L. (1998). Deciphering rules of helix stability in peptides. Meth. Enzymol. 295, 1–26. 10 ANDREWS, M. J. I. and TABOR, A. B. (1999). Forming stable helical peptides using natural and artiﬁcial amino acids. Tetrahedron 55, 11711–11743. 11 SERRANO, L. (2000). The relationship between sequence and structure in elementary folding units. Adv. Protein Chem. 53, 49–85. 12 RICHARDSON, J. S. and RICHARDSON, D. C. (1988). Amino acid preferences for speciﬁc locations at the ends of α helices. Science 240, 1648–1652. 13 PRESTA, L. G. and ROSE, G. D. (1988). Helix signals in proteins. Science 240, 1632–1641. 14 DOIG, A. J., MACARTHUR, M. W., STAPLEY, B. J., and THORNTON, J. M. (1997). Structures of N-termini of helices in proteins. Protein Sci. 6, 147–155. 15 MACGREGOR, M. J., ISLAM, S. A., and STERNBERG, M. J. E. (1987). Analysis of the relationship between side-chain conformation and secondary structure in globular proteins. J. Mol. Biol. 198, 295–310. 16 SWINDELLS, M. B., MACARTHUR, M. W., and THORNTON, J. M. (1995). Intrinsic ψ propensities of amino acids derived from the coil regions of known

35

36 Design and Stability of "-Helices

structures. Nat. Struct. Biol. 2, 596–603. 17 DUNBRACK, R. and KARPLUS, M. (1994). Conformational analysis of the backbone-dependent rotamer preferences of protein side-chains. Nat. Struct. Biol. 1, 334–339. 18 DUNBRACK, R. L. (2002). Rotamer libraries in the 21(st) century. Curr. Opin. Struct. Biol. 12, 431–440. 19 PENEL, S., HUGHES, E., and DOIG, A. J. (1999). Side-chain structures in the ﬁrst turn of the α-helix. J. Mol. Biol. 287, 127–143. 20 SERRANO, L. and FERSHT, A. R. (1989). Capping and α-helix stability. Nature 342, 296–299. 21 BELL, J. A., BECKTEL, W. J., SAUER, U., BAASE, W. A., and MATTHEWS, B. W. (1992). Dissection of helix capping in T4 lysozyme by structural and thermodynamic analysis of six amino acid substitutions at Thr 59. Biochemistry 31, 3590–3596. 22 CHAKRABARTTY, A., DOIG, A. J., and BALDWIN, R. L. (1993). Helix capping propensities in peptides parallel those in proteins. Proc. Natl Acad. Sci. USA 90, 11332–11336. 23 FOROOD, B., FELICIANO, E. J., and NAMBIAR, K. P. (1993). Stabilization of α-helical structures in short peptides via end capping. Proc. Natl Acad. Sci. USA 90, 838–842. 24 DASGUPTA, S. and BELL, J. A. (1993). Design of helix ends. Amino acid preferences, hydrogen bonding and electrostatic interactions. Int. J. Pept. Protein Res. 41, 499–511. 25 HARPER, E. T. and ROSE, G. D. (1993). Helix stop signals in proteins and peptides: the capping box. Biochemistry 32, 7605–7609. 26 SEALE, J. W., SRINIVASAN, R., and ROSE, G. D. (1994). Sequence determinants of the capping box, a stabilizing motif at the N-termini of α-Helices. Protein Sci. 3, 1741–1745. ˜ OZ, V. and SERRANO, L. (1995). The 27 MUN hydrophobic-staple motif and a role for loop residues in α-helix stability and protein folding. Nat. Struct. Biol. 2, 380–385.

28 VIGUERA, A. R. and SERRANO, L. (1999). Stable proline box motif at the N-terminal end of α-Helices. Protein Sci. 8, 1733–1742. 29 DOIG, A. J. and BALDWIN, R. L. (1995). Nand C-capping preferences for all 20 amino acids in α-helical peptides. Protein Sci. 4, 1325–1336. 30 ROHL, C. A., CHAKRABARTTY, A., and BALDWIN, R. L. (1996). Helix propagation and N-cap propensities of the amino acids measured in alanine-based peptides in 40 volume percent triﬂuoroethanol. Protein Sci. 5, 2623–2637. 31 SCHELLMAN, C. (1980). The αL conformation at the ends of helices. Protein Folding 53–61. 32 AURORA, R., SRINIVASAN, R., and ROSE, G. D. (1994). Rules for α-helix termination by glycine. Science 264, 1126–1130. 33 AURORA, R. and ROSE, G. D. (1998). Helix capping. Protein Sci. 7, 21–38. 34 PRIETO, J. and SERRANO, L. (1997). C-capping and helix stability: the Pro C-capping motif. J. Mol. Biol. 274, 276–288. 35 SIEDLECKA, M., GOCH, G., EJCHART, A., STICHT, H., and BIERZYNSKI, A. (1999). α-Helix nucleation by calcium-binding peptide loop. Proc. Natl Acad. Sci. USA 96, 903–908. 36 JERNIGAN, R., RAGHUNATHAN, G., and BAHAR, I. (1994). Characterisation of interactions and metal-ion binding-sites in proteins. Curr. Opin. Struct. Biol. 4, 256–263. 37 ALBERTS, I. L., NADASSY, K., and WODAK, S. J. (1998). Analyisis of zinc binding sites in protein crystal structures. Protein Sci. 7, 1700–1716. 38 GHADIRI, M. R. and CHOI, C. (1990). Secondary structure nucleation in peptides -transition-metal ion stabilized α-Helices. J. Am. Chem. Soc. 112, 1630–1632. 39 KISE, K. J. and BOWLER, B. E. (2002). Induction of helical structure in a heptapeptide with a metal cross-link: Modiﬁcation of the Lifson-Roig Helix-Coil theory to account for covalent cross-links. Biochemistry 41, 15826–15837.

References 40 RUAN, F., CHEN, Y., and HOPKINS, P. B. (1990). Metal ion enhanced helicity in synthetic peptides containing unnatural, metal-ligating residues. J. Am. Chem. Soc. 112, 9403–9404. 41 CLINE, D. J., THORPE, C., and SCHNEIDER, J. P. (2003). Effects of As(III) binding on α-helical structure. J. Am. Chem. Soc. 125, 2923–2929. 42 KARPEN, M. E., De HASET, P. L., and NEET, K. E. (1992). Differences in the amino acid distributions of 310 -helices and α-Helices. Protein Sci. 1, 1333–1342. 43 BAKER, E. N. and HUBBARD, R. E. (1984). Hydrogen bonding in globular proteins. Prog. Biophys. Mol. Biol. 44, 97–179. 44 N´EMETHY, G., PHILLIPS, D. C., LEACH, S. J., and SCHERAGA, H. A. (1967). A second right-handed helical structure with the parameters of the Pauling-Corey α-helix. Nature 214, 363–365. 45 MILLHAUSER, G. L. (1995). Views of helical peptides-a proposal for the position of 310 -helix along the thermodynamic folding pathway. Biochemistry 34, 3872–3877. 46 BOLIN, K. A. and MILLHAUSER, G. L. (1999). α and 310 : The split personality of polypeptide helices. Acc. Chem. Res. 32, 1027–1033. 47 MIICK, S. M., MARTINEZ, G. V., FIORI, W. R., TODD, A. P., and MILLHAUSER, G. L. (1992). Short alanine-based peptides may form 310 -helices and not α-Helices in aqueous solution. Nature 359, 653–655. 48 ZHOU, H. X. X., LYU, P. C. C., WEMMER, D. E., and KALLENBACH, N. R. (1994). Structure of C-terminal α-helix cap in a synthetic peptide. J. Am. Chem. Soc. 116, 1139–1140. 49 FIORI, W. R. and MILLHAUSER, G. L. (1995). Exploring the peptide 310 -helix/α-helix equilibrium with double label electron spin resonance. Biopolymers 37, 243–250. 50 TONIOLO, C. and BENEDETTI, E. (1991). The polypeptide 310 -helix. Trends Biochem. Sci. 16, 350–353. 51 RAMACHANDRAN, G. N. and SASISEKHARAN, V. (1968). Conformation of polypeptides and proteins. Adv. Protein Chem. 23, 283–437.

52 LOW, B. W. and GRENVILLE-WELLS, H. J. (1953). Generalized mathematical relations for polypeptide chain helixes. The coordinates for the π helix. Proc. Natl Acad. Sci. USA 39, 785–801. 53 SHIRLEY, W. A. and BROOKS, C. L. (1997). Curious structure in “canonical” alanine based peptides. Proteins: Struct. Funct. Genet. 28, 59–71. 54 LEE, K. H., BENSON, D. R., and KUCZERA, K. (2000). Transitions from α to π helix observed in molecular dynamics simulations of synthetic peptides. Biochemistry 39, 13737–13747. 55 WEAVER, T. M. (2000). The π -helix translates structure into function. Protein Sci. 9, 201–206. 56 MORGAN, D. M., LYNN, D. G., MILLER-AUER, H., and MEREDITH, S. C. (2001). A designed Zn2+-binding amphiphilic polypeptide: Energetic consequences of π -helicity. Biochemistry 40, 14020–14029. 57 FODJE, M. N. and AL-KARADAGHI, S. (2002). Occurrence, conformational features and amino acid propensities for the π -helix. Protein Eng. 15, 353–358. 58 BALDWIN, R. L. (2003). In search of the energetic role of peptide hydrogen bonds. J. Biol. Chem. 278, 17581–17588. 59 EPAND, R. M. and SCHERAGA, H. A. (1968). Biochemistry 7, 2864–2872. 60 TANIUCHI, J. H. and ANFINSEN, C. B. (1969). J. Biol. Chem. 244, 2864–2872. 61 SPEK, E. J., GONG, Y., and KALLENBACH, N. R. (1995). Intermolecular interactions in a helical oligo- and poly(L-glutamic acid) at acidic pH. J. Am. Chem. Soc. 117, 10773–10774. 62 BROWN, J. E. and KLEE, W. A. (1971). Biochemistry 10, 470–476. 63 BIERZYNSKI, A., KIM, P. S., and BALDWIN, R. L. (1982). A salt bridge stabilizes the helix formed by isolated C-peptide of ribonuclease A. Proc. Natl Acad. Sci. USA 79, 2470–2474. 64 KIM, P. S., BIERZYNSKI, A., and BALDWIN, R. L. (1982). A competing salt-bridge suppresses helix formation by the isolated C-peptide carboxylate of Ribonuclease A. J. Mol. Biol. 162, 187–199.

37

38 Design and Stability of "-Helices

65 KIM, P. S. and BALDWIN, R. L. (1984). A helix stop signal in the isolated S-peptide of ribonuclease A. Nature 307, 329–334. 66 SHOEMAKER, K. R., KIM, P. S., BREMS, D. N. et al. (1985). Nature of the charged-group effect on the stability of the C-peptide helix. Proc. Natl Acad. Sci. USA 82, 2349–2353. 67 STREHLOW, K. G. and BALDWIN, R. L. (1989). Effect of the substitution Ala-Gly at each of 5 residue positions in the C-peptide helix. Biochemistry 28, 2130–2133. 68 OSTERHOUT, J. J., BALDWIN, R. L., YORK, E. J., STEWART, J. M., DYSON, H. J., and WRIGHT, P. E. (1989). H-1 NMR studies of the solution conformations of an analog of the C-peptide of ribonuclease-A. Biochemistry 29, 7059–7064. 69 SHOEMAKER, K. R., FAIRMAN, R., SCHULTZ, D. A. et al. (1990). Sidechain interactions in the C-peptide helix -Phe8-His12+ . Biopolymers 29, 1–11. 70 FAIRMAN, R., SHOEMAKER, K. R., YORK, E. J., STEWART, J. M., and BALDWIN, R. L. (1990). The Glu2-Arg10+ side-chain interaction in the C-peptidce helix of ribonuclease A. Biophys. Chem. 37, 107–119. 71 STREHLOW, K. G., ROBERTSON, A. D., and BALDWIN, R. L. (1991). Proline for alanine subsitutions in the C-peptide helix of ribonuclease A. Biochemistry 30, 5810–5814. 72 FAIRMAN, R., ARMSTRONG, K. M., SHOEMAKER, K. R., YORK, E. J., STEWART, J. M., and BALDWIN, R. L. (1991). Position effect on apparent helical propensities in the C-peptide helix. J. Mol. Biol. 221, 1395–1401. 73 BLANCO, F. J., JIMENEZ, M. A., RICO, M., SANTORO, J., HERRANZ, J., and NIETO, J. L. (1992). The homologous angiogenin and ribonuclease N-terminal fragments fold into very similar helices when isolated. Biochem. Biophys. Res. Commun. 182, 1491–1498. 74 RICO, M., NIETO, J. L., SANTORO, J., BERMEJO, F. J., and HERRANZ, J. (1983). H1-NMR parameters of the N-terminal 13-residue C-peptide of ribonuclease in

75

76

77

78

79

80

81

82

83

aqueous-solution. Org. Magn. Res. 21, 555–563. GALLEGO, E., HERRANZ, J., NIETO, J. L., RICO, M., and SANTORO, J. (1983). H1-NMR parameters of the N-terminal 19-residue S-peptide of ribonuclease in aqueous-solution. Int. J. Pept. Protein Res. 21, 242–253. RICO, M., NIETO, J. L., SANTORO, J., BERMEJO, F. J., HERRANZ, J., and GALLEGO, E. (1983). Low temperature H1-NMR evidence of the folding of isolated ribonuclease S-peptide. FEBS Lett. 162, 314–319. RICO, M., GALLEGO, E., SANTORO, J., BERMEJO, F. J., NIETO, J. L., and HERRANZ, J. (1984). On the fundamental role of the Glu2∼ . . . Arg10+ salt bridge in the folding of isolated ribonuclease A S-peptide. Biochem. Biophys. Res. Commun. 123, 757–763. NIETO, J. L., RICO, M., JIMENEZ, M. A., HERRANZ, J., and SANTORO, J. (1985). Amide H1-NMR study of the folding of ribonuclease C-peptide. Int. J. Biol. Macromol. 7, 66–70. NIETO, J. L., RICO, M., SANTORO, J., and BERMEJO, J. (1985). NH resonanaces of ribonuclease S-peptide in aqueous solution - low temperature NMR study. Int. J. Pept. Protein Res. 25, 47–55. SANTORO, J., RICO, M., NIETO, J. L., BERMEJO, J., HERRANZ, J., and GALLEGO, E. (1986). C13 NMR spectral assignment of ribonuclease S-peptide - Some new structural information about its low temperature folding. J. Mol. Struct. 141, 243–248. RICO, M., HERRANZ, J., BERMEJO, J. et al. (1986). Quantitative interpretation of the helix coil transition in RNase A S-peptide. J. Mol. Struct. 143, 439–444. RICO, M., SANTORO, J., BERMEJO, J. et al. (1986). Thermodynamic parameters for the helix coil thermal transition of ribonuclease S-peptide and derivatives from H1 NMR data. Biopolymers 25, 1031–1053. MARQUSEE, S. and BALDWIN, R. L. (1987). Helix stabilization by Glu− . . . Lys+ salt bridges in short peptides of de novo design. Proc. Natl Acad. Sci. USA 84, 8898–8902.

References 84 PARK, S. H., SHALONGO, W., and STELLWAGEN, E. (1993). Residue helix parameters obtained from dichroic analysis of peptides of deﬁned sequence. Biochemistry 32, 7048–7053. 85 MARQUSEE, S., ROBBINS, V. H., and BALDWIN, R. L. (1989). Unusually stable helix formation in short alanine based peptides. Proc. Natl Acad. Sci. USA 86, 5286–5290. 86 LYU, P. C., MARKY, L. A., and KALLENBACH, N. R. (1989). The role of ion-pairs in α-helix stability -2 new designed helical peptides. J. Am. Chem. Soc. 111, 2733–2734. 87 LYU, P. C., LIFF, M. I., MARKY, L. A., and KALLENBACH, N. R. (1990). Side-chain contributions to the stability of α-helical structure in peptides. Science 250, 669–673. 88 GANS, P. J., LYU, P. C., MANNING, P. C., WOODY, R. W., and KALLENBACH, N. R. (1991). The helix-coil transition in heterogeneous peptides with speciﬁc side chain interactions: Theory and comparison with circular dichroism. Biopolymers 31, 1605–1614. 89 PENEL, S., MORRISON, R. G., MORTISHIRE-SMITH, R. J., and DOIG, A. J. (1999). Periodicity in α-helix lengths and C-capping preferences. J. Mol. Biol. 293, 1211–1219. 90 WADA, A. (1976). The α-helix as an electric macro-dipole. Adv. Biophys. 9, 1–63. 91 HOL, W. G. J., van DUIJNEN, P. T., and BERENDSEN, H. J. C. (1978). The α-helix dipole and the properties of proteins. Nature 273, 443–446. 92 AQVIST, J., LUECKE, H., QUIOCHO, F. A., and WARSHEL, A. (1991). Dipoles located at helix termini of proteins stabilize charges. Proc. Natl Acad. Sci. USA 88, 2026–2030. 93 ZHUKOVSKY, E. A., MULKERRIN, M. G., and PRESTA, L. G. (1994). Contribution to global protein stabilization of the N-capping box in human growth hormone. Biochemistry 33, 9856–9864. 94 TIDOR, B. (1994). Helix-capping interaction in λ Cro protein: a free energy simulation analysis. Proteins: Struct. Funct. Genet. 19, 310–323.

95 COCHRAN, D. A. E., PENEL, S., and DOIG, A. J. (2001). Contribution of the N1 amino acid residue to the stability of the α-helix. Protein Sci. 10, 463–470. 96 COCHRAN, D. A. E. and DOIG, A. J. (2001). Effects of the N2 residue on the stability of the α-helix for all 20 amino acids. Protein Sci. 10, 1305–1311. 97 IQBALSYAH, T. M. and DOIG, A. J. (in press). Effect of the N3 residue on the stability of the α-helix. Protein Sci. 13, 32–39. 98 DECATUR, S. M. (2000). IR spectro-scopy of isotope-labeled helical peptides: Probing the effect of N-acetylation on helix stabiliTy. Biopolymers 180–185. 99 DOIG, A. J., CHAKRABARTTY, A., KLINGLER, T. M., and BALDWIN, R. L. (1994). Determination of free energies of N-capping in α-Helices by modiﬁcation of the Lifson-Roig helix-coil theory to include N- and C-capping. Biochemistry 33, 3396–3403. 100 STAPLEY, B. J., ROHL, C. A., and DOIG, A. J. (1995). Addition of side chain interactions to modiﬁed Lifson-Roig helix-coil theory: application to energetics of phenylalanine-methionine interactions. Protein Sci. 4, 2383–2391. 101 ANDREW, C. D., PENEL, S., JONES, G. R., and DOIG, A. J. (2001). Stabilising non-polar/polar side chain interactions in the α-helix. Proteins: Struct. Funct. Genet. 45, 449–455. 102 HUYGHUES-DESPOINTES, B. M., SCHOLTZ, J. M., and BALDWIN, R. L. (1993). Effect of a single aspartate on helix stability at different positions in a neutral alanine-based peptide. Protein Sci. 2, 1604–1611. 103 BRANDTS, J. R. and KAPLAN, K. J. (1973). Derivative spectroscopy applied to tyrosyl chromophores. Studies on ribonuclease, lima bean inhibitor, and pancreatic trypsin inhibitor. Biochemistry 10, 470–476. 104 EDELHOCH, H. (1967). Spectroscopic determination of tryptophan and tyrosine in proteins. Biochemistry 6, 1948–1954. 105 CHAKRABARTTY, A., KORTEMME, T., PADMANABHAN, S., and BALDWIN, R. L. (1993). Aromatic side-chain contribution

39

40 Design and Stability of "-Helices

106

107

108

109

110

111

112

to far-ultraviolet circular dichroism of helical peptides and its effect on measurement of helix propensities. Biochemistry 32, 5560–5565. ANDREW, C. D., BHATTACHARJEE, S., KOKKONI, N., HIRST, J. D., JONES, G. R., and DOIG, A. J. (2002). Stabilising interactions between aromatic and basic side chains in α-helical peptides and proteins. Tyrosine effects on helix circular dichroism. J. Am. Chem. Soc. 124, 12706–12714. KEMP, D. S., BOYD, J. G., and MUENDEL, C. C. (1991). The helical s-constant for alanine in water derived from template-nucleated helices. Nature 352, 451–454. KEMP, D. S., ALLEN, T. J., and OSLICK, S. L. (1995). The energetics of helix formation by short templated peptides in aqueous solution. 1. Characterization of the reporting helical template Ac-HE1(1). J. Am. Chem. Soc. 117, 6641–6657. GROEBKE, K., RENOLD, P., TSANG, K. Y., ALLEN, T. J., MCCLURE, K. F., and KEMP, D. S. (1996). Template-nucleated alanine-lysine helices are stabilized by position-dependent interactions between the lysine side chain and the helix barrel. Proc. Natl Acad. Sci. USA 93, 4025–4029. KEMP, D. S., OSLICK, S. L., and ALLEN, T. J. (1996). The structure and energetics of helix formation by short templated peptides in aqueous solution. 3. Calculation of the helical propagation constant s from the template stability constants t/c for Ac-Hel(1)-Ala(n)-OH, n = 1–6. J. Am. Chem. Soc. 118, 4249–4255. KEMP, D. S., ALLEN, T. J., OSLICK, S. L., and BOYD, J. G. (1996). The structure and energetics of helix formation by short templated peptides in aqueous solution. 2. Characterization of the helical structure of Ac-Hel(1)-Ala(6)-OH. J. Am. Chem. Soc. 118, 4240–4248. AUSTIN, R. E., MAPLESTONE, R. A., SEFLER, A. M. et al. (1997). Template for stabilization of a peptide α-helix: Synthesis and evaluation of conformational effects by circular dichroism and NMR. J. Am. Chem. Soc. 119, 6461–6472.

113 ARRHENIUS, T. and SATTHERTHWAIT, A. C. (1990). Peptides: Chemistry, Structure and Biology: Proceedings of the 11th American Peptide Symposium, ESCOM, Leiden. 114 MULLER, K., OBRECHT, D., KNIERZINGER, A. et al. (1993). Perspectives in Medicinal Chemistry, pp. 513–531, Verlag Chemie, Weinheim. 115 GANI, D., LEWIS, A., RUTHERFORD, T. et al. (1998). Design, synthesis, structure and properties of an α-helix cap template derived from N-[(2S)-2-chloropropionyl](2S)-Pro-(2R)-Ala-(2S,4S)-4-thioPro-OMe which initiates α-helical structures. Tetrahedron 54, 15793–15819. 116 ALEMAN, C. (1997). Conformational properties of α-amino acids disubstituted at the α-carbon. J. Phys. Chem. 101, 5046–5050. 117 SACCA, B., FORMAGGIO, F., CRISMA, M., TONIOLO, C., and GENNARO, R. (1997). Linear oligopeptides. 401. In search of a peptide 310 -helix in water. Gazz. Chim. Ital. 127, 495–500. 118 TANAKA, M. (2002). Design and conformation of peptides containing α,α-disubstituted α-amino acids. J. Synth. Org. Chem. Japan 60, 125–136. 119 LANCELOT, N., ELBAYED, K., RAYA, J. et al. (2003). Characterization of the 310 -helix in model peptides by HRMAS NMR spectroscopy. Chem. Eur. J. 9, 1317–1323. 120 KARLE, I. L. and BALARAM, P. (1990). Structural characteristics of α-helical peptide molecules containing Aib residues. Biochemistry 29, 6747–6756. 121 BANERJEE, R. and BASU, G. (2002). A short Aib/Ala-based peptide helix is as stable as an Ala-based peptide helix double its length. ChemBiochem 3, 1263–1266. 122 YOKUM, T. S., GAUTHIER, T. J., HAMMER, R. P., and MCLAUGHLIN, M. L. (1997). Solvent effects on the 310 -/α-helix equilibrium in short amphipathic peptides rich in α,α-disubstituted amino acids. J. Am. Chem. Soc. 119, 1167–1168. 123 MILLHAUSER, G. L., STENLAND, C. J., HANSON, P., BOLIN, K. A., and van De VEN, F. J. M. (1997). Estimating the relative populations of 310 -helix and α-helix in Ala-rich peptides. A hydrogen

References

124

125

126

127

128

129

130

131

132

133

exchange and high ﬁeld NMR study. J. Mol. Biol. 267, 963–974. FIORI, W. R., LUNDBERG, K. M., and MILLHAUSER, G. L. (1994). A single carboxy-terminal arginine determines the amino-terminal helix conformation of an alanine-based peptide. Nat. Struct. Biol. 1, 374–377. YODER, G., POLESE, A., SILVA, R. A. G. D. et al. (1997). Conformation characterization of terminally blocked L-(αMe)Val homopeptides using vibrational and electronic circular dichroism. 310 -Helical stabilization by peptide-peptide interaction. J. Am. Chem. Soc. 119, 10278–10285. KENNEDY, D. F., CRISMA, M., TONIOLO, C., and CHAPMAN, D. (1991). Studies of peptides forming 310 - and α-Helices and β-bend ribbon structures in organic solution and in model biomembranes by Fourier transform infrared spectroscopy. Biochemistry 30, 6541–6548. HUNGERFORD, G., MARTINEZ-INSUA, M., BIRCH, D. J. S., and MOORE, B. D. (1996). A reversible transition between an α-helix and a 310 -helix in a ﬂuorescence-labelled peptide. Angew. Chem. Int. Ed. Engl. 35, 326–329. SHEINERMAN, F. B. and BROOKS, C. L. (1995). 310 -Helices in peptides and proteins as studied by modiﬁed Zimm-Bragg theory. J. Am. Chem. Soc. 117, 10098–10103. ROSE, G. D. and CREAMER, T. P. (1994). Protein-folding -predicting predicting. Proteins: Struct. Funct. Genet. 19, 1–3. DALAL, S., BALASUBRAMANIAN, S., and REGAN, L. (1997). Protein alchemy: Changing β-sheet into α-helix. Nat. Struct. Biol. 4, 548–552. ROSE, G. D. (1997). Protein folding and the paracelsus challenge. Nat. Struct. Biol. 4, 512–514. POLAND, D. and SCHERAGA, H. A. (1970). Theory of Helix-Coil Transitions in Biopolymers. Academic Press, New York and London. QIAN, H. and SCHELLMAN, J. A. (1992). Helix/coil theories: A comparative study for ﬁnite length poly-peptides. J. Phys. Chem. 96, 3987–3994.

134 DOIG, A. J. (2002). Recent advances in helix-coil theory. Biophys. Chem. 101–102, 281–293. 135 ZIMM, B. H. and BRAGG, J. K. (1959). Theory of the phase transition between helix and random coil in polypeptide chains. J. Chem. Phys. 31, 526–535. 136 LIFSON, S. and ROIG, A. (1961). On the theory of the helix-coil transition in polypeptides. J. Chem. Phys. 34, 1963–1974. 137 FLORY, P. J. (1969). Statistical Mechanics of Chain Molecules. Wiley, New York. 138 PAPPU, R. V., SRINIVASAN, R., and ROSE, G. D. (2000). The Flory isolated-pair hypothesis is not valid for polypeptide chains: Implications for protein folding. Proc. Natl Acad. Sci. USA 97, 12565–12570. 139 TIFFANY, M. L. and KRIMM, S. (1968). New chain conformations of poly(glutamic acid) and polylysine. Biopolymers 6, 1379–1382. 140 KRIMM, S. and TIFFANY, M. L. (1974). The circular dichroism spectrum and structure of unordered polypeptides and proteins. Israel J. Chem. 12, 189–200. 141 WOODY, R. W. (1992). Circular dichroism and conformations of unordered polypeptides. Adv. Biophys. Chem. 2, 37–79. 142 BARRON, L. D., HECHT, L., BLANCH, E. W., and BELL, A. F. (2000). Solution structure and dynamics of biomolecules from Raman optical activity. Prog. Biophys. Mol. Biol. 73, 1–49. 143 BLANCH, E. W., MOROZOVA-ROCHE, L. A., COCHRAN, D. A. E., DOIG, A. J., HECHT, L., and BARRON, L. D. (2000). Is polyproline II helix the killer conformation? A Raman optical activity study of the amyloidogenic preﬁbrillar intermediate of human lysozyme. J. Mol. Biol. 301, 553–563. 144 SMYTH, E., SYME, C. D., BLANCH, E. W., HECHT, L., VASAK, M., and BARRON, L. D. (2000). Solution structure of native proteins irregular folds from Raman optical activity. Biopolymers 58, 138–151. 145 SYME, C. D., BLANCH, E. W., HOLT, C. et al. (2002). A Raman optical activity study of the rheomorphism in caseins,

41

42 Design and Stability of "-Helices

146

147

148

149

150

151

152

153

154

155

156

synucleins and tau. Eur. J. Biochem. 269, 148–156. RUCKER, A. L. and CREAMER, T. P. (2002). Polyproline II helical structure in protein unfolded states: Lysine peptides revisited. Protein Sci. 11, 980–1059. SHI, Z. S., WOODY, R. W., and KALLENBACH, N. R. (2002). Is polyproline II a major backbone conformation in unfolded proteins? Adv. Protein Chem. 62, 163–240. PAPPU, R. V. and ROSE, G. D. (2002). A simple model for polyproline II structure in unfolded states of alanine-based peptides. Protein Sci. 11, 2437–2455. SHI, Z., OLSON, C. A., ROSE, G. D., BALDWIN, R. L., and KALLENBACH, N. R. (2002). Polyproline II structure in a sequence of seven alanine residues. Proc. Natl Acad. Sci. USA 99, 9190–9195. RUCKER, A. L., PAGER, C. T., CAMPBELL, M. N., QUALIS, J. E., and CREAMER, T. P. (2003). Host-guest scale of left-handed polyproline II helix formation. Proteins: Struct. Funct. Genet. 53, 68–75. ANDERSEN, N. H. and TONG, H. (1997). Empirical parameterization of a model for predicting peptide helix/coil equilibrium populations. Protein Sci. 6, 1920–1936. SCHOLTZ, J. M., QIAN, H., ROBBINS, V. H., and BALDWIN, R. L. (1993). The energetics of ion-pair and hydrogen-bonding interactions in a helical peptide. Biochemistry 32, 9668–9676. SHALONGO, W. and STELLWAGEN, E. (1995). Incorporation of pairwise interactions into the Lifson-Roig model for helix prediction. Protein Sci. 4, 1161–1166. ARGOS, P. and PALAU, J. (1982). Amino acid distribution in protein secondary structures. Int. J. Pept. Protein Res. 19, 380–393. KUMAR, S. and BANSAL, M. (1998). Dissecting α-Helices: position-speciﬁc analysis of α-Helices in globular proteins. Proteins 31, 460–476. SUN, J. K., PENEL, S., and DOIG, A. J. (2000). Determination of α-helix N1 energies by addition of N1, N2 and N3

157

158

159

160

161

162

163

164

165

166

preferences to helix-coil theory. Protein Sci. 9, 750–754. ROHL, C. A. and DOIG, A. J. (1996). Models for the 310 -helix/coil, π -helix/coil, and α-helix/310 -helix/coil transitions in isolated peptides. Protein Sci. 5, 1687–1696. SUN, J. K. and DOIG, A. J. (1998). Addition of side-chain interactions to 310 -helix/coil and α-helix/310 -helix/coil theory. Protein Sci. 7, 2374–2383. ˜ OZ, V. and SERRANO, L. (1994). MUN Elucidating the folding problem of helical peptides using empirical parameters. Nat. Struct. Biol. 1, 399–409. ˜ OZ, V. and SERRANO, L. (1997). MUN Development of the multiple sequence approximation within the AGADIR model of α-helix formation: comparison with Zimm-Bragg and Lifson-Roig formalisms. Biopolymers 41, 495–509. FERNA´ NDEZ-RECIO, J., V´ASQUEZ, A., CIVERA, C., SEVILLA, P., and SANCHO, J. (1997). The tryptophan/histidine interaction in α-Helices. J. Mol. Biol. 267, 184–197. LACROIX, E., VIGUERA, A. R., and SERRANO, L. (1998). Elucidating the folding problem of α-Helices: local motifs, long-range electrostatics, ionic-strength dependence and prediction of NMR parameters. J. Mol. Biol. 284, 173–191. ˜ OZ, V. and SERRANO, L. (1995). MUN Elucidating the folding problem of helical peptides using empirical parameters. II. Helix macrodipole effects and rational modiﬁcation of the helical content of natural peptides. J. Mol. Biol. 245, 275–296. ˜ OZ, V., YUMOTO, N., PETUKHOV, M., MUN YOSHIKAWA, S., and SERRANO, L. (1998). Position dependence of nonpolar amino acid intrinsic helical propensities. J. Mol. Biol. 278, 279–289. LOMIZE, A. L. and MOSBERG, H. I. (1997). Thermodynamic model of secondary structure for α-helical peptides and proteins. Biopolymers 42, 239–269. V´ASQUEZ, M. and SCHERAGA, H. A. (1988). Effect of sequence-speciﬁc interactions on the stability of helical

References

167

168

169

170

171

172

173

174

175

conformations in polypeptides. Biopolymers 27, 41–58. ROBERTS, C. H. (1990). A hierarchical nesting approach to describe the stability of α helices with side chain interactions. Biopolymers 30, 335–347. GUZZO, A. V. (1965). The inﬂuence of amino acid sequence on protein structure. Biophys. J. 5, 809–822. DAVIES, D. R. (1964). A correlation between amino acid composition and protein structure. J. Mol. Biol. 9, 605–609. CHOU, P. Y. and FASMAN, G. D. (1974). Conformational parameters for amino acids in helical, b-sheet and random coil regions calculated from proteins. Biochemistry 13, 211–221. CHOU, P. Y. and FASMAN, G. D. (1978). Empirical predictions of protein conformation. Annu. Rev. Biochem. 47, 251–276. VON DREELE, P. H., POLAND, D., and SCHERAGA, H. A. (1971). Helix-coil stability constants for the naturally occurring amino acids in water. I. Properties of copolymers and approximate theories. Macromolecules 4, 396–407. VON DREELE, P. H., LOTAN, N., ANANTHANARAYANAN, V. S., ANDREATTA, R. H., POLAND, D., and SCHERAGA, H. A. (1971). Helix-coil stability constants for the naturally occurring amino acids in water. II. Characterization of the host polymers and application of the host-guest technique to random poly-(hydroxypropylglutamine-cohydroxybutylglutamine). Macromolecules 4, 408–417. WOJCIK, J., ALTMANN, K. H., and SCHERAGA, H. A. (1990). Helix-coil stability constants for the naturally occurring amino acids in water. XXIV. Half-cysteine parameters from random poly-(hydroxybutylglutamine-co-Smethylthio-L-cysteine). Biopolymers 30, 121–134. PADMANABHAN, S., YORK, E. J., GERA, L., STEWART, J. M., and BALDWIN, R. L. (1994). Helix-forming tendencies of amino acids in short (hydroxybutyl)-L-glutamine peptides: An

176

177

178

179

180

181

182

183

184

185

evaluation of the contradictory results from host-guest studies and short alanine-based peptides. Biochemistry 33, 8604–8609. CHAKRABARTTY, A., KORTEMME, T., and BALDWIN, R. L. (1994). Helix propensities of the amino acids measured in alanine-based peptides without helix-stabilizing side-chain interactions. Protein Sci. 3, 843–852. BLABER, M., ZHANG, X.-J., and MATTHEWS, B. W. (1993). Structural basis of amino acid α-helix propensity. Science 260, 1637–1640. BLABER, M. W., BAASE, W. A., GASSNER, N., and MATTHEWS, B. W. (1993). Structural basis of amino acid α-helix propensity. Science 260, 1637–1640. BLABER, M., ZHANG, X. J., LINDSTROM, J. D., PEPIOT, S. D., BAASE, W. A., and MATTHEWS, B. W. (1994). Determination of α-helix propensity within the context of a folded protein: sites 44 and 131 in bacteriophage-T4 lysozyme. J. Mol. Biol. 235, 600–624. HOROVITZ, A., MATTHEWS, J. M., and FERSHT, A. R. (1992). A-Helix stability in proteins. II. Factors that inﬂuence stability at an internal position. J. Mol. Biol. 227, 560–568. O’NEIL, K. T. and DEGRADO, W. F. (1990). A thermodynamic scale for the helix-forming tendencies of the commonly occurring amino acids. Science 250, 646–651. PACE, C. N. and SCHOLTZ, J. M. (1998). A helix propensity scale based on experimental studies of peptides and proteins. Biophys. J. 75, 422–427. VILA, J., WILLIAMS, R. L., GRANT, J. A., WOJCIK, J., and SCHERAGA, H. A. (1992). The intrinsic helix-forming tendency of L-alanine. Proc. Natl Acad. Sci. USA 89, 7821–7825. VILA, J. A., RIPOLL, D. R., and SCHERAGA, H. A. (2000). Physical reasons for the unusual α-helix stabilization afforded by charged or neutral polar residues in alanine-rich peptides. Proc. Natl Acad. Sci. USA 97, 13075–13079. ROHL, C. A., FIORI, W., and BALDWIN, R. L. (1999). Alanine is helix-stabilizing in both template-nucleated and standard

43

44 Design and Stability of "-Helices

186

187

188

189

190

191

192

193

194

peptide helices. Proc. Natl Acad. Sci. USA 96, 3682–3687. KENNEDY, R. J., TSANG, K.-Y., and KEMP, D. S. (2002). Consistent helicities from CD and template t/c data for N-templated polyalanines: progress toward resolution of the alanine helicity problem. J. Am. Chem. Soc. 124, 934–944. LYU, P. C., ZHOU, H. X. X., JELVEH, N., WEMMER, D. E., and KALLENBACH, N. R. (1992). Position-dependent stabilizing effects in α-Helices -N-terminal capping in synthetic model peptides. J. Am. Chem. Soc. 114, 6560–6562. PETUKHOV, M., UEGAKI, K., YUMOTO, N., YOSHIKAWA, S., and SERRANO, L. (1999). Position dependence of amino acid intrinsic helical propensities II: Non-charged polar residues: Ser, Thr, Asn, and Gln. Protein Sci. 8, 2144–2150. PETUKHOV, M., UEGAKI, K., YUMOTO, N., and SERRANO, L. (2002). Amino acid intrinsic α-helical propensities III: Positional dependence at several positions of C-terminus. Protein Sci. 11, 766–777. ERMOLENKO, D. N., RICHARDSON, J. M., and MAKHATADZE, G. I. (2003). Noncharged amino acid residues at the solvent-exposed positions in the middle and at the C terminus of the α-helix have the same helical propensity. Protein Sci. 12, 1169–1176. SZALIK, L., MOITRA, J., KRYLOV, D., and VINSON, C. (1997). Phosphorylation destabilizes α-helices. Nat. Struct. Biol. 4, 112–114. LIEHR, S. and CHENAULT, H. K. (1999). A comparison of the α-helix forming propensities and hydrogen bonding properties of serine phosphate and α-amino-g-phosphonobutyric acid. Bioorg. Med. Chem. Lett. 9, 2759–2762. ANDREW, C. D., WARWICKER, J., JONES, G. R., and DOIG, A. J. (2002). Effect of phosphorylation on α-helix stability as a function of position. Biochemistry 41, 1897–1905. HOROVITZ, A., SERRANO, L., AVRON, B., BYCROFT, M., and FERSHT, A. R. (1990). Strength and co-operativity of contributions of surface salt bridges to

195

196

197

198

199

200

201

202

203

204

protein stability. J. Mol. Biol. 216, 1031–1044. MERUTKA, G. and STELLWAGEN, E. (1991). Effect of amino acid ion pairs on peptide helicity. Biochemistry 30, 1591–1594. STELLWAGEN, E., PARK, S.-H., SHALONGO, W., and JAIN, A. (1992). The contribution of residue ion pairs to the helical stability of a model peptide. Biopolymers 32, 1193–1200. HUYGHUES-DESPOINTES, B. M., SCHOLTZ, J. M., and BALDWIN, R. L. (1993). Helical peptides with three pairs of Asp-Arg and Glu-Arg residues in different orientations and spacings. Protein Sci. 2, 80–85. HUYGHUES-DESPOINTES, B. M. and BALDWIN, R. L. (1997). Ion-pair and charged hydrogen-bond interactions between histidine and aspartate in a peptide helix. Biochemistry 36, 1965–1970. HUYGHUES-DESPOINTES, B. M., KLINGLER, T. M., and BALDWIN, R. L. (1995). Measuring the strength of side-chain hydrogen bonds in peptide helices: the Gln.Asp (i,i + 4) interaction. Biochemistry 34, 13267–13271. STAPLEY, B. J. and DOIG, A. J. (1997). Hydrogen bonding interactions between Glutamine and Asparagine in α-helical peptides. J. Mol. Biol. 272, 465–473. PADMANABHAN, S. and BALDWIN, R. L. (1994). Tests for helix-stabilizing interactions between various nonpolar side chains in alanine-based peptides. Protein Sci. 3, 1992–1997. PADMANABHAN, S. and BALDWIN, R. L. (1994). Helix-stabilizing interaction between tyrosine and leucine or valine when the spacing is i,i + 4. J. Mol. Biol. 241, 706–713. VIGUERA, A. R. and SERRANO, L. (1995). Side-chain interactions between sulfur-containing amino acids and phenylalanine in α-helices. Biochemistry 34, 8771–8779. TSOU, L. K., TATKO, C. D., and WATERS, M. L. (2002). Simple cation-p interaction between a phenyl ring and a protonated amine stabilizes an α-helix in water. J. Am. Chem. Soc. 124, 14917–14921.

References 205 TAYLOR, J. W. (2002). The synthesis and study of side-chain lactam-bridged peptides. Biopolymers 66, 49–75. 206 CAMPBELL, R. M., BONGERS, J., and FELXI, A. M. (1995). Rational design, synthesis, and biological evaluation of novel growth-hormone releasing-factor analogs. Biopolymers 37, 67–68. 207 CHOREV, M., ROUBINI, E., MCKEE, R. L. et al. (1991). Cyclic parathyroid-hormone related protein antagonists -lysine 13 to aspartic acid 17 [I to (I + 40] side-chain to side-chain lactamization. Biochemistry 30, 5968–5974. 208 OSAPAY, G. and TAYLOR, J. W. (1992). Multicyclic polypeptide model compounds. 2. Synthesis and conformational properties of a highly α-helical uncosapeptide constrained by 3 side-chain to side-chain lactam bridges. J. Am. Chem. Soc. 114, 6966–6973. 209 BOUVIER, M. and TAYLOR, J. W. (1992). Probing the functional conformation of Neuropeptide-T through the design and study of cyclic analogs. J. Med. Chem. 35, 1145–1155. 210 KAPURNIOTU, A. and TAYLOR, J. W. (1995). Structural and conformational requirements for human calcitonin activity–design, synthesis, and study of lactam-bridged analogs. J. Med. Chem. 38, 836–847. 211 OSAPAY, G. and TAYLOR, J. W. (1990). Multicyclic polypeptide model compounds. 1. Synthesis of a tricyclic amphiphilic α-helical peptide using an oxime resin, segment-condensation approach. J. Am. Chem. Soc. 112, 6046–6051. 212 BRACKEN, C., GULYAS, J., TAYLOR, J. W., and BAUM, J. (1994). Synthesis and nuclear-magnetic-resonance structure determination of an α-helical, bicyclic, lactam-bridged hexapeptide. J. Am. Chem. Soc. 116, 6431–6432. 213 CHEN, S. T., CHEN, H. J., YU, H. M., and WANG, K. T. (1993). Facile synthesis of a short peptide with a side-chain-constrained structure. J. Chem. Res. (S) 6, 228–229. 214 ZHANG, W. T., and TAYLOR, J. W. (1996). Efﬁcient solid-phase synthesis of peptides with tripodal side-chain bridges

215

216

217

218

219

220

221

222

223

224

and optimization of the solvent conditions for solid-phase cyclizations. Tetrahedron Lett. 37, 2173–2176. LUO, P. Z., BRADDOCK, D. T., SUBRAMANIAN, R. M., MEREDITH, S. C., and LYNN, D. G. (1994). Structural and thermodynamic characterization of a bioactive peptide model of apolipoprotein-E-side-chain lactam bridges to constrain the conformation. Biochemistry 33, 12367–12377. JACKSON, D. Y., KING, D. S., CHMIELEWSKI, J., SINGH, S., and SCHULTZ, P. G. (1991). General approach to the synthesis of short α-helical peptides. J. Am. Chem. Soc. 113, 9391–9392. KUMITA, J. R., SMART, O. S., and WOOLLEY, G. A. (2000). Photo-control of helix content in a short peptide. Proc. Natl Acad. Sci. USA 97, 3803–3808. FLINT, D. G., KUMITA, J. R., SMART, O. S., and WOLLEY, G. A. (2002). Using an azobenzene cross-linker to either increase or decrease peptide helix content upon trans-to-cis photoisomerization. Chem. Biol. 9, 391–397. KUMITA, J. R., FLINT, D. G., SMART, O. S., and WOOLLEY, G. A. (2002). Photo-control of peptide helix content by an azobenzene cross-linker: steric interactions with underlying residues are not critical. Protein Eng. 15, 561–569. ˜ OZ, V., RICO, M., JIME´ NEZ, M. A., MUN and SERRANO, L. (1994). Helix stop and start signals in peptides and proteins. The capping box does not necessarily prevent helix elongation. J. Mol. Biol. 242, 487–496. KALLENBACH, N. R. and GONG, Y. X. (1999). C-terminal capping motifs in model helical peptides. Bioorg. Med. Chem. 7, 143–151. PETUKHOV, M., YUMOTO, N., MURASE, S., ONMURA, R., and YOSHIKAWA, S. (1996). Factors that affect the stabilization of α-helices in short peptides by a capping box. Biochemistry 35, 387–397. VIGUERA, A. R. and SERRANO, L. (1995). Experimental analysis of the Schellman motif. J. Mol. Biol. 251, 150–160. GONG, Y., ZHOU, H. X., GUO, M., and KALLENBACH, N. R. (1995). Structural

45

46 Design and Stability of "-Helices

225

226

227

228

229

230

231

232

233

analysis of the N- and C-termini in a peptide with consensus sequence. Protein Sci. 4, 1446–1456. ARMSTRONG, K. M. and BALDWIN, R. L. (1993). Charged histidine affects α-helix stability at all positions in the helix by interacting with the backbone charges. Proc. Natl Acad. Sci. USA 90, 11337–11340. SCHOLTZ, J. M., YORK, E. J., STEWART, J. M., and BALDWIN, R. L. (1991). A neutral water-soluble, α-helical peptide: The effect of ionic strength on the helix-coil equilibrium. J. Am. Chem. Soc. 113, 5102–5104. SMITH, J. S. and SCHOLTZ, J. M. (1998). Energetics of polar side-chain interactions in helical peptides: Salt effects on ion pairs and hydrogen bonds. Biochemistry 37, 33–40. LOCKHART, D. J. and KIM, P. S. (1993). Electrostatic screening of charge and dipole interactions with the helix backbone. Science 260, 198–202. JELESAROV, I., DURR, E., THOMAS, R. M., and BOSSHARD, H. B. (1998). Salt effects on hydrophobic interaction and charge screening in the folding of a negatively charged peptide to a coiled coil (leucine zipper). Biochemistry 37, 7539–7550. SCHOLTZ, J. M., MARQUSEE, S., BALDWIN, R. L. et al. (1991). Calorimetric determination of the enthalpy change for the α-helix to coil transition of an alanine peptide in water. Proc. Natl Acad. Sci. USA 88, 2854–2858. YODER, G., PANCOSKA, P., and KEIDERLING, T. A. (1997). Characterization of alanine-rich peptides, Ac-(AAKAA)n-GY-NH2 (n) 1–4), using vibrational circular dichroism and fourier transform infrared conformational determination and thermal unfolding. Biochemistry 36, 5123–5133. HUANG, C. Y., KLEMKE, J. W., GETAHUN, Z., DEGRADO, W. F., and GAI, F. (2001). Temperature-dependent helix-coil transition of an alanine based peptide. J. Am. Chem. Soc. 123, 9235–9238. GOCH, G., MACIEJCZYK, M., OLESZCZUK, M., STACHOWIAK, D., MALICKA, J., and BIERZYNSKI, A. (2003). Experimental

234

235

236

237

238

239

240

241

242

243

investigation of initial steps of helix propagation in model peptides. Biochemistry 42, 6840–6847. NELSON, J. W. and NELSON, N. R. K. (1986). Stabilization of the ribonuclease S-peptide α-helix by triﬂuoroethanol. Proteins: Struct. Funct. Genet. 1, 211–217. NELSON, J. W. and NELSON, N. R. K. (1989). Persistence of the α-helix stop signal in the S-peptide in triﬂuoroethanol solutions. Biochemistry 28, 5256–5261. SONNICHSEN, F. D., van EYK, J. E., HODGES, R. S., and SYKES, B. D. (1992). Effect of triﬂuoroethanol on protein secondary structure: an NMR and CD study using a synthetic actin peptide. Biochemistry 31, 8790–8798. WATERHOUS, D. V. and JOHNSON, W. C. (1994). Importance of environment in determining secondary structure in proteins. Biochemistry 33, 2121–2128. JASANOFF, A. and FERSHT, A. R. (1984). Quantitative determination of helical propensities. Analysis of data from triﬂuoroethanol titration curves. Biochemistry 33, 2129–2135. ALBERT, J. S. and HAMILTON, A. D. (1995). Stabilization of helical domains in short peptides using hydrophobic interactions. Biochemistry 34, 984–290. WALGERS, R., LEE, T. C., and CAMMERS-GOODWIN, A. (1998). An indirect chaotropic mechanism for the stabilization of helix conformation of peptides in aqueous triﬂuoroethanol and hexaﬂuoro-2-propanol. J. Am. Chem. Soc. 120, 5073–5079. LUO, P. and BALDWIN, R. L. (1999). Interaction between water and polar groups of the helix backbone: An important determinant of helix propensities. Proc. Natl Acad. Sci. USA 96, 4930–4935. KENTSIS, A. and SOSNICK, T. R. (1998). Triﬂuoroethanol promotes helix formation by destabilizing backbone exposure: Desolvation rather than native hydrogen bonding deﬁnes the kinetic pathway of dimeric coiled coil folding. Biochemistry 37, 14613–14622. REIERSEN, H. and REES, A. R. (2000). Triﬂuoroethanol may form a solvent

References

244

245

246

247

248

249

250

matrix for assisted hydrophobic interactions between peptide sidechains. Protein Eng. 13, 739–743. MYERS, J. K., PACE, N., and SCHOLTZ, J. M. (1998). Triﬂuoroethanol effects on helix propensity and electrostatic interactions in the helical peptide from ribonuclease T1. Protein Sci. 7, 383–388. NOZAKI, Y. and TANFORD, C. (1967). Intrinsic dissociation constants of aspartyl and glutamyl carboxyl groups. J. Biol. Chem. 242, 4731–4735. KYTE, J. (1995). Structure in Protein Chemistry, Garland Publishing, New York and London. KORTEMME, T. and CREIGHTON, T. E. (1995). Ionisation of cysteine residues at the termini of model α-helical peptides. Relevance to unusual thiol pKa values in proteins of the thio-redoxin family. J. Mol. Biol. 253, 799–812. MIRANDA, J. J. (2003). Position-dependent interactions between cysteine and the helix dipole. Protein Sci. 12, 73–81. EISENBERG, D., WEISS, R. M., and TERWILLIGER, T. C. (1982). The helical hydrophobic moment: a measure of the amphiphilicity of a helix. Nature 299, 371–374. ˜ OZ, V. and SERRANO, L. (1995). MUN Analysis of i; i + 5 and i; i + 8 hydrophobic interactions in a helical

251

252

253

254

255

model peptide bearing the hydro-phobic staple motif. Biochemistry 34, 15301–15306. THOMAS, S. T., LOLADZE, V. V., and MAKHATADZE, G. I. (2001). Hydration of the peptide backbone largely deﬁnes the thermodynamic propensity scale of residues at the C position of the C-capping box of α-Helices. Proc. Natl Acad. Sci. USA 98, 10670–10675. SHI, Z., OLSON, C. A., BELL, A. J., and KALLENBACH, N. R. (2002). Non-classical helix-stabilizing interactions: C–H. . .O H-bonding between Phe and Glu side chains in α-helical peptides. Biophys. Chem. 101–2, 267–279. LUO, R., DAVID, L., HUNG, H., DEVANEY, J., and GILSON, M. K. (1999). Strength of solvent-exposed salt-bridges. J. Phys. Chem. B 103, 366–380. MARQUSEE, S. and SAUER, R. T. (1994). Contributions of a hydrogen bond/salt bridge network to the stability of secondary and tertiary structure in lambda repressor. Protein Sci. 3, 2217–2225. SHI, Z., OLSON, C. A., BELL, A. J., and KALLENBACH, N. R. (2001). Stabilization of α-helix structure by polar side-chain interactions: Complex salt bridges, cation π interactions and C–H . . . O–H bonds. Biopolymers 60, 366–380.

47

1

Design and Stability of Peptide β-sheets Mark S. Searle University of Nottingham, Nottingham, United Kingdom

Originally published in: Protein Folding Handbook. Part I. Edited by Johannes Buchner and Thomas Kiefhaber. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30784-2

1 Introduction

The pathway by which the polypeptide chain assembles from the unfolded, “disordered” state to the final active folded protein has been the subject of intense investigation. Hierarchical models of protein folding emphasize the importance of local interactions in restricting the conformational space of the polypeptide chain in the search for the native state. These nuclei of structure promote interactions between different parts of the sequence leading ultimately to a cooperative ratelimiting step from which the native state emerges [1–4]. Designed peptides that fold autonomously in water (α-helices and β-sheets) have proved extremely valuable in probing the relationship between local sequence information and folded conformation (the stereochemical code) in the absence of the tertiary interactions found in the native state of proteins. This has allowed intrinsic secondary structure propensities to be investigated in isolation, and enabled the nature and strength of the weak interactions relevant to a wide range of molecular recognition phenomena in chemistry and biology to be put on a quantitative footing. While the literature is rich in studies of α-helical peptides [5–8], water-soluble, nonaggregating monomeric β-sheets have emerged relatively recently [9–11]. For reasons of design and chemical synthesis, these are almost exclusively antiparallel β-sheets, although others have used nonnatural linkers to engineer parallel strand alignments [12–14]. Here we focus on contiguous antiparallel β-sheet systems. Autonomously folding β-hairpin motifs, consisting of two antiparallel β-strands linked by a reverse β-turn (Figure 1), represent the simplest systems for probing weak interactions in β-sheet folding and assembly, although more recently a number of three- and four-stranded β-sheet structures have been described. From this growing body of data, key factors have come into focus that are important in rational Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Design and Stability of Peptide $-sheets

Fig. 1 a) Beta-strand alignment and interstrand hydrogen bonds in an antiparallel $-hairpin peptide; R groups represent amino acid side chains, main chain N and angles are shown. N-terminal $-hairpin of ubiquitin in which the native TLTGK turn se-

quence has been replaced by the sequence NPDG. b) Native strand alignment giving rise to a type I NPDG turn. c) Nonnative strand alignment with a G-bulged type I turn (NPDGT).

design. The following will be considered: the role of the β-turn in promoting and stabilizing antiparallel β-sheet formation; the role of intrinsic backbone φ, ψ propensities in preorganizing the extended conformation of the polypeptide chain; the role of cooperativity in β-sheet folding and stability and in the propagation of multistranded β-sheets, and quantitative approaches to estimating the energetics of β-sheet stability and folding. 2 β-Hairpins Derived from Native Protein Sequences

The early focus on peptides excised from native protein structures provided the first insights into autonomously folding β-hairpins, preceding the more rational

3 Role of $-turns in Nucleating $-hairpin Folding

approach to β-sheet design. Peptides derived from tendamistat [15], B1 domain of protein G [16, 17], ubiquitin [18, 19], and ferredoxin [20] showed that these sequences could exist in the monomeric form without aggregating, but that in most cases they showed a very limited tendency to fold in the absence of tertiary contacts. The use of organic co-solvents appeared to induce native-like conformation [17, 18, 20]. The study by Blanco et al. [20] of a peptide derived from the B1 domain of protein G provided the first example of native-like folding in water of a fully native peptide sequence. In contrast, the studies of hairpins isolated from the proteins ubiquitin and ferredoxin, which are structurally homologous to the G B1 domain (all form an α/β-roll fold) showed these to be unfolded in water [18, 20]. More recent studies of the native ubiquitin peptide have now revealed evidence for a small population of the folded state in water [21], while a β-turn mutation has been shown to significantly enhance folding [22]. The apparent lack of evidence for folding of the peptide derived from residues 15–23 of tendamistat (YQSWRYSQA) [15], and from the N-terminal sequence of ubiquitin (MQIFVKTLTGKTITLEV) [19] led to partial redesign of the sequence to enhance folding through modification to the β-turn sequence by introducing an NPDG type I turn, which is the most common type I turn sequence in proteins. Thus, in the former peptide SWRY was replaced by NPDG [15], and in the latter the G-bulged type I turn TLTGK was replaced by NPDG [19]. By introducing this tight two-residue loop across PD it was envisaged that both hairpins would be stabilized. This was certainly the case, however, the most striking observation was that both peptides folded into nonnative conformations with a three-residue G-bulged type I turn (PDG) reestablished across the turn (Figure 1b and 1c). These initial studies in rational redesign led to strikingly irrational results, prompting a much more systematic approach to β-hairpin design. The above results revealed that the βturn sequence, which dictates the preferred backbone geometry, appears to be an important factor in dictating the β-strand alignment. From the protein folding viewpoint, as demonstrated by the redesigned ubiquitin hairpin sequence, it is evident that one important role of the native turn sequence may be to preclude the formation of nonnative conformations that may be incompatible with formation of the native state.

3 Role of β-turns in Nucleating β-hairpin Folding

The systematic classification of β-turns in proteins reveals a wide variety of geometries and sizes of loop [2, 23, 24]. For the design of β-hairpins, the emphasis has been on incorporation of the smallest turn sequence possible to limit the entropic destabilization effects. While two-residue type I and type II turns are generally common in β-turns, the backbone conformation (φ and torsion angles) of a two-residue type I or type II turn results in a local left-handed twist, which is not compatible with the right-handed twist found in protein β-sheets. Consequently, these turns are less commonly found between contiguous antiparallel β-strands. However, the diastereomeric type I and II turns have the φ and and angles complementary

3

4 Design and Stability of Peptide $-sheets

to the right-handed twisted orientation of the β-strands. These conclusions appear to rationalize, at least in part, the above observations with β-turn modifications introduced into the β-hairpins of tendamistat and ubiquitin. The introduced NPDG type I turn is not compatible with the right-handed twist of the β-strands resulting in a refolding to a more flexible G-bulged type I turn. The work of Gellman et al. [25] has shown the importance of backbone φ, angle preferences for the residues in the turn sequence by comparing the stability of a number of β-hairpin peptides derived from the ubiquitin sequence (MQIFVKSXXKTITLVKV) containing either XX = L-Pro-Xaa or D-Pro-Xaa. Replacing L-Pro with D-Pro switches the twist from left-handed (type I or II) to right-handed (type I or II ), making the latter compatible with the right-handed twist of the two β-strands. NMR data (Hα shifts and long-range nuclear Overhauser effects (NOEs)) indicate that each of the D-Pro containing peptides showed a significant degree of folding, whereas the L-Pro analogs appeared to be unfolded. Similar conclusions were drawn from studies of a series of 12-mers containing XG turn sequences, with X = L-Pro or D-Pro [26]. A number of natural L-amino acids are commonly found in the α L region of conformational space and are compatible with the type I or II turn conformation. Statistical analyses from a number of groups have identified Xaa-Gly as a favored type I turn. A number of studies have used the Asn-Gly sequence to design βhairpin motifs and have demonstrated a high population of the folded structure with the required turn conformation and strand alignment [27–32]. The work by de Alba et al. [28] identified two possible conformations of the peptide ITSYNGKTYGR. The NOE data appear to be compatible with rapidly inter-converting conformations involving an YNGK type I turn and a NGKT type II turn, each giving a distinct pattern of cross-strand NOEs. Ramirez-Alvarado et al. [27] also described an NGcontaining 12-mer (RGITVXGKTYGR; X = N), which they subsequently extended to a series of hairpins to examine the correlation between hairpin stability and the database frequency of occurrence of residues in position X [29]. Using nuclear magnetic resonance (NMR) and circular dichroism (CD) measurements they concluded that X = Asn > Asp > Gly > Ala > Ser in promoting hairpin folding, in agreement with the intrinsic φ, preferences of these residues. Despite extensive analysis of NG type I turns in the protein database (PDB), it is still not entirely clear why Asn at the first position is so effective in promoting turn formation. There is no evidence for specific side chain to main chain hydrogen bonds that might stabilise the desired backbone conformation, although specific solvation effects cannot be ruled out [30]. Evidence that the NG turn is able to nucleate folding in the absence of cross-strand interactions was demonstrated in a truncated analog of one designed 16-residue β-hairpin sequence KKYTVSINGKKITVSI (β1 in Figure 2), in which the sequence was shortened to SINGKKITVSI, lacking the N-terminal six residues [30]. Evidence from NOE data showed that the turn was significantly populated with interactions observed between Ser6 Hα ↔ Lys11 Hα and Ile7 NH ↔ Lys10 NH (Figure 2b) that are only compatible with a folded type I turn around NG. Two destabilized β-hairpin mutants (KKYTVSINGKKITKSK with electrostatic repulsion between the N- and

3 Role of $-turns in Nucleating $-hairpin Folding

Fig. 2 Structure of the 16-residue $-hairpin sequence $1, and mutants $2;,$3 and $4 in which cross-strand salt bridges (Lys-Glu) have been introduced at position X2 -X15 and X6 -X11 . In b), hairpin $1 has been truncated by removing the N-terminal hydrophobic

residues (1–5). The resulting peptide still shows the ability to fold around the turn sequence as evident from medium range NOEs (indicated by arrows) that show that the INGK sequence is adopting a type I NG turn.

C-terminal Lys residues, and KKATASINGKKITVSI with the loss of key hydrophobic residues in one strand) showed no evidence from NMR chemical shift data for cross-strand interactions; however, careful examination of NOE data revealed evidence for NG turn nucleation [30]. Titration with organic co-solvent showed both peptides to fold significantly, indicating that the turn sequence probably already predisposes the peptide to form a β-hairpin but that favorable cross-strand interactions are required for stability. The work of de Alba et al. convincingly illustrated this principle in a series of six hairpin sequences (10-mers) where strand residues were conserved but turn sequences varied [28]. Using a number of NMR criteria they were able to show that changes in turn sequence could result in a variety of turn conformations including two residue 2:2 turns, 3:5 turns and 4:4 turns (see earlier nomenclature [22–24]), with different pairings of amino acid side chains. As with the earlier examples cited with the turn modification described for hairpins derived from tendamistat and ubiquitin, the bulged-type I turn (3:5 turn) appears to be an intrinsically stable turn with the necessary right-handed twist. Together these data strongly support a model

5

6 Design and Stability of Peptide $-sheets

for hairpin folding in which the turn sequence strongly dictates its preferred conformation, and that strand alignment, cross-strand interactions and subsequently conformational stability are dictated by the specificity of the turn.

4 Intrinsic φ, Propensities of Amino Acids

Statistical analyses of high-resolution structures in the PDB have provided significant insights into residue-specific intrinsic backbone φ; ψ preferences in polypeptide chains. A novel approach presented by Swindells et al., was to determine φ, ψ propensities of different residues in nonregular regions of protein structure where backbone geometry is free of interactions associated with regular hydrogen bonded β-sheet or α-helical secondary structure [33]. The striking observation is that in this context φ and ψ angles (see Figure 3) are far from randomly distributed, and that most occupy regions of Ramachandran space associated with regular secondary structure. The observed φ; ψ distributions for individual residues has been taken as representative of those found in denatured states of proteins providing the basis of a “random coil” state from which residue-specific NMR parameters (3 JNH-H α and NOE intensities) can be derived as a reference state for folding studies [34, 35].

Fig. 3 Ramachandran plots of residue backbone N and angles taken from a database of 512 high-resolution protein Xray structures showing: a) residues in regular $-sheet (N, −120◦ , 120◦ ) and "-helix (N;, −60◦ , 60◦ ), and b) residues in the irregular coil regions of the same structures

(N, 60◦ , 0◦ is the "L region of conformationa space mainly occupied by Gly). The distribution in b) shows that residues have a natura propensity to occupy the " and $ regions of conformational space even when they are not involved in regular protein secondary structure. Taken from Ref [36].

4 Intrinsic N, Propensities of Amino Acids 7

While β-propensity is found to vary significantly from one residue to the next, context-dependent effects also appear to play an important part [36]. While V, I, F, and Y, for example, have a high intrinsic preference to be in the βregion of the Ramachandran plot where steric interactions with flanking residues are minimized, their conformation is relatively insensitive to the nature of the flanking residues. In contrast, small or unbranched side chains have a higher preference for the α-helical conformation, however, this preference can be significantly modulated by its neighbors through a combination of steric and hydrophobic interactions, as well as both repulsive and attractive electrostatic interactions. General effects of flanking residues (grouped as α-like or β-like, reflecting φ propensities) on the central residue of a XXX triplet are illustrated in Figure 4 as an average over all residues at the central position. More specific effects are also shown for

Fig. 4 Effects of neighboring residues on residue $-propensities within the triplets XNX calculated from the data in Figure 3. The $-propensity of residue N is a measure of the number of times a particular residues is found in the $-region of the Ramachandran plot as a fraction of the total distribution between "- and $-space [$/(" + $)]. The context dependence of the $-propensity is estimated by considering the nature of the neighboring residue (X) (X = any residue, " is a residue that prefers the "-helical region – Asp, Glu, Lys, or Ser, $ is

a residue that prefers the $-sheet region – Ile, Val, Phe, or Tyr). The effects of neighboring residues on the averages-propensity is shown in (a), specific effects on Ser (b), Val (c) and Lys (d) are also shown. While Val is relatively insensitive to the nature of the flanking residues, the smaller Ser residue can be forced to adopt a higher $propensity if it has bulky neighbors. Thus, the intrinsic $-propensity of a particular residue is highly context dependent. Taken in part from Ref. [36].

8 Design and Stability of Peptide $-sheets

Ser, Val, and Lys (Figure 4). With Ser, for example, having bulky flanking residues either side with high β-propensity (denoted βSβ, where β could be V, I, F or Y), significantly increases the β-propensity of the Ser residue to minimize the steric repulsion between the two bulky neighboring residues [36]. Thus, intrinsic structural propensities appear to be highly context dependent. This statistical framework has been extended to a number of experimental systems to examine the extent to which isolated β-strand sequences (in the absence of secondary structure interactions) are predisposed by the primary sequence to adopt an extended β-like conformation. The isolated 8-mer (GKKITVSI), corresponding to the C-terminal β-strand of the hairpin β1 (Figure 2; KKYTVSINGK-KITVSI), was examined by NMR analysis of 3 JNH-Hα values and backbone NOE intensities. Surprisingly, many of these parameters are similar to those for the folded hairpin despite the monomeric nature of the 8-mer [32, 36]. In an analogous study of the C-terminal strand of the ubiquitin hairpin described above [21, 37], similarly large deviations of coupling constants and NOE intensities from random coil values suggested that the isolated β-strands are partially preorganized into an extended conformation supporting a model for hairpin folding which may not require a significant further organisation of the peptide backbone, a factor that may contribute significantly to hairpin stability. Several studies of denaturated states of proteins have also highlighted the influence of neighboring residues in modulating main chain conformational preferences [38–41], and the importance of residual structure in the unfolded state in guiding the conformational search to the native state [42, 43].

5 Side-chain Interactions and β-hairpin Stability

There is a strong case, at least in the context of isolated peptide fragments, that the origin of the specificity of β-hairpin folding is largely dictated by the conformational preferences of the turn sequence. However, the stability of the folded state has been attributed to interstrand hydrogen bonding and/or hydrophobic interactions, though which dominates is still a matter of debate. Ramirez-Alvarado et al. [27] reported that the population of the folded state of the hairpin RGITVNGKTYGR was significantly diminished by replacing residues on the N-terminal strand, and then the C-terminal strand, by Ala. The loss of stability was attributed firstly to a reduction in hydrophobic surface burial, but also due to the intrinsically lower β-propensity of Ala, the latter contributing through an adverse conformational entropy term. To compensate for this de Alba et al. [44] described a family of hairpins derived from the sequence IYSNSDGTWT. The effects of residue substitutions in the first three positions was examined while maintaining the overall β-character of the two strands. Several favorable cross-strand pair-wise interactions were identified that were apparent in earlier, and more recent, PDB analysis of β-sheet interactions [45, 46]. For example, Thr-Thr and Tyr-Thr cross-strand pairs produced stabilizing interactions, whilst Ile-Thr and Ile-Trp had a destabilizing effect. Other studies of

5 Side-chain Interactions and $-hairpin Stability

Ala substitution in one β-strand have similarly highlighted hydrophobic burial as a key factor in conformational stability [30], while the observation of large numbers of side chain NOEs have been used as evidence for hydrophobic stabilization in water (see below) [19, 25–27, 30]. 5.1 Aromatic Clusters Stabilize β-hairpins

The first example of a natural β-hairpin sequence that folded autonomously in water (residues 41–56 of the B1 domain of protein G) identified an aromatic-rich cluster of residues that appears to impart considerable stability [17]. The interstrand pairing of Trp/Val and Tyr/Phe has subsequently been exploited in the design of a number of model hairpin systems, in particular to examine the relationship between the position of this stabilizing cluster and the β-turn sequence. The separation between the loop sequence and cluster strongly influences stability and the extent of participation in β-sheet forming interactions (Figure 5a). In an isomeric family of 20-mers (peptides 1, 2, and 3), all of which contain exactly the same residue composition, the most stable hairpin is that in which the smallest cost in conformational entropy is paid to bring the cluster together, i.e., where the cluster is closest to the loop sequence [47]. This arrangement results in the largest Hα chemical shift deviations, indicating a well-formed core; however, the terminal residues show a lower propensity to fold. Thus, there exists a strong interplay between the two key stabilizing components of the hairpin. A statistical model was developed to rationalize the experimental observations and estimate the free energy change versus the number of peptide hydrogen bonds formed (Figure 5b). It is a well-known phenomenon that α-helical peptides become more stable as the length increases, reflecting the fact that while helix nucleation is energetically unfavorable, the propagation step has a small net increase in stability. In β-hairpin systems, Stanger et al. [48] suggest that this may not be the case. There is some evidence for an increase in stability as strands lengthen from five to seven residues, however, further extension (to nine) does not lead to a further stability increment, suggesting that there may be an intrinsic limit to strand length. Since the choice of sequence extension in this study was limited to an all Thr extension or an alternating Ser-Thr (ST)n extension the conclusions should be viewed with caution. Since crossstrand Ser/Ser and Thr/Thr pairings do not bury very much hydrophobic surface area, it is not surprising perhaps that cross-strand side-chain interactions may only just compensate for the entropic cost of organizing the peptide backbone. Studies with other more favorable pairings may be enlightening. Undoubtedly the most successfully designed structural motif to date has been the tryptophan zipper (trpzip) motif [49], whose stability exceeds substantially all those already described. The design is based around stabilizing nonhydrogen-bonded cross-strand Trp-Trp pairs (Table 1). The most successful designs involved two such Trp-Trp pairs which NMR structural analysis reveals are interdigitated in a zipper-like manner stabilized through face-face offset π-stacking giving a compact structure (Figure 6). This arrangement

9

10 Design and Stability of Peptide $-sheets

Fig. 5 Interplay between hydrophobic cluster and turn position in a family of isomeric $-hairpins. The distance of the WVYF cluster from the turn is shown in (a) for peptides 1, 2, and 3. A statistical mechanical model was used to estimate the free energy of folding for the three peptides according to the number of peptide hydrogen bonds formed and the relative position of the stabilizing hydrophobic cluster (b). Thus, peptide 1, which has the hydrophobic cluster

closest to the turn, has the smallest energetic cost of forming the cluster which then significantly stabilizes this core motif, but leads to fraying of the N- and C-termini. In contrast, in the case of peptide 3, there is a large energy penalty in ordering the peptide backbone to bring the residues of the cluster together. Overall the core hairpin is less stable but more residues are involved in ordered structure. Adapted from Ref. [47].

5 Side-chain Interactions and $-hairpin Stability Table 1 trpzip β-hairpin peptides, sequences and stability

β-hairpin

Sequence

Turn type

Tm

H (kJ mol−1 )

trpzip1 trpzip2 trpzip3

SWTWEGNKWTWK SWTWENGKWTWK SWTWED PNKWTWK

(Type II turn) (Type I turn) (Type II turn)

323 345 352

−45.4 −70.6 −54.8

Fig. 6 NMR structures of trpzips 1 and 2. A) Ensemble of 20 structures of trpzip1 (residues 2–11) showing the relative orientations of the indole rings. B) Overlay of trpzip 1 and 2 aligned to the peptide backbone of residues 2–5 and 8–11 (top),

and rotated by 90◦ for the end-on view of the indole rings. The backbone carbonyl of residue 6 is labeled to illustrate the different turn geometries of the two hairpins (type II and type I ). Taken from Ref [49].

11

12 Design and Stability of Peptide $-sheets

of the indole rings results in a pronounced signature in the CD spectrum with intense exciton-coupling bands at 215 and 229 nm indicative of interactions between aromatic chromophores in a highly chiral environment (see Figure 7). The high sensitivity of the CD bands permits thermal denaturation curves to be determined with high sensitivity, in contrast to other β-sheet systems where only small changes in the CD spectrum at 216 nm are evident, resulting in poor signal-to-noise. The thermal unfolding curves are sigmoidal and reversible, fitting to a two-state folding model (Figure 7). The trpzip hairpins 1, 2, and 3 show high T m values (see Table 1) with folding strongly enthalpy driven. Changing the turn sequence (GN versus NG versus DPN) has a significant impact on stability. The unfolding curve for trpzip2 with the NG type I turn gives the most cooperative unfolding transition and appears to be the most stable of the three at room temperature, despite other studies that suggest that the unnatural D-PN (type II ) turn is the most stabilizing [25]. Clearly, context-dependent factors are at work. Also, the strongly enthalpy-driven signature for folding is different to that reported previously for other systems [32], reflecting differences in the nature of the stabilizing interactions which in the case of the trpzip peptides involve π-π stacking interactions. Despite the fact that the trpzip hairpins represent a highly stabilized motif, no such examples have been found in the protein structure database. The authors suggest that on steric grounds it may be difficult to accommodate the trpzip motif within a multistranded β-sheet or through packing against other structural elements. 5.2 Salt Bridges Enhance Hairpin Stability

The high abundance of salt bridges on the surface of hyperthermophilic proteins [50, 51], together with examples of rational enhancement of protein stability through redesign of surface charge, has strongly implicated ionic interactions as a stabilizing force [52, 53]. In model peptides that are only weakly folded in water, the relative importance of ionic versus hydrophobic interactions in stabilizing local secondary structure has been less well investigated in terms of a detailed quantitative description. To this end, peptide β1 (Figure 2) was mutated to introduce Lys-Glu salt bridges at two positions within the hairpin involving substitution of Ser → Glu: one salt bridge is postioned adjacent to the β-turn, while the other involves residues close to the N- and C-termini (see Figure 2) [54]. Using NMR chemical shift data from Ha resonances we have estimated the net contribution to stability from these two interactions. Although the contribution to stability in each case is small, Hα chemical shifts are extremely sensitive to small shifts in the population of the folded state, as evident from the δHα data in Figure 8. On this basis, the individual contributions of these two interactions was estimated (K2-E15, peptide β2, and E6-K11, peptide β3) compared with their K2-S15 and S6-K11 counterparts (peptide β1) and found to be similar (−1.2 and −1.3 kJ mol−1 at 298K). When the two salt bridges are introduced simultaneously into the hairpin sequence (β4), the energetic contribution of the two interactions together (−3.6 kJ mol−1 ) is

5 Side-chain Interactions and $-hairpin Stability

Fig. 7 Folding data for trpzips 1 to 3 determined from near- and far-UV CD spectra. a) CD spectrum of trpzip 1, with inset of nearUV CD showing buried aromatic residues (10-fold expansion). b) Reversible thermal

denaturation of trpzip 1 monitored by CD at 229 nm (unfolding and refolding curves overlayed). c) Temperature dependence of folding for trpzips 1 to 3 plotted as fraction folded. Taken from Ref. [49].

13

14 Design and Stability of Peptide $-sheets

Fig. 8 Effects of salt bridges (Lys-Glu) on $-hairpin stability. H" chemical shift deviation from random coil values (*H" values) for $-hairpin peptides $2 to $4 compared with those of the reference hairpin

protein $1 containing Lys-Ser cross-strand pairs at the X2 -X15 and X6 -X11 mutation sites (see Figure 2). a) $1 versus $2; b) $1 versus $3, and c) $1 versus $4. All data at 298K and pH 5.5. Taken from Ref. [54].

5 Side-chain Interactions and $-hairpin Stability

significantly greater than the sum of the individual interactions. This effect is readily apparent from the large increase in -δ-Hα values in Figure 8, indicating that the contribution of a given interaction appears to depend on the relative stability of the system in which the interaction is being measured. Similar observations have been reported using a disulfide cyclized β-hairpin scaffold [55]. The indication is

Fig. 9 a) Thermodynamic cycle showing the context-dependent energetic contribution to hairpin stability of each electrostatic interaction in the $1 to $4 family of hairpins (Figure 2) by comparing relative hairpin stabilities at 298K determined from NMR chemical shift data; G values are shown for Lys-Glu interactions at pH 5.5. The stabilizing contribution of each salt bridge is context dependent, showing some degree

of cooperativity between the two mutation sites according to the degree of preorganization of the hairpin structure. b) Schematic illustration of the abundance of cross-strand NOEs in hairpin peptide $4. c) Family of five NMR structures of hairpin $4 showing the peptide backbone alignment, with some fraying at the N- and C-termini. Taken from Ref. [54].

15

16 Design and Stability of Peptide $-sheets

that the strength of the interaction appears to depend on the degree of preorganization of the β-hairpin template that pays varying degrees of the entropic cost in bringing the pairs of side chains together. This is more clearly illustrated by the schematic representation shown in Figure 9 that shows the thermodynamic cycle indicating the effects on stability of the introduction of each mutation. Thus, the energetic contribution of the S15/E15 mutation could be measured either from β1 ←→ β2 or from β3 ←→ β4. The values obtained are quite different, the latter suggesting that the interaction is more favorable (−1.2 versus −2.3 kJ mol−1 ). This appears to correlate with the fact that the stability of β3 is greater than β1 and that the degree of preorganization of the former determines the energetic contribution of the interaction. The same principle is evident when determining the energetics of the E6-K11 interaction. This can be estimated from β1 ←→ β3 or β2 ←→ β4. Again, the energetics are quite different (−1.3 versus −2.4 kJ mol−1 ) with the larger contribution from the β2 ←→ β4 pair, where the intrinsic stability of the β2 reference state is higher. The NMR data show that the folded conformation of β4 is highly populated (> 70%) giving rise to an abundance of cross-strand NOEs (Figure 9) [55]. On the basis of 173 restraints (NOEs and torsion angle restraints from 3 JNH-Hα values) we have calculated a family of structures compatible with the NMR data (Figure 9). The large number of van der Waals contacts between hydrophobic residues (V, I, and Y)

Fig. 10 CD melting curves for $4 at various concentrations (% v/v) of MeOH, as indicated. The nonlinear least squares fit is shown in each case from which thermodynamic data for folding were determined

(see Ref. [54]). The change in heat capacity on folding in 30% and 50% aqueous methanol is assumed to be zero; there is evidence for cold-denaturation in water and 10% methanol. Taken from Ref. [54].

7 Quantitative Analysis of Peptide Folding 17

evident from the NOE data leads us to conclude that hydrophobic interactions still provide the overall driving force for folding with structure and stability further consolidated by additional Coulombic interactions from the salt bridges. CD melting curves for hairpin β4 in water and various concentrations of methanol (Figure 10) show the same characteristics described above from temperature-dependent NMR data for β1: a shallow melting curve for β4 in water with evidence for cold denaturation is consistent with the hydrophobic interaction again providing the dominant driving force for folding (H = +11:9 kJ mol−1 and S = +38 J K−1 mol−1 ), while folding becomes strongly enthalpy driven in 50% aqueous methanol (H = −39:8 kJ mol−1 and S = −106 J K−1 mol−1 ).

6 Cooperative Interactions in β-sheet Peptides: Kinetic Barriers to Folding

Folding kinetics for a β-hairpin derived from the C-terminus of the B1 domain of protein G have been described from measurements of tryptophan fluorescence following laser-induced temperature-jump [56, 57]. Kinetic analysis of this peptide (and a dansylated analogue) reveals a single exponential relaxation process with time constant 3.7 ± 0:3 µs. The data indicate a single kinetic barrier separating folded and unfolded states, consistent with a two-state model for folding. Subsequently, the authors developed a statistical mechanical model based on these observations, describing the stability in terms of a minimal numbers of parameters: loss of conformational entropy, backbone stabilization by hydrogen bonding and formation of a stabilizing hydrophobic cluster between three key residues. This model seems sufficient to reproduce all of the features observed experimentally, with a rough, funnel-like energy landscape dominated by two global minima representing the folded and unfolded states. The formation of the hydrophobic cluster appears to be a key folding event. Nucleation by the turn seems most likely, consistent with experimental measurements of loop formation on the timescale of ≈1 µs [58]. However, simulation studies by others suggest that folding may proceed by hydrophobic collapse followed by rearrangement to form the hydrophobic cluster, with hydrogen bonds then propagating outward from the cluster in both directions [59]. Such a model does not appear to require a turn-based nucleation event.

7 Quantitative Analysis of Peptide Folding

Quantitative analysis of the population of folded β-sheet structures in solution still presents a challenge, largely as a consequence of uncertainties in limiting spectroscopic parameters for the fully folded state. Far UV-CD has been considered to be unreliable as a consequence of the complicating influence of the β-turn conformation and possibly aromatic residues, where present [10]. Added to this, the CD spectrum of β-sheet is intrinsically weak compared with α-helical secondary structure.

18 Design and Stability of Peptide $-sheets

The trpzip peptides [49] represent the exception to the rule, as discussed above. The use of NMR parameters (Hα chemical shifts, 3 JNH-Hα values and NOE intensities) to quantify folded populations has been discussed [9–11, 32, 60]. NMR offers the advantage that several independent parameters can be used in quantitative analysis to provide a consensus picture of the folded state. There still appear to be significant discrepancies between CD analysis and NMR, with peptides that appear to be significantly folded by NMR giving rise to a largely random coil CD spectrum. One interpretation of this observation is that in aqueous solution the peptides fold as a collapsed state with an ill-defined hydrogen bonding network dominated by sidechain interactions. Interestingly, in many cases the addition of organic co-solvents changes the CD spectrum dramatically. The interpretation of co-solvent-induced folding is also subject to some uncertainty. Does trifluorethanol or methanol actually significantly perturb the equilibrium between the folded and unfolded states (induce folding), or do these solvents exert their influence by changing the nature of the folded state such as to stabilize interstrand hydrogen bonding interactions without significantly changing the folded population? The latter hypothesis would appear to more readily account for the observation of solvent-induced effects on the CD spectrum, and finds some support from studies of cyclic β-hairpin analogs where the folded population is fixed, but whose CD spectrum undergoes large solvent-induced changes [60]. The use of cyclic β-hairpin analogs has been exploited in a number of studies to generate a fully folded NMR reference state for comparison with the folding of acyclic analogs [61, 62]. Backbone cyclization through amide bond formation or through a disulfide bridge seem to work equally well. Such an approach has been used effectively to measure the thermodynamics of folding of a short hairpin carrying a motif of aromatic residues [62]. The cyclic analogs show a much higher stability than their acyclic counterparts, including significant protection from amide H/D exchange due to enforced interstrand hydrogen bonding. While peptide cyclization seems a worthwhile approach to defining the fully folded state, there may also be some caveats to this approach that have not been tested. When conformational constraints (β-turns) are imposed at both ends of the structure this may affect the intrinsic twist of the two β-strands, resulting in a more pronounced twist in the cyclic analog than in the acyclic hairpin. Further, the latitude for conformational dynamics in the fully folded state will also be different. Both of these factors are likely to influence Hα shifts chemical shifts to some degree.

8 Thermodynamics of β-hairpin Folding

The number of β-hairpin model systems is expanding rapidly with a greater focus now on quantitative analysis and thermodynamic characterization. The thermodynamic signature for the folding of β1 (Figure 2) presents an insight into the nature of the stabilizing weak interactions in various solvent milieu [30, 32]. β1 exhibits

8 Thermodynamics of $-hairpin Folding

Fig. 11 a) Temperature-dependent stability of the hairpin peptide $1 (see Figure 2) at pH 5.5 in water, 20% methanol and 50% methanol. Temperature is plotted against the RMS value for the deviation of H" chemical shifts from random coil values, assuming a two-state folding model. In water the peptide shows “hot” and “cold” denaturation, but in 50% methanol the folded population increases at low temperature. The best fits to the three sets of data are shown; in water folding is slightly entropydriven with a large negative C◦ p value, while in 50% methanol folding is strongly

enthalpy-driven with C◦ p close to zero. In 20% methanol the values are in between. (Reproduced with permission from the Journal of the American Chemical Society [32]). b) Variable-temperature FTIR spectra of i) 2 mM aqueous solution of the 8-mer peptide GKKITVSI (C-terminal $-strand of hairpin $1); ii) 2 mM solution of $-hairpin peptide $1; iii) 10 mM solution of $1, all in the temperature range 278–330 K and in D2 O solution, pH 5.0 (uncorrected). In iii) the band that appears around 1620 cm−1 is formed irreversibly indicative of peptide aggregation. Reproduced from Ref [64].

the property of cold denaturation, with a maximum in the stability curve occurring at 298K as judged by changes in Hα chemical shift (Figure 11). Such pronounced curvature is clear evidence for entropy-driven folding accompanied by a significant change in heat capacity. Both of these thermodynamic signatures are hallmarks of the hydrophobic effect contributing strongly to hairpin stability. More recent studies by Tatko and Waters [63] have also shown that cold denaturation effects can be observed for simple model peptides, consistent with substantial changes in heat capacity on folding. Further examination of folding in the presence of methanol co-solvent shows that the signature changes such that folding becomes strongly enthalpy driven, and that in 50% aqueous methanol the temperaturestability profile is indicative of the absence of any significant contribution of the hydrophobic effect to folding [32]. The population of the folded state appears to be enhanced by methanol, reflecting similar observations in helical peptides where the phenomenon has been attributed to the effects of the co-solvent destabilizing

19

20 Design and Stability of Peptide $-sheets

the unfolded peptide chain so promoting hydrogen bonding interactions in the folded state. It is unlikely that the folded state of a model β-hairpin peptide resembles a β-sheet in a native protein, with the former sampling a much larger number of conformations of similar energy stabilized by a fluctuating ensemble of transient interactions. IR analysis of the amide I band of β1 does not show significant differences in the region expected for β-sheet formation (≈ 1630 cm−1 ) from data on a nonhydrogen-bonded short reference peptide [64]. However, this band does appear under aggregating conditions (Figure 11). In other cases, IR spectral features characteristic of β-hairpins have been shown to form reversibly at relatively high peptide concentrations [65]. Similarly, β-hairpins that appear to be well folded on the basis of various NMR criteria seem to be weakly folded by CD analysis [60]. This discrepancy has been attributed largely to weak interstrand hydrogen bonding interactions in the folded state. Indeed, molecular dynamics simulations using ensemble-averaging approaches or time-averaged NOEs tend to de-emphasize the role of hydrogen bonding between the peptide backbone of the two strands, but emphasize the role of hydrophobic side chains interactions [30, 60, 67]. In all cases described, side chain NOEs across the β-strands support such interactions, however, cross-strand backbone NOEs only imply the possibility of hydrogen bonds since weakly populated folded states lead to only small NH ←→ ND protection factors in water. In studies (both calorimetric and NMR) of the folding of the C-terminal hairpin from the B1 domain of protein G, enthalpy-driven folding is observed in water [68]. In contrast with the above data, where the stabilizing hydrophobic interactions involved aliphatic side chains, here an aromatic cluster is responsible for folding. A strongly enthalpy-driven transition is also apparent for the trpzip motifs described above [49] all of which is consistent with π-π interactions stabilizing β-hairpin structures through fundamentally electrostatic interactions rather than through only solvophobic effects. Studies of a designed β-hairpin system, also carrying the same motif of three aromatic residues, demonstrate qualitatively similiar enthalpydriven folding [62], while the thermodynamics of the N-terminal hairpin component of a designed three-stranded antiparallel β-sheet enables similar conclusions to be drawn (see further below) [69]. The difference in the thermodynamic signature for aliphatic versus aromatic side-chain interactions has been suggested to have its origins in enthalpy-entropy compensation effects, such that enthalpy-driven interactions may be fundamentally a consequence of tighter interfacial interactions, giving rise to stronger electrostatic (enthalpic) interactions [62]. More recent work by Tatko and Waters [70] probing the energetics of aromatic-aliphatic interactions in hairpins suggests that there is a preference for the self-association of aromatics over aromatic-aliphatic interactions, and that the unique nature of this interaction may impart some degree of sequence selectivity. The limited data available from such systems suggest that the thermodynamic driving force for β-hairpin folding is highly dependent on the nature of the side-chain interactions involved. Further quantitative analysis of peptide folding is required to substantiate these hypotheses.

9 Multistranded Antiparallel $-sheet Peptides

9 Multistranded Antiparallel β-sheet Peptides

The natural extension of the earlier studies on β-hairpin peptides was to design three- and four-stranded antiparallel β-sheets using the design principles already discussed, focusing on the importance of turn sequence in defining stability and strand alignment, and employing motifs of interacting side chains already identified to impart stability. An overriding question concerns the extent to which cooperative interactions perpendicular to the strand direction are important in stabilizing these structures (see Figure 12); in other words, how good is a pre-organized βhairpin motif at templating the interaction of a third strand. Several studies have attempted to address this important question. The earliest study described a 24residue peptide incorporating two NG turns (KKFTLSINGKKYTISNGKTYITGR) that showed little evidence for folding in water but was significantly stabilised in aqueous methanol solutions [71, 72]. By comparison with a 16-residue β-hairpin analog consisting of the same C-terminal sequence (GKKYTISNGKTYITGR), it was possible to show that the Hα shift perturbations for the C-terminal β-hairpin were greater in the presence of the interactions of the third strand. Subsequent design strategies, incorporating the D-Pro-Gly loop, together with other Asn-Glycontaining turn sequences have illustrated that it is possible to design structures that fold in water [69, 73–75], some of which show a degree of cooperative stabilization between different strands. Other studies have reported peptides that fold to three- (and four-) stranded β-sheets in organic solvents [76], but a quantitative dissection of stabilizing interactions between β-strands has not been presented. Redesign of the three-stranded Betanova, following earlier work [74], reported the use of an automated approach using the algorithm PERLA (protein engineering rotamer library algorithm) to define stabilizing and destabilizing single and multipleresidue mutations producing incremental increases in stability over the original design of up to ≈4 kJ mol−1 [77]. Schenck and Gellman [73] demonstrated cooperative interactions using their D-ProGly to L-ProGly switch, the latter destabilizing one hairpin component selectively. From chemical shift analysis they showed that the individual β-hairpins are cooperatively stabilized by the presence of the third strand. De Alba et al. [75] also reported a designed β-sheet system, but were unable to find convincing evidence for cooperative stabilization of either hairpin by the third strand. It is clear that folding is not a highly cooperative process in these peptide systems. Thus, the folded three-stranded β-sheet is more than likely in equilibrium with populations of the individual hairpins and the unfolded state (see Figure 12). One quantitative study of cooperative interactions between the strands of a threestranded β-sheet [69] was based on a designed system incorporating a previously studied β-hairpin with the third strand capable of forming a stabilizing motif of aromatic residues (Figure 13), similar to that already described [17, 62]. The earlier study of the isolated C-terminal β-hairpin showed cold denaturation [30], approximating to two-state unfolding. In the designed three-stranded system, the same hairpin component shows the same cold denaturation profile, however, the N-

21

22 Design and Stability of Peptide $-sheets

Fig. 12 a) Models for the folding of $hairpin and three-stranded $-sheet peptides illustrating the possibility of cooperative interactions being propagated parallel i) and perpendicular ii) to the $-strand direction. b) Four-state model for the folding of a three-stranded antiparallel $-sheet peptide showing the presence at equilibrium of the

unfolded state and partially folded states in which the C-terminal $-hairpin is formed but the N-terminus is disordered, the Nterminal $-hairpin is formed but the Cterminus is disordered, and the fully folded state. The preformed hairpins can act as templates for the folding of the third strand.

terminal hairpin (sharing a common central strand) showed increased folding at low temperature. While the first process is characterized by entropy-driven folding, the latter is enthalpy-driven (Figure 13). Clearly, two different thermodynamic profiles are not consistent with a single two-state folding model, but the data could be rationalized in terms of a four-state model in which the individual hairpins with an unfolded tail are also populated (Figure 12). Examination of the folding of the isolated C-terminal hairpin, and comparison of the data with that of the three-

9 Multistranded Antiparallel $-sheet Peptides

Fig. 13 a) Mean NMR structure of the fully folded state of the three-stranded antiparallel $-sheet structure of sequence KGEWTFVNG9 KYTVSING17 KKITVSI showing the core cluster of hydrophobic residues (underlined). b) Temperature-dependent stability profiles for the various $-hairpin components of the three-stranded sheet using the H" splitting of Gly9 and Gly17 in the two $-turns: Gly9 (filled circles), Gly17 (open circles). Gly17 (open squares)

in the isolated C-terminal hairpin peptide (G9 KYTVSING17 KKITVSI) is also shown. A larger Gly H" splitting indicates a higher population of the folded hairpin. Different profiles for Gly9 and Gly17 in the threestranded sheet show that the peptide cannot be folding via a simple two-state model involving only random coil and fully folded peptide. The data have been fitted assuming the four-state model shown in Figure 12. Reproduced from Ref. [69].

23

24 Design and Stability of Peptide $-sheets

stranded analog, shows good evidence that the C-terminal hairpin is cooperatively stabilized by the interaction of the third strand, even though overall folding is not highly cooperative. The data in Figure 13 show the temperature dependence of the splitting of the Gly Hα resonances in the two NG turns. Larger values indicate a higher folded population. The splitting for G17 is greater in the three-stranded sheet than for the isolated C-terminal hairpin, while many Hα resonances are further downfield shifted than in the isolated hairpin. The temperature dependence of the stability shows the C-terminal hairpin in both cases to undergo cold denaturation. The N-terminal hairpin carrying the aromatic motif of residues increases in population at low temperature; fitting the data shows the former to be entropy driven and the latter enthalpy driven [69]. Entropic factors seem to be the likely explanation for the small cooperative stabilization effect on the C-terminal hairpin (< 2 kJ mol−1 ), with each hairpin providing a possible template against which the third strand can interact. With one strand preorganized, the entropic

Fig. 14 Structure of the four-stranded $-sheet peptide D PD PD P-cc (a), and the cyclic reference peptide c(D P)2 -II (b) used for estimating limiting values for the fully folded state of the C-terminal hairpin of D PD PD P-cc. Adapted from the work of Ref. [78].

10 Conclusions

cost of association of an additional strand is largely confined to the associating strand [73, 78]. The nature of the folded state is unlikely to compare with that of a β-sheet in a native protein, more likely, hydrophobic contacts between side chains stabilize a collapsed conformation where interstrand hydrogen bonds may play a minor stabilizing role. These “loosely” defined interactions between side chains, rather than a native-like “crystalline” array of hydrogen bonds, may explain why cooperative interactions have only a small effect on overall stability because only a small energy barrier separates the folded from partially or fully unfolded states. Several of the above studies have attempted to address the issue of whether cooperative interactions are propagated orthogonal to the strand direction as the number of β-strands increases. Gellman and coworkers have examined the influence of strand number on antiparallel β-sheet stability in designed three- and four-stranded structures (see Figure 14), using the D-Pro-Gly β-turn sequence to define β-turn position and strand length. The results are not dissimilar to those described above; a third strand stabilizes an existing two-stranded β-sheet by ≈2 kJ mol−1 , while a fourth strand seems to confer a similar small stability increment [79].

10 Conclusions

In contrast to the highly cooperative folding behavior characteristic of native globular proteins, simple model β-sheet systems, which lack defining tertiary interactions, do not possess this characteristic. The limited number of three- and fourstranded β-sheet peptides so far described confirms this, although some evidence for cooperative interactions between β-strands have been presented and rationalized on the basis of the entropic benefits associated with adding an additional β-strand to a preformed template. In all cases to date, it seems that designed threeand four-stranded β-sheet structures are in equilibrium with their partially folded β-hairpin components. This is an indication that the energy landscape is relatively flat unlike that for most native proteins where there is a significant energy difference between the single native structure and other conformations. It is interesting to look at examples of the smallest known β-sheet proteins and make comparisons with the designed systems described above. The WW domains [80–82] form a single folded motif consisting of three strands of antiparallel β-sheet but with well-defined tertiary interactions arising from folding back of the N- and C-termini to form a small compact hydrophobic core. This defining feature appears to allow the WW domains to fold co-operatively despite their small size (some as small as 35 residues) (Figure 15). Apart from the desire to be able to design molecules to order with specific tailored properties, the model βsheet systems described have enabled considerable progress to be made in testing our understanding of the basic design principles of β-sheets and fundamental aspects of weak interactions.

25

26 Design and Stability of Peptide $-sheets

Fig. 15 NMR structure of the WW domain of the formin binding protein (PDB code: 1EOI), consisting of a three-stranded antiparallel $sheet motif; the side chains of conserved residues are shown with tertiary contacts evident between W8 and P33.

References 1 HONIG, B. J. Mol. Biol. 1999; 293, 283–293. 2 BROCKWELL, D. J., SMITH, D. A., RADFORD, S. E. Curr. Opin. Struct. Biol. 2000; 10, 16–25. 3 BALDWIN, R. L., ROSE, G. Trends Biochem. Sci. 1999; 24, 26–76. 4 BALDWIN, R. L., ROSE, G. Trends Biochem. Sci. 1999; 24, 77–83. 5 CHAKRABARTTY, A., BALDWIN, R. Adv. Protein Chem. 1995; 46, 141–176. 6 MUNOZ, V., SERRANO, L. J. Mol. Biol. 1995; 245(3), 275–296. 7 O’NEIL, K. T., DEGRADO, W. F. Science 1990; 250(4981), 646–651. 8 HILL, R., DEGRADO, W. J. Am. Chem. Soc. 1998; 120(6), 1138–1145. 9 GELLMAN, S. H. Curr. Opin. Chem. Biol. 1998; 2, 717–725. 10 RAMIREZ-ALVARADO, M., KORTEMME, T., BLANCO, F. J., SERRANO, L. Bioorg. Med. Chem. 1999; 7, 93–103. 11 SEARLE, M. S. J. Chem. Soc. Perkin Trans. 2001; 2, 1011–1020. 12 KEMP, D. S., BOWEN, B. R., MEUNDEL, C. C. J. Org. Chem. 1990; 55, 4650–4657.

13 NOWICK, J. S., SMITH, E. M., PAIRISH, M. Chem. Soc. Rev. 1996; 401–415. 14 DIAZ, H., TSANG, K. Y., CHOO, D., ESPINA, J. R., KELLY, J. W. J. Am. Chem. Soc. 1993; 115, 3790–3791. 15 BLANCO, F. J., JIMENEZ, M. A., HERRANZ, J., RICO, M., SANTORO, J., NIETO, J. L. J. Am. Chem. Soc. 1993; 115, 5887–5889. 16 BLANCO, F. J., RIVAS, G., SERRANO, L. Nat. Struct. Biol. 1994; 1, 584–590. 17 BLANCO, F. J., JIMENEZ, M. A., PINEDA, A., RICO, M., SANTORO, J., NIETO, J. L. Biochemistry 1994; 33, 6004–6014. 18 COX, J. P. L., EVANS, P. A., PACKMAN, L. C., WILLIAMS, D. H., WOOLFSON, D. N. J. Mol. Biol. 1993; 234, 483–492. 19 SEARLE, M. S., WILLIAMS, D. H., PACKMAN, L. C. Nat. Struct. Biol. 1995; 2, 999–1006. 20 SEARLE, M. S., ZERELLA, R., WILLIAMS, D. H., PACKMAN, L. C. Protein Eng. 1996; 9, 559–565. 21 ZERELLA, R., EVANS, P. A., IONIDES, J. M. C. et al. Protein Sci. 1999; 8, 1320– 1331.

References 22 ZERELLA, R., CHEN, P. Y., EVANS, P. A., RAINE, A., WILLIAMS, D. H. Protein Sci. 2000; 9, 2142–2150. 23 SIBANDA, B. L., BLUNDELL, T. L., THORNTON, J. M. J. Mol. Biol. 1989; 206, 759–777. 24 HUTCHINSON, E. G., THORNTON, J. M. Protein Sci. 1994; 3, 2207–2216. 25 HAQUE, T. S., GELLMAN, S. H. J. Am. Chem. Soc. 1997; 119, 2303–2304. 26 STANGER, H. E., GELLMAN, S. H. J. Am. Chem. Soc. 1998; 120, 4236–4237. 27 RAMIREZ-ALVARADO, M., BLANCO, F. J., SERRANO, L. Nat. Struct. Biol. 1996; 3, 604–612. 28 De ALBA, E., JIMENEZ, M. A., RICO, M. J. Am. Chem. Soc. 1997; 119, 175–183. 29 RAMIREZ-ALVARADO, M., BLANCO, F. J., NIEMANN, H., SERRANO, L. J. Mol. Biol. 1997; 273, 898–912. 30 GRIFFITHS-JONES, S. R., MAYNARD, A. J., SEARLE, M. S. J. Mol. Biol. 1999; 292, 1051–1069. 31 MAYNARD, A. J., SEARLE, M. S. J. Chem. Soc. Chem. Commun. 1997; 1297–1298. 32 MAYNARD, A. J., SHARMAN, G. J., SEARLE, M. S. J. Am. Chem. Soc. 1998; 120, 1996–2007. 33 SWINDELLS, M. B., MACARTHUR, M. W., THORNTON, J. M. Nat. Struct. Biol. 1995; 2, 596–603. 34 FIEBIG, K. M., SCHWALBE, H., BUCK, M., SMITH, L. J., DOBSON, C. M. J. Phys. Chem. 1996; 100, 2661–2666. 35 SMITH, L. J., BOLIN, K. A., SCHWALBE, H., MACARTHUR, M. W., THORNTON, J. M., DOBSON, C. M. J. Mol. Biol. 1996; 255, 494–506. 36 GRIFFITHS-JONES, S. R., SHARMAN, G. J., MAYNARD, A. J., SEARLE, M. S. J. Mol. Biol. 1998; 284, 1597–1609. 37 JOURDAN, M., GRIFFITHS-JONES, S. R., SEARLE, M. S. Eur. J. Biochem. 2000; 267, 3539–3548. 38 NERI, D., BILLETER, M., WIDER, G., WUTHRICH, K. Science 1992; 257, 1559–1563. 39 FRANK, M., CLORE, G. M., GRONENBORN, A. Protein Sci. 1995; 4, 2605–2615. 40 BUCKLER, D., HAAS, E., SHERAGA, H. Biochemistry 1995; 34, 15965–15978. 41 SHORTLE, D. FASEB J. 1996; 10, 27–34.

42 KLEIN-SEETHARAMAN, J., OIKAWA, M., GRIMSHAW, S. B. et al. Science 2002; 295, 1719–1722. 43 SHORTLE, D., ACKERMAN, M. S. Science 2001; 293, 487–489. 44 De ALBA, E., RICO, M., JIMENEZ, M. A. Protein Sci. 1997; 6, 2548–2560. 45 WOUTERS, M. A., CURMI, P. M. G. Protein Struct. Funct. Genet. 1995; 22, 119–131. 46 HUTCHINSON, E. G., SESSIONS, R. B., THORNTON, J. M., WOOLFSON, D. N. Protein Sci. 1998; 7, 2287–2300. 47 ESPINOSA, J. F., MUNOZ, V., GELLMAN, S. H. J. Mol. Biol. 2001; 306, 397–402. 48 STANGER, H. E., SYUD, F. A., ESPINOSA, J. F., GIRIAT, I., MUIR, T., GELLMAN, S. H. Proc. Natl Acad. Sci. USA 2001; 98, 12105–12120. 49 COCHRAN, A. G., SKELTON, N. J., STAROVASNIK, M. A. Proc. Natl Acad. Sci. USA 2001; 98, 5578–5583. 50 KARSHIKOFF, A., LADENSTEIN, R. Trends Biochem. Sci. 2001; 26, 550–556. 51 KUMAR, S., NUSSINOV, R. Cell. Mol. Life Sci. 2001, 58, 1216–1233. 52 TAKANO, K., TSUCHIMORI, K., YAMAGATA, Y., YUTANI, K. Biochemistry 2000, 39, 12375–12381. 53 LOLADZE, V. V., IBARRA-MOLERO, B., SANCHEZ-RUIZ, J., MAKHATADZE, G. I. Biochemistry 1999, 38, 16419–16423. 54 CIANI, B., JOURDAN, M., SEARLE, M. S. J. Am. Chem. Soc. 2003; 125, 9038–9347. 55 RUSSELL, S. J., BLANDL, T., SKELTON, N. J., COCHRAN, A. G. J. Am. Chem. Soc. 2003; 125, 388–395. 56 MUNOZ, V., THOMPSON, P. A., HOFRICHTER, J., EATON, W. A. Nature 1997; 390, 196–199. 57 MUNOZ, V., HENRY, E. R., HOFRICHTER, J., EATON, W. A. Proc. Natl Acad. Sci. USA 1998; 95, 5872–5879. 58 HAGEN, S. J., HOFRICHTER, J., SZABO, A., EATON, W. A. Proc. Natl Acad. Sci. USA 1996; 93, 11615–11617. 59 DINNER, A. R., LAZARIDIS, T., KARPLUS, M. Proc. Natl Acad. Sci. USA 1999; 96, 9068–9073. 60 LACROIX, E., KORTEMME, T., LOPEZ DE LA PAZ, M., SERRANO, L. Curr. Opin. Struct. Biol. 1999; 487–493.

27

28 Design and Stability of Peptide $-sheets

61 SYND, F. A., ESPINOSA, J. F., GELLMAN, S. H. J. Am. Chem. Soc. 1999; 121, 11577–11578. 62 ESPINOSA, J. F., GELLMAN, S. H. Angew. Chem. 2000; 39, 2330–2333. 63 TATKO, C. D., WATERS, M. L. J. Am. Chem. Soc. 2004; 126, 2018–2034. 64 COLLEY, C. S., GRIFFITHS-JONES, S. R., GEORGE, M. W., SEARLE, M. S. J. Chem. Soc. Chem. Commun. 2000; 593–594. 65 HILARIO, J., KUBELKA, J., KEIDERLING, T. A. J. Am. Chem. Soc. 2003; 125, 7562–7574. 66 MA, B., NUSSINOV, R. J. Mol. Biol. 2000; 296, 1091–1104. 67 WANG, H., SUNG, S.-S. Biopolymers 1999; 50, 763–776. 68 HONDA, S., KOBAYASHI, N., MUNEKATA, E. J. Mol. Biol. 2000; 295, 269–278. 69 GRIFFITHS-JONES SEARLE, M. S. J. Am. Chem. Soc. 2000; 122, 8350–8356. 70 TATKO, C. D., WATERS, M. L. J. Am. Chem. Soc. 2002; 124, 9372–9373. 71 SHARMAN, G. J., SEARLE, M. S. J. Chem. Soc. Chem. Commun. 1997; 1955–1956. 72 SHARMAN, G. J., SEARLE, M. S. J. Am. Chem. Soc. 1998; 120, 5291–5300.

73 SCHENCK, H. L., GELLMAN, S. H. J. Am. Chem. Soc. 1998; 120, 4869–4870. 74 KORTEMME, T., RAMIREZ-ALVARADO, M., SERRANO, L. Science 1998; 281, 253–256. 75 De ALBA, E., SANTORO, J., RICO, M., JIMENEZ, M. A. Protein Sci. 1999; 8, 854–865. 76 DAS, C., RAGHOTHAMA, S., BALARAM, P. J. Am. Chem. Soc. 1998; 120, 5812–5813; DAS, C., RAGHOTHAMA, S., BALARAM, P. Chem. Commun. 1999; 967–968. 77 LOPEZ DE LA PAZ, M., LACROIX, E., RAMIREZ-ALVARADO, M., SERRANO, L. J. Mol. Biol. 2001; 312, 229–246. 78 FINKELSTEIN, A. V. Proteins 1991; 9, 23–27. 79 SYUD, F. A., STANGER, H. E., SCHENCK MORTELL, H. et al. J. Mol. Biol. 2003; 326, 553–568. 80 MACIAS, M. J., GERVAIS, V., CIVERA, C., OSCHKINAT, H. Nat. Struct. Biol. 2000; 7, 375–379. 81 KOEPF, E. K., PETRASSI, H. M., RATNASWAMY, G., HUFF, M. E., SUDOL, M., KELLY, J. W. Biochemistry 1999; 38, 14338–14351. 82 KANELIS, V., ROTIN, D., FORMAN-KAY, J. D. Nat. Struct. Biol. 2001; 8, 407–412.

1

Engineering Proteins for Stability and Efficient Folding Bernhard Schimmele, and Andreas Pl¨uckthun Universit¨at Z¨urich, Z¨urich, Switzerland

Originally published in: Protein Folding Handbook. Part II. Edited by Johannes Buchner and Thomas Kiefhaber. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30784-2

1 Introduction

The industrial, biotechnological, and medical applications of proteins are often limited by an insufficient protein stability or related problems. Such applications commonly require that proteins be produced on a large scale and remain stable enough to fulfill their functions for a reasonable length of time, often under harsh conditions. However, natural proteins are typically only marginally stable, and it is thus a major challenge for protein engineers to optimize stability and folding efficiency. The approaches that have been successfully employed to achieve this goal are rational design, semi-rational strategies based on sequence comparisons, and the methods of directed protein evolution. Of course, these methods are not mutually exclusive and can be combined to solve practical problems. All studies employing these methods have revealed important rules for protein engineering and at the same time shed light on the principles and mechanisms responsible for the folding and stability of proteins. Recent advances in stability engineering have demonstrated that merely small changes in a given protein sequence can have profound effects on its biophysical properties. The major challenge is therefore to correctly identify and remedy these shortcomings. It is the goal of this chapter to summarize the biophysical principles and technological approaches useful in improving the biophysical properties of proteins through sequence modification. Considering the enormous array of technologies involved in this endeavor, ranging from computer algorithms to selection technologies, it is not possible to give detailed experimental protocols in this chapter; instead, we will guide the reader to the cited literature.

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Engineering Proteins for Stability and Efficient Folding

2 Kinetic and Thermodynamic Aspects of Natural Proteins 2.1 The Stability of Natural Proteins

Evolution does not per se provide proteins with high stability. In fact, stability is just one of many evolutionary constraints on proteins. Proteins have to fold to a defined structure with adequate yield in a reasonable time and then have to be just stable enough to perform their function over a certain period. There is no evolutionary incentive to make a protein any “better” than what is needed to fulfill its cellular functions. In contrast, the use of a protein in a formulation at high concentration, its prolonged activity at 37 ◦ C, and its large-scale expression and crystallization, just to name a few conditions, may put far higher demands on the protein than its natural environment. Thus, the natural sequence may not be able to provide these properties, but a mutant sequence may. Proteins exist and have evolved in order to fulfill a given function, and evolution drives the structural properties of a protein mainly towards increased functionality [1]. In fact, most proteins are only marginally stable, with Gfolding in the range of −20 to −60 kJ mol−1 . It is still a matter of debate whether this marginal stability is actually a “design feature,” e.g., to allow degradation at a certain rate, whether it is caused by the selection pressure towards higher functionality that may not be compatible with high stability, or whether it is just a side effect of the lack of selection for high stability. Function is often linked to higher structural flexibility in certain regions of a protein. Lower stability as a result of this higher local structural flexibility might therefore simply represent an adaptation to increased functionality [2]. If this were generally true, however, stability engineering would fail in most cases, as it would not be able to reconcile stability with preserved protein function. An alternative, more optimistic view for the protein engineer is that marginal stability can be interpreted as a result of genetic drift [3]. In other words, lower stability is not intended; but it simply does not matter, provided that function is maintained. Random mutations occurring during evolution are more likely to destabilize the structure of a given parental protein sequence than to stabilize it or be neutral. However, as long as this stability decrease is not sufficient to render the protein nonfunctional, these destabilizing mutations are likely to accumulate in the sequence. This tendency has also been referred to as “sequence entropy” [4]. As a consequence, stability engineering could be interpreted as the art of identifying these unfavorable mutations in order to reestablish a more stable sequence. The concept discussed above also sets the basis for the consensus approach to stability engineering, which is discussed in Section 3.1 Based on the physical principles of protein folding and a structure-based analysis of the interactions between amino acid residues, rational design can give hints as to which residues need to be altered to achieve a desired effect, and this is discussed in Section 3.2. The third focus will be set on the methods of directed evolution, which mimic the

2 Kinetic and Thermodynamic Aspects of Natural Proteins

mechanisms of Darwinian evolution to evolve proteins with enhanced folding and stability properties (see Section 4). 2.2 Different Kinds of “Stability”

Before discussing in detail different strategies for rendering a protein more stable, it is worth taking a closer look at some basic features associated with protein stability and folding properties. The term “stability” itself is rather vague, and its precise meaning has often been adapted depending on the problem being addressed. This leads to different definitions of “stability.” We will now briefly analyze the definitions and differences between thermodynamic, kinetic, and thermal stability, as well as folding efficiency, which is also sometimes discussed in this context. Even though these properties are interconnected, they are not equivalent. This has important implications for protein engineering, as it is difficult to predict in advance how a given mutation will influence each of these properties. These influences will in fact be different for any protein under investigation according to the free energies of the native and unfolded states and the folding intermediates, as well as the folding pathway and the respective rate constants. 2.2.1 Thermodynamic Stability Thermodynamics describes the global unfolding behavior of a protein. The corresponding thermodynamic stability G describes the differences between the free energies of the native (N) and the unfolded (U) states. Importantly, G is an equilibrium property for a reaction involving two or more states. The simplest model of an unfolding reaction is the equilibrium of a two-state unfolding reaction:

where kunfolding and kfolding are the rate constants of the respective unfolding and folding reactions. As a consequence, values for G can be deduced only if the described process is fully reversible and no intermediate states are populated to a significant extent, unless they are explicitly known and measured. G describes to what degree the two states are populated at a given temperature according to G unfolding = −RT In K

(1)

where K is the equilibrium constant of the unfolding process. The treatment of experimental data is much simplified in such a model, as stability is affected only by the free energies of the folded and unfolded states. Because G also provides an overall measure of all energetic contributions of interactions occurring upon protein folding, it is a convenient quantity for comparing the energetic effects of single amino acid replacements in a given folded structure. Because of the great scientific interest in deducing the principles of protein folding on a quantitative

3

4 Engineering Proteins for Stability and Efficient Folding

basis, many studies have been carried out with carefully selected model systems in which these strict requirements that allow the determination of G are fulfilled. Moreover, G represents an intrinsic quantity, at a given temperature in a given buffer, that is independent of the experimental setup and therefore reproducible in any case. The major problem in such studies, however, is confirmation that the observed transition can in fact be described as a fully reversible process, and denaturant-induced transition data alone are often not sufficient to allow reliable judgment. Only if data derived from measurements on the basis of different spectral probes (or better yet, additionally by differential scanning calorimetry, yielding the model-independent enthalpy change of unfolding) agree with each other are the deduced G values likely to be correct. Nevertheless, many useful conclusions can be drawn about the effects of mutations even if these strict requirements cannot be completely fulfilled in all cases. Another fact complicates the use of G as a measure for protein stability. Mostly as a result of the fact that the hydrophobic effect is the major driving force of protein folding [5], G itself is a characteristic, curved function of temperature. It is defined as G(T ) = G U −G N = H(T )−T S(T ) = −RT In K

(2)

emphasizing especially that the enthalpy change H is itself a function of temperature. This change of H with T can be described by the heat capacity change C p =

∂H(T ) ∂T

(3) p

Although often ignored, the temperature dependence of G should thus not be left out of the account if one is dealing with the stability engineering of proteins. The large heat capacity change upon the transfer of nonpolar solutes to water, which is the basis of the hydrophobic effect, results in a curved function of G versus temperature. By using the definition of Cp , G can be approximated as a function of T, the melting temperature T m , the enthalpy change at T m H(T m ), and Cp . G(T ) = (1−

T T )H(Tm ) + (T −Tm )C p −T C p In Tm Tm

(4)

If G(T) is plotted as a function of T, the curve increases at low temperatures and decreases at high temperatures (Figure 1). The temperature at which folded and unfolded states are equally populated, and thus G = 0, is called the melting temperature T m . The respective curvature of G versus temperature is strongly dependent on the change of heat capacity Cp upon unfolding. A mutation may change Cp , H(T m ), and T m in any combination, thereby altering the shape and the position of this curve. Higher thermodynamic stability at a given temperature (G(T)) can

2 Kinetic and Thermodynamic Aspects of Natural Proteins

Fig. 1 The complex relationship between the melting temperature and the free energy of folding. Schematic representations of the free energy difference between the folded and unfolded states of a protein, GN , as a function of temperature, T. A typical protein is shown in curve 1 (–––). The shape of this curve changes if a mutation affects C p ; H(T m ), and/or T m of the protein. In the examples shown, not just one but several of these parameters are changed. Higher thermodynamic stability at a given temperature can phenomenologically be the result of a curve upshift (. . .. . .. . .). Right-shifting the

curved function (----) results in a right-shift of the maximum of the GN function as well as in a shift of the melting temperature T m towards higher temperatures (T m ). If the GN function is flattened because of a small C p (. . .), a higher melting temperature (T m ) can result, even though the thermodynamic stability is actually decreased over a broad temperature range. All depicted curves represent extreme cases, and typically a combination of the described alterations will occur upon mutating a protein.

thus be achieved, e.g., by an upshift of this curve with constant maximum. Rightshifting of the curve will decrease G at lower temperatures but increase it at higher temperatures, which goes along with an increase of T m . Flattening of the curve due to a lower change of heat capacity upon unfolding may cause lower thermodynamic stability at most temperatures, even though T m is increased. These considerations contain important implications for the analysis of engineered mutants. First, a measured decrease of G at a certain temperature does not necessarily mean that at higher temperatures the thermodynamic stability might not have been increased. Second, by determining the change of G, no conclusions can be drawn about a potential change of the melting temperature T m . While they are typically related, a low G at a given temperature does not necessarily mean that T m will be decreased. T m and G must therefore be regarded as different properties. It should also be recalled that thermodynamic parameters, while easily reproduced, give no information about how long it will take until equilibrium is reached or, in the case of a nonreversible reaction, until a certain fraction of proteins is inactivated. Thus, thermodynamic stability does not necessarily provide information about whether a protein will meet the stability requirements for an intended application. T m itself often serves as a measure of protein stability, more precisely of thermal stability. In the literature, the expression “thermal stability” is again used with different meanings. In the described case, it represents the melting temperature for a reversible process. However, a complete thermodynamic analysis of the vast majority of proteins is not possible, because either intermediate folding states are

5

6 Engineering Proteins for Stability and Efficient Folding

populated or folding in the absence of denaturants that solubilize the unfolded state is not a fully reversible process. As will be discussed in the following section, thermal stability can nevertheless serve as a very practical means of describing protein stability, defined either as the transition temperature of an irreversible process or as the half-life of a protein under a given set of conditions. 2.2.2 Kinetic Stability In most cases, outside the biophysical research lab, protein unfolding is an irreversible reaction. Initially, the aggregation of unfolded molecules or of folding intermediates prevent the back-reaction of folding, and this is followed, after prolonged times at high temperature, by chemical inactivation or, in impure samples, proteolysis. In a simple model, kinetic inactivation can be described as

As discussed in the Introduction, one evolutionary constraint on proteins is that their three-dimensional structure remains viable for a certain period of time. In the case of non-equilibrium conditions and irreversible reactions, reaction kinetics becomes the important parameter. The folded state can resist high temperatures for a considerably long time if either the rate constant of inactivation or the rate constant of unfolding is sufficiently low. The rate constants are determined by the free energy of activation, e.g., the difference in energy between the folded state and the transition state of unfolding. Proteins can thus be kinetically stabilized by increasing this activation barrier. This kinetic stability is different in some crucial aspects from the thermodynamic stability mentioned above. Engineering of proteins for enhanced stability has to deal with both aspects. The best mutations for enhancing kinetic stability will not necessarily be the best mutations for enhancing thermodynamic stability, and not every mutation that increases thermodynamic stability will automatically have a positive effect on protein half-life. Kinetic stabilization is a common theme in nature, and there are several indications that many proteins from thermophilic organisms are indeed stabilized kinetically rather than thermodynamically [6, 7]. Another important example is that of proteins from the coats of viruses and phages that have to protect their genetic material under very adverse conditions [8]. In extreme cases the native state of a protein can even be less stable than its denatured state, but the native fold can still be kinetically trapped, and large kinetic unfolding barriers can provide the protein with an extremely long half-life [9]. There are different ways of describing and determining kinetic stability. Even if the reaction proceeds in an irreversible way, a practical “melting curve” can still be determined, and the observed transition is cooperative. The midpoint of this transition can serve as a practical means to compare the thermal resistance of different protein variants. Alternatively, one can use the half-life of the protein at a given temperature as an empirical means of stability. However, one has to be aware

2 Kinetic and Thermodynamic Aspects of Natural Proteins

that this will not reflect equilibrium conditions. The observed values are actually kinetic values and thus are not independent of the exact experimental conditions, such as the protein concentration, the heating rate, etc. A more thorough analysis of kinetic stabilization must include kinetic measurements to determine the respective unfolding rates at different temperatures [10, 11]. For medical applications, engineered proteins usually have to fulfill defined stability requirements such as the absence of aggregation in the formulation used, long-term stability, and prolonged activity at 37 ◦ C. The mere analysis of thermodynamic and kinetic parameters under defined reaction conditions in vitro is not always a reliable indicator of protein behavior under in vivo conditions. Mechanisms other than the intrinsic properties of the protein, such as proteolysis or aggregation with other proteins, can affect half-life. For practical utility, the half-life can therefore also be determined by measuring the percentage of molecules that retain their function after incubation under the respective conditions, e.g., in human serum at 37 ◦ C for several days [12]. 2.2.3 Folding Efficiency A different issue of kinetics is related not to unfolding but to the folding of the protein in vivo. More precisely, the question is, which percentage of a protein will actually fold to the native state, as opposed to going to misfolded states, soluble aggregates, or inclusion bodies? Even though the folding efficiency in vivo is not directly related to stability, correlations can often be observed [13]. Additionally, the efficiency of protein folding in vivo is usually the predominant factor influencing the expression yield and is therefore also crucial for large-scale production of functionally intact proteins. Importantly, many mammalian proteins with a high potential for medical applications, especially those secreted or expressed on the cell surface, can rely on the complex folding machinery of the eukaryotic cell to reach their final native state, and the secretory quality-control system of eukaryotic cells allows discrimination of proteins by their folding behavior [14]. Moreover, they usually do not need to be expressed in high amounts in their native physiological context. The resulting lack of selection pressure on their efficiency of folding during evolution is likely to be one of the causes for the difficulties often observed when attempting their overexpression. Unfortunately, these tend to be the proteins of greatest pharmacological utility. Despite the fact that thermodynamic stability underlies folding efficiency, the kinetic partitioning into productive folding or aggregation is influenced by many different factors. For illustrative purposes, we can again describe this by a very simple scheme:

Folding intermediates are often the source of aggregation, and the overall folding efficiency will therefore depend mainly on the nature of these intermediates. This includes the free energy of the folding intermediates themselves, as well as their

7

8 Engineering Proteins for Stability and Efficient Folding

half-lives and any efficient pathways to aggregation. in vivo, the situation becomes much more complex, as additional parameters such as interactions with cellular components and chaperones or degradation by host cell proteases come into play. The final output of a properly folded protein will therefore depend on all of these kinetic competitions [15]. Protein expression in the bacterial cytoplasm in many cases shows correlations between soluble expression yield and thermodynamic stability of the protein [16, 17]. Additional complications can arise from transport steps. For example, the expression of proteins in the periplasm of E. coli is dependent on the prior transport of the polypeptide chain through the inner membrane, and the folding yield is subsequently influenced by the folding and aggregation reaction in the periplasmic space as well as by interactions with periplasmic factors such as chaperones and proteases. In some cases, mutations that show positive effects on in vivo folding yield have no influence on the overall stability of the protein [18]. Conversely, mutations that strongly increase thermodynamic stability sometimes result in lower folding yields [19]. Nevertheless, many mutations act synergistically on both properties, because they are likely to reduce the free energy of the folded state as well as the free energy of folding intermediates and thereby lower the energetic activation barrier to folding [13].

3 The Engineering Approach 3.1 Consensus Strategies 3.1.1 Principles As discussed in the preceding sections, marginal protein stability is likely to be a side effect of “sequence entropy” occurring during natural evolution, because the major driving force of evolution is positive selection towards an enhanced functional property, while stability has to be maintained at only a minimum level to secure function. Mutations are likely to occur in a random fashion during this process; the probability that a mutation will have a stabilizing effect on the protein is very low, whereas the probability that the mutation will have destabilizing effects is very high. However, as long as the remaining amino acid sequence is still able to fold into a given structure and the overall domain stability does not fall below a certain threshold, the resulting protein sequences will not be eliminated during the course of evolution [20]. Destabilizing mutations are therefore often selectively neutral and thus accumulate in a given parental sequence. The same should also be true for folding efficiencies. Most proteins are not needed at high concentrations or may even become harmful to the organism in such a case. Similar to stability, folding yield is selectively neutral, provided that the minimum level for cellular function is maintained. This sets the basis for a semi-rational approach to protein stabilization, which is called the consensus approach [21] and is based on sequence statistics. Because mutations occur randomly, the distribution of amino acids at a given position in

3 The Engineering Approach

a set of homologous proteins can be described, in a very crude approximation, by Boltzmann’s law. The consensus approach assumes that at a specific position in a sequence alignment of homologous proteins, the contribution of the respective consensus amino acid to the stability of the protein is on average higher than the contribution of any non-consensus amino acid. Replacement of all non-consensus amino acids in a sequence by the respective consensus amino acid should therefore increase the overall stability of the protein. Obvious advantages of the consensus approach are that it is comparatively simple and is not strictly dependent on structural information at high resolution. The prerequisite for building a non-biased consensus is the availability of sequences homologous to the protein under investigation. The number of sequences should be large enough to make the sequence statistics reliable and to exclude bias in the resulting consensus sequences. Figure 2 shows an alignment of homologous sequences of single repeat modules, the smallest structural entity of a class of proteins known as leucine-rich repeat (LRR) proteins. Because the length of these modules varies among the different classes of LRR proteins – influencing their topology – only repeat modules of a length of 24 amino acid residues have been used for this alignment. The probability of each amino acid occurring at a given position is calculated to derive a consensus sequence, representing the most frequently occurring amino acid residue at each position. The distribution of residues at each position can provide information on structurally forbidden residues and allows weighing the consensus with respect to variability. In most cases, the consensus will contain the residues important for defining the structure of the proteins. In the case of an enzyme family, it will also include the “functional” residues, i.e., those of the active site. In the case of binding proteins, such as antibodies or repeat proteins, the “functional” residues (those involved in binding) are not conserved but are different for each individual molecule, which has to adapt to its target. Although at first sight no sophisticated structural analysis seems to be required, this is true in only the simplest of cases, where a single family of related sequences can be represented by a single consensus. Frequently, multiple families have emerged that use mutually incompatible solutions of packing. A good example is that of antibodies for which subgroup-specific consensus building has been very fruitful [22]. Averaging over all families would simply yield the consensus of the most-represented family and, if they are equally represented, may result in mutually incompatible residues. Thus, structural analysis can be very helpful in deciding whether an “averaging” of different sequence families is permissible or not. Because of this problem of interacting residues, a simple averaging may lead to incompatible pairs; therefore, these residues should be changed only as groups. The danger of disrupting these interactions by substitutions with consensus residues is especially high in cases where a very broad set of sequences is used for the alignment. To minimize this risk, the sequence statistics can be extended by analysis of covariance in order to derive probabilities that describe the joint occurrence of amino acid residues at two defined positions [23]. As explained above, before deriving a consensus, the aligned sequences can be divided into subclasses, which are likely to contain interacting pairs or groups of residues. Certain

9

10 Engineering Proteins for Stability and Efficient Folding

Fig. 2 Example of the analysis for deriving and analyzing a consensus sequence. (a) From a sequence alignment of 3077 sequences of 24 amino acid LRR motifs of the Pfam database (http://www.sanger.ac.uk/Software/Pfam/),

the relative frequencies of amino acid residues at each position are calculated. (b) Based on the relative frequencies calculated from the alignment, a consensus sequence can be derived representing the most frequently occurring residue at each position.

3 The Engineering Approach

variations of residues involved in salt bridges, distinct hydrogen-bonding patterns, or packing of the hydrophobic core are characterized by complementary changes between these subclasses, with mutations to a certain residue at one position being compensated by a mutation to a complementary residue at another position. The subclasses can be built either by being based on sequence homology alone or by including structural information if available. Even though the definition of these subclasses is always dependent on the homology cutoff set by the investigator, a simple dendrogram analysis can be used to group the complete set of sequences into distinct families. Additionally, by building the consensus sequence of each family separately, followed by a comparison of these consensus sequences, distinct structural features of each group can be recognized in some cases. The consensus concept has been applied successfully to a large variety of proteins and structural motifs to date. Important lessons for an effective application of the consensus approach can be learned from these studies, and we will therefore give a few examples to briefly discuss some of the advantages and limitations of the method. 3.1.2 Examples There is always the concern that the stabilizing and destabilizing effects of introduced mutations will counterbalance each other and that the overall change in protein stability will be small. Steipe et al. [20] applied the method for the first time on immunoglobulin VL domains and predicted 10 potentially stabilizing mutations. Six mutations were indeed stabilizing, three had no effect, and one was destabilizing. When applied to GroEL mini-chaperones, 34 predicted amino acid replacements were individually checked, out of which 13 were stabilizing, five showed no effect, and 16 were destabilizing [24]. Lehmann et al. [25] extended the approach to an entire protein. In a first set of experiments, 13 homologous sequences of

←-----------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------Fig. 2 (Continued) The color code is given on the right hand side of the panel. Residues occurring in more than 80% of all sequences at the respective position are colored in red. Based on these values, consensus sequences with a given homology threshold can be derived (not shown), i.e., with a higher threshold, more positions will be “undefined.” (c) Preferences at a given position are reflected not only by the absolute frequency but also by the total number of different amino acid types occurring at each position (residue variability). As an example, the relative frequencies of each amino acid are shown for the highly conserved position 2 and the highly variable position 4. By plotting the relative frequencies of amino acids at a particular position,

preferences for certain amino acid types as well as “forbidden” residues can be identified. At position 2, a strong preference for leucine can be observed, and the occurrence of other residues is restricted mostly to hydrophobic side chains. At position 4, all residues are “allowed” except for proline, which is “disallowed” due to secondary structure propensity reasons. (d) By normalizing the relative frequency of the consensus amino acid with the number of “allowed” residues at a given position, a modified consensus sequence can be deduced. The variability V is calculated according to V = 100×N/F, where N is the total number of different residue types occurring at each position and F is the frequency of the most frequent residue in percentage.

11

12 Engineering Proteins for Stability and Efficient Folding

a fungal phytase were used to build the consensus, and the resulting consensus enzyme showed an increase of 15–22 degrees in unfolding temperature and an increase of the temperature optimum for catalysis of 16–26 degrees compared with each of its parents. In a second set of experiments, additional mutations were predicted by simply adding more sequences to the alignment. By checking the effects of the individual mutations on thermal stability and combining mutations with positive effects, the unfolding temperature could be increased by an additional 21 degrees to 90 ◦ C. No loss of catalytic activity of the enzyme was observed in any case. This work showed that the number of sequences used for the alignment is indeed an important factor because it might help to optimize ambiguous positions. In addition, the observed large increase in thermal stability could not be attributed to the effect of one single amino acid substitution but rather to the synergistic effects of many replacements. Even though these and other studies show clearly that the consensus approach allows one to predict stabilizing mutations with a rather high success rate in a rapid way, the effect of each predicted mutation carries some uncertainty, and it is possible that some may contribute destabilizing effects that can counterbalance the stabilizing ones. However, the effects of stabilizing mutations were often found to be additive. Therefore, instead of examining each mutation individually, it is often useful to combine groups of “rather certain” mutations and others that are more speculative. The application of the consensus concept to families of repeat proteins [23, 26–28] represents, in some respects, a special case due to a number of favorable features. It can nevertheless illustrate the importance of some principles of the approach. The non-globular fold of repeat proteins consists of repeated motifs of 20–40 amino acids. Several results indicate that consensus repeat proteins are indeed much more stable than natural repeat proteins [29, 30]. Repeat proteins might represent an extreme case in which the principles of the consensus approach become very apparent. The structural entities (the repeat modules) are small and thus each protein contributes several repeats to the databases; the number of available sequences consequently becomes very large compared with other proteins. Therefore, a consensus sequence for a single repeat module can easily be assigned and ambiguous positions will occur with lower frequency in the statistical output (Figure 2). In addition, interactions that are present within or between several repeats will add to the free energy of folding multiple times, while problem spots would equally be potentiated. Therefore, effects on stability are likely to be consecutively added by introducing additional modules to the array. Even though more-detailed analyses are still needed, initial results pinpoint some of the structural reasons for the stability gains observed upon building a consensus in each of these studies. The regular arrangement of structural motifs gives rise to a more regular H-bonding pattern with a higher number of inter- and intra-repeat H bonds [30]. Loop insertions in natural repeat proteins that are likely to result in more flexible local regions are removed, thereby eliminating local centers of unfolding. In the left-handed helical and disallowed regions of the Ramachandran plot, glycine residues are always present in the consensus proteins, while they are avoided in other places by this design, where their flexibility is not needed and may be harmful to stability.

3 The Engineering Approach

3.2 Structure-based Engineering

Structure-based engineering relies on a detailed analysis of 3D structures, followed by site-directed mutagenesis. We avoid the term “rational” engineering, as it would elicit expectations of perfect predictability and implicitly suggest that all other approaches are free of logical reasoning. In structure-based engineering, positions have to be identified at which suboptimal amino acids in the original sequences lead to a loss of stability. Subsequently it needs to be specified which amino acids should be introduced as a replacement. The ab initio prediction of protein structure, however, is still not a feasible task due to the multitude of potential interactions within the protein and between protein and solvent, which leads to an extremely high number of possible conformations and intermediates of similar energy [31]. Hence, a prerequisite for structure-based engineering is, next to experimentally determined structures, usually the existence of a large experimental data-set within a group of structural homologues that can be used as a basis for predictions. High-resolution structures are necessary to allow the estimation of possible conformational, energetic, and steric influences upon replacement of particular amino acid side chains and can thus help to avoid unfavorable strain in the resulting mutants. Because of the present efforts in the field of structural genomics, these structure-based approaches are likely to become even more important in the future. With this structural information in hand, the goal of designing more stable variants is then to pinpoint particular regions and positions associated with possible stability defects and to subsequently find a better solution to the problem. In contrast, semi-rational approaches like the previously discussed consensus concept are rather crude methods for introducing stabilizing features. Structural and energetic analysis can be used to reexamine the changes proposed by the consensus approach and to fine-tune the system to reintroduce structural features that might have been lost in the averaging process. Proteins of hyperthermophilic organisms have been of special interest for examining the structural mechanisms of thermostabilization and have been contemplated as guides for the engineering of “problem” proteins for better properties. From a phenomenological point of view, the basis of increased thermostability is frequently set by a flattened G-versus-T curve that is due to a smaller change of heat capacity upon unfolding (Figure 1) or by a kinetic stabilization that is due to a strong decrease in the rate of unfolding. The crucial question is, however, what the molecular differences are that give these proteins their favorable properties. Genome-wide comparisons between hyperthermophilic and mesophilic organisms with respect to amino acid composition did not yield any obvious common rules of how these effects are achieved [32]. Therefore, hope was placed on the increasing number of pairwise high-resolution structure comparisons of thermophilic proteins with their mesophilic counterparts. While they provided a more differentiated picture, a “global” rule still could not be derived. A few highly specific mutations are often enough to provide considerably stabilizing effects, but the additive effect of many small contributions, none of them dramatic by itself, may be the usual case.

13

14 Engineering Proteins for Stability and Efficient Folding

Moreover, rather than relying on one universal strategy, nature utilizes a variety of strategies for the thermal adaptation of proteins [33]. In fact, the list of stabilizing structural features in hyperthermophilic proteins reflects the diverse principles of protein stability and folding that protein engineers try to exploit and that will be discussed in this section. High-resolution structures of thermophilic proteins can thus provide a detailed view of how nature implements these principles to create proteins of higher stability [34]. However, the lack of a unifying “rule” and the multitude of strategies nature uses provide an important lesson for protein engineers. Depending on the protein under investigation, the strategy of choice can be different, and even for a given protein there may be more than one optimal solution to the problem. Before choosing from the available set of strategies, the focus should therefore be on identifying potential “weak points” responsible for stability defects in a given protein structure. We will now discuss some structural features associated with protein stability as well as strategies for altering these features towards more favorable biophysical properties. The given list of course lays no claim to completeness but should point out some important principles. Any replacement in a given sequence may have multiple effects on protein stability, and destabilizing effects can often outweigh the stabilizing ones. An assessment of potential destabilizing effects is therefore crucial. Wherever possible, references are made to the different forms of “stability” discussed in the Introduction. In the case study provided at the end of this section, an example will be given to demonstrate how consensus approaches and structural analysis can be combined to yield useful results. 3.2.1 Entropic Stabilization An obvious strategy for increasing the free-energy difference between the folded and unfolded states, and thus the thermodynamic stability, is to decrease the entropy of the unfolded state. The underlying concept is to decrease the flexibility of the polypeptide chain, usually by introducing an additional intrachain linkage. Such entropic stabilization has become a common strategy for protein engineers. The prerequisite for success is that the mutations rendering the unfolded protein less flexible do not introduce unfavorable strain in the folded three-dimensional structure or result in any steric incompatibilities [35]. We now discuss several ways to achieve an entropic stabilization. 3.2.1.1 Introduction of Disulfide Bridges The introduction of additional disulfide bridges is a straightforward way of establishing an intrachain linkage to reduce the entropy of the unfolded state [36, 37]. The magnitude of the entropic effect is thought, as a crude approximation, to be proportional to the logarithm of the number of residues between the two bridged cysteines [38]. The spatial distance between the residues to be replaced with cysteines has to be evaluated with care in the model of the folded protein in order to prevent perturbations of the native structure upon formation of the disulfide bridge. It should be noted, however, that the energetic effect of additional disulfide bridges is not only entropic but also of a far more complex nature, giving rise to entropic as well as enthalpic contributions

3 The Engineering Approach

to the change in the free energy of folding [39, 40]. For example, an additional decrease in the free energy of folding can result from the reduced solvation energy of the unfolded state [40]. In contrast, a reduced solvation energy of the folded state would have the opposite effect, while residual structure in the denatured state would again push the equilibrium to the side of the folded protein. Because the disulfide bond itself is hydrophobic in nature, it is often engineered into the interior of the protein. This is not an easy task, as it can negatively affect core packing. Even though there are several examples of successful protein stabilization by introducing artificial disulfide bridges [41, 42], the complex energetic effects can also cause a destabilization of the protein [43]. Furthermore, the introduction of additional cysteines often results in a rather drastic decrease in folding efficiency, because incorrect and intermolecular disulfide formation can remove large portions of expressed protein by aggregation. Since disulfide formation does not occur in the cytoplasm, secretion to the bacterial periplasm or to the eukaryotic ER is required for functional expression, usually associated with lower yields than for the production of cytoplasmic proteins. Alternatively, if the protein is produced by refolding, redox conditions have to be adjusted, which can be difficult if the native protein also contains free cysteines. 3.2.1.2 Circularization An alternative approach with the same underlying concept is the circularization of proteins by fixing the loose N- and C-termini via a peptide bond. In addition to the entropic effect, the fixing of the loose ends can prevent local unfolding events occurring at the termini and thereby kinetically stabilize the native structure. With the discovery of inteins, which mediate protein-splicing reactions, a tool that allows the directed formation of peptide bonds between ends fused to different parts of the intein became available. Intein-mediated protein ligation has been used to covalently link the termini of β-lactamase, a protein that is especially amenable to this strategy due to the close proximity of its N- and C-termini [44]. In accordance with polymer theory, the thermal stability of the protein was enhanced by about 6 degrees, from 45 ◦ C to 51 ◦ C. For circularized DHFR [45] an increased half-life at elevated temperature was observed. The close proximity of the termini is of course a prerequisite for this procedure, and the stabilizing effect is likely to become marginal if the loose ends are linked via long unstructured loops. Similar to the situation upon introduction of artificial disulfide bridges, destabilizing enthalpic effects may negate the favorable entropic contribution [46]. In addition, low protein ligation efficiencies and difficulties in separating circular from linear forms of the protein often cause additional technical challenges. It remains to be seen whether this technology is robust enough for biotechnological or biomedical applications. 3.2.1.3 Shortening Solvent-exposed Loops Short, solvent-exposed loops are rather fixed in the native state, but a comparably large number of additional conformations become accessible in the unfolded state, while long loops have a large number of conformations also available in the native state. Thus, the shortening of loops should in principle lead to a relative decrease in the loss of conformational entropy upon folding. Conversely, increasing the loop lengths by insertion of glycine residues into

15

16 Engineering Proteins for Stability and Efficient Folding

the loops of the four-helix bundle protein Rop has indeed resulted in a strong and continuous decrease in thermodynamic stability [47]. In addition, loop shortening can have the effect of abolishing hot spots of local unfolding events and may result in kinetic stabilization. Even though it has become obvious that loop shortening or tying down of loops by external interactions is a common theme in thermostable proteins of thermophilic organisms [48, 49], the strategy is often hard to realize for a given protein target, as the danger of introducing additional strain in the native state is high, and solvent-exposed loops are often important with respect to function. 3.2.1.4 Reduction of Chain Entropies By considering the conformational entropies of amino acid side chains, another strategy for decreasing the entropy of the unfolded state becomes apparent. Because of the five-membered-ring nature of the proline side chain, it not only restricts the possible conformations of the preceding residue but also can adopt only a few conformations itself. It therefore has the lowest conformational entropy of all amino acids [50]. In contrast, glycine, which has no side chain, has the highest conformational entropy. Substitutions of non-glycine residues with proline or the replacement of glycines by other residues should therefore reduce the entropy of the unfolded state. Positions that allow substitutions with proline are, however, very rare. Because proline is poorly compatible with α-helices and incompatible with β-strands, the position of a new proline must not be part of these secondary structure elements. At most positions in the native structure, the respective torsion angles will be incompatible, and the mutations are thus very much restricted to loop and turn regions. Again, care should be taken not to remove any favorable interactions of the replaced amino acid side chain [51]. In order to examine in advance whether the respective site is permissive for a substitution with proline, the dihedral angles of the site can be checked and should lie in the range of φ/ −50 to −80/120 to 180 or, alternatively, −50 to −70/−10 to −50. Similar restrictions apply for the replacement of glycines with any other residue. In many cases this will create steric overlaps, and such negative structural crowding effects can outweigh the positive energetic benefits. 3.2.2 Hydrophobic Core Packing Exposed residues are often directly involved in ligand or substrate binding and therefore often play a functional role. In contrast, the residues of a protein’s interior usually play mostly a structural role, and the associated hydrophobic effect is thought to be the main driving force of protein folding and thermodynamic stability. In known structures the core residues fill almost the entire interior space, provide many favorable van der Waals interactions, and maximize hydrophobic stabilization by exclusion of the solvent. In principle, an increase in thermodynamic stability of 4–8 kJ mol−1 can be achieved for each additional methylene group buried [52]. Paradoxically, the importance of the hydrophobic effect for folding and stability of proteins simultaneously limits its applicability for protein engineering. Because

3 The Engineering Approach

the hydrophobic core is already densely packed in almost all native proteins, most changes here will create over-packing or packing defects, causing an overall destabilization rather than an improvement in stability. In addition, even subtle changes of core residues can lead to a rearrangement of external residues and thereby alter the functional properties of the protein [53]. Care should also be taken to avoid the introduction of conformational strain by the mutation of core residues, as destabilizing effects from a strained conformation can sometimes compensate the energetic gain of an increase in buried hydrophobic volume [54]. Improvement of core packing must therefore be based on analyses made from high-resolution structural information in combination with sequence comparisons. This allows one to specifically look for cavities in the core that indicate imperfect packing. If the hydrophobic surface area around the cavity is large, additional van der Waals interactions can be provided by the introduction of sterically fitting alkyl or aryl groups from hydrophobic side chains, thereby decreasing the size of the cavity [55, 56]. 3.2.3 Charge Interactions Oppositely charged amino acid residues, if appropriately positioned, have the potential to form salt bridges, whereas like-charged residues lead to repulsions. The magnitude of the effect of charge-charge interactions on overall protein stability is still a matter of discussion [57]. In the case of ionic interactions between side chains buried in the hydrophobic core, the high energetic cost of transferring charged ions from aqueous solution to the low-dielectric interior of the protein also has to be taken into account. If a single charge were buried in a protein, which would be extremely rare in a natural protein, the design of an ion pair would be very attractive. However, if a hydrophobic pair were to be replaced by an ion pair, the resulting energy would have to be higher than the loss of the previous pair plus the cost of burying the charge. Nevertheless, the high contribution of buried salt bridges to the overall stability of the native protein structure underlines their potential for introducing additional stability [58]. The optimal spatial arrangement of the interacting side chains and their respective charges is, however, crucial. Moreover, buried charged side chains are often not only part of interacting charge pairs but also part of complex charge clusters built from many side chains, which are able to magnify the effect. Similar rules apply for the interactions of charged residues on the protein surface. Only the perfect arrangement of charges seems to be able to make up for the desolvation penalty that has to be paid upon formation of a salt bridge. The effect on thermal stability, however, can be drastic. Increasing the free energy of unfolding by the changing of charges includes maximizing the number of salt bridges and, equally important, the removal of repulsive interactions [59], which are not uncommon in natural proteins. Predictions on a structural basis can be difficult due to the often higher flexibility of side chains on the protein surface, but simple models that allow predictions about potential stabilizing and destabilizing surface charges can be used [60]. The key is to consider not only nearest neighbors but also a whole network of charges that have to optimally interact and avoid repulsions.

17

18 Engineering Proteins for Stability and Efficient Folding

Because the surface charge distribution can have a huge impact on stability, but is defined by many residues at different positions, these residues are also a valuable target to be combined with selection techniques as discussed in Section 4. A special case of electrostatic interaction is the “helix dipole.” By reducing the net partial charges at the helical ends through placement of side-chains, which productively interact with the helix dipole, the helical structure is stabilized [61]. Introduction of negatively charged residues at the N-terminal end and positively charged residues at the C-terminal end leads to this stabilizing effect. The provided stability gain is, however, marginal (less than 4 kJ mol−1 ). 3.2.4 Hydrogen Bonding There was no initial reason to believe that intrachain hydrogen bonds in the native state would be more energetic than those of the unfolded chain to water [5]. By including terms of entropy change of the solvent and additional van der Waals interactions upon polar group burial, however, the positive contribution of hydrogen bonding to protein stability has become generally accepted [58, 62]. Despite this ongoing discussion, hydrogen-bonding patterns are a highly valuable target for the stability engineering of proteins. Because engineering deals with improving folded proteins, the major concern has to be how to satisfy the existing hydrogen-bonding network in a structural context. The basis for this endeavor is structural information of high resolution, and the most lucrative goal is to identify potential residues that represent buried but unsatisfied donors of hydrogen bonds. Site-directed mutagenesis of a nearby residue to provide a hydrogen-bonding acceptor can cause a stability gain in the range of 2–10 kJ mol−1 , depending on the geometry and other compensating effects [63, 64]. A special structural context that can provide significant additional stabilization by either hydrogen bonds or ionic interactions is the anchoring of relatively loose structural elements like loop structures or the N- and C-termini, thereby tightening “hot spots” of local unfolding [49, 65]. 3.2.5 Disallowed Phi-Psi Angles The stereochemistry of the polypeptide backbone can be defined by the dihedral angles φ and , and any individual residue in a structure is defined by a single set of φ, values. For conformational analysis of protein structures, the Ramachandran plot representing the dihedral angle space is an excellent starting point [66]. The φ, values of amino acid residues in protein structures usually reside in three preferred or “allowed” regions of the Ramachandran plot, called right-handed helical, extended, and left-handed helical. The right-handed and extended conformations correspond to α-helix and β-strand secondary structures, respectively, and the vast majority of non-glycine residues lie within these two regions. The left-handed region corresponds to structural features at the termini of secondary structure elements and describes regions involved in the reversal of the polypeptide chain. There is a high preference in this region for glycine residues, as the β-carbons of non-glycine residues can sterically interact with the polypeptide backbone, resulting in unfavorable energies. In some cases, the substitution of non-glycine residues

3 The Engineering Approach

by glycines in the left-handed helical region increases thermodynamic stability (up to 8 kJ mol−1 in RNase H) [67]. Although the strict introduction of glycines in such positions is an important point to consider for de novo protein design, it does not represent a general rule for stabilizing the native states of proteins. In some cases, the energy penalty for the accommodation of unfavorable strain can be offset by lost unfavorable or new favorable local interactions, such as hydrogen bonding or hydrophobic interactions. Some replacements of this kind can therefore even lead to destabilization rather than stabilization [68]. The same rules apply in principle to the disallowed regions of the Ramachandran plot. Steric clashes result in a high energetic cost in the folded structure, especially for non-glycine residues in all disallowed regions. Stabilizing mutations to glycine have been introduced with energy gains of up to 18 kJ mol−1 [69]. It has been noted, however, that certain non-glycine residues also have propensities to occur in the disallowed regions – such as Asn and Ala in the type II turn region– and the energetic cost of their occurrences is often low [70]. Other residues found in the disallowed regions are small polar residues [71] that compensate for the energy cost by making additional hydrogen bonds. Unfavorable conformations often occur in very short loops [72], where the rest of the structure may constrain the loop efficiently, and interactions with the solvent may also offset the energy costs. Conformational analysis by the Ramachandran plot therefore provides a convenient and fast way to assess possible conformational strain in the tertiary structure associated with particular target residues. However, a close inspection of possible side chain interactions is required, and the analysis should be extended to identify potential compensating features of the residues to be replaced. 3.2.6 Local Secondary Structure Propensities Effects on the overall stability of a protein can also be influenced by the respective secondary structure propensities of amino acid residues for the α-helical and βsheet conformations. However, these effects are usually marginal. Nevertheless, even at the expense of other favorable trends, such as avoiding the exposure of hydrophobic side chains to the solvent, a given residue is often favored at a certain position due to its secondary structure propensity [73]. If a particular secondary structure element does not form efficiently, many interactions between this element and other parts of the protein can be lost. The energetic consequences and the influences on folding efficiency are, however, hard to predict at this point. 3.2.7 Exposed Hydrophobic Side Chains The removal of exposed hydrophobic side chains increases the polar surface of a protein. Such mutations are not likely to affect thermodynamic stability but presumably do affect the folding efficiency by influencing the rate of aggregation of intermediates. Interestingly, they also do not affect the solubility per se (the amount of native protein that can be dissolved in buffer) and seem to act mostly on folding intermediates [74]. Because of lateral interactions, e.g., between neighboring loops, partially exposed hydrophobic amino acids can even increase stability. It must be kept in mind however, that not only the aggregation pathway, but also the stabilities

19

20 Engineering Proteins for Stability and Efficient Folding

of folding intermediates are an important parameter for kinetic stability, which can possibly be influenced by the existence of exposed hydrophobic side chains. Moreover, hydrophobic cavities at the protein surface are often involved in specific binding functions of the protein, and the removal of these “functional hot spots” has to be avoided. Therefore, the hydrophobicity of the protein surface must be carefully balanced. 3.2.8 Inter-domain Interactions In proteins consisting of more than one domain, additional principles come into play. The overall stability not only reflects the intrinsic stabilities of the single domains but is also influenced by the stability of the interface of the domains in cases where they are interacting with each other. Stabilization of this interface can be achieved mainly by increasing and optimizing the hydrophobic surface area of the interface. Because hydrophobic side chains at the interface are usually exposed during folding and transient opening of the domain interface, a tradeoff between interface stability and folding efficiency is often observed. Additional dramatic effects on stability can be observed in cases where the two domains exhibit very different intrinsic stabilities [75]. In such a scenario, the unfolding of the protein and the loss of function are strongly related to the unfolding of the less stable domain [76]. An important aspect of kinetic stabilization can be observed in twodomain proteins, where one domain can slow down the unfolding, and therefore the aggregation, of the other. Thus, the native state becomes kinetically stabilized in the assembly and a stable domain interface reduces the extent of its transient openings and, thereby, the resulting exposure of hydrophobic patches that would favor aggregation. Alternatively, covalent cross-links (e.g., disulfide bonds) between the interfaces of multimeric proteins can be introduced. For example, the introduction of disulfide bonds between the interfaces of Lactobacillus thymidylate synthase not only increases their thermal stability but also leads to reversible thermal unfolding [77]. 3.3 Case Study: Combining Consensus Design and Rational Engineering to Yield Antibodies with Favorable Biophysical Properties

The following example illustrates how consensus approaches, rational design, and experimental data can be combined in a synergistic fashion to iteratively optimize biophysical properties. The smallest form of an antibody able to bind the antigen in the same manner as the whole IgG consists of two domains, the variable domain of the heavy chain (VH ) and the variable domain of the light chain (VL ). Both domains interact with each other via an interfacial region of highly hydrophobic character. In single-chain Fv (scFv) antibody fragments, the two domains are covalently linked by a flexible linker region of typically 15–20 amino acids (Figure 3a). The binding site for the antigen usually involves three loop regions in each of the domains, named complementarity-determining regions (CDRs). Antibodies that are based

3 The Engineering Approach

Fig. 3 (a) Structure of an antibody Fv fragment consisting of the variable heavy chain (VH ) and the variable light chain (VL ). Each domain VH and VL is characterized by three hydrophobic core regions (upper [green],

central [yellow], and lower [orange] core) and a charge cluster at the base of each domain (red). Even though the residues defining these regions are conserved within the same germ-line family, these sequence

21

22 Engineering Proteins for Stability and Efficient Folding

on human antibody sequences possess great potential for many medical applications, either directly as an antibody fragment or after reconstructing an IgG [78]. Antibody fragments can be expressed in a convenient manner in E. coli, thereby providing rapid access to these proteins [79]. Nevertheless, there are often drastic differences between individual antibodies concerning their expression yield and their stabilities. Ideally, the recombinant antibodies would all provide favorable biophysical properties. The starting point for such recombinant antibodies is a library. One fully synthetic library of this kind, the human combinatorial antibody library (HuCAL), was designed based on the consensus concept [22]. Based on human antibody germline sequences, several sequence families were created. Importantly, instead of averaging over all possible human antibody sequences, the consensus sequences were built for each family separately, resulting in seven consensus frameworks for VH (VH 1a, VH 1b, VH 2, VH 3, VH 4, VH 5, and VH 6) and seven frameworks for VL (Vκ 1, Vκ 2, Vκ 3, Vκ 4, Vλ 1, Vλ 2, and Vλ 3). This diversity is important, as the use of different frameworks allows a variety of non-CDR contacts to the target, thereby greatly increasing the range of targets being recognized. By using this strategy, each human VH and VL subfamily that is frequently used during an immune response is represented by one consensus framework, and thus the immune response is closely mimicked. The consensus building was further restricted to the framework regions, while the CDRs were diversified in a manner guided by structure. Thereby, functional diversity is maintained in optimized sets of frameworks. Instead of fully relying on the statistical output of the consensus building, structural modeling was employed in order to decrease the risk of disrupting interactions between certain amino acid residues that might be in contact with each other in the three-dimensional structure. Because many structures of antibody domains are available, the modeled structures of each framework family could be compared with the respective natural template structures. The models were checked according to several principles, which have been outlined in the preceding section. At this point ←-----------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------Fig. 3 (Continued) motifs differ between different germ-line families. (b) Arrangement of the residues defining the charge cluster of VH 3 and Vk 3 domains [13]. Importantly, the charge cluster consists of a network of buried charges and hydrogen bonds rather than pairwise interactions between individual residues. (c) Furthermore, subtype-dependent packing differences occur. As an example, the residues defining the upper hydrophobic core region of different human VH subtypes are shown as structural superpositions. Structural alignments are shown of VH 4 (PDB entry 1DHV), VH 1a (1DHA), and VH 5 (1DHW) [22], with the most stable framework, VH 3

(1DHU) (left panel). While the upper core residues of VH 3 are densely packed, cavities occur for the other subtypes. In the least stable subtypes, VH 4 and VH 6, the bulky aromatic residues Phe29 and Phe31 are replaced by smaller residues, and the created space is only partly filled up by compensating residues Trp41 and Val25. The loss of the phenyl ring by replacement with Gly in VH 1a (middle panel) as well as the substitution of Leu89 by Ala are not compensated for by larger side chains at other positions, thereby creating hydrophobic cavities. The same is true for position 89 in VH 5 (right panel). Adapted from Ewert et al. [13].

3 The Engineering Approach

the models were mostly checked for whether the interactions in natural antibody domains were correctly recreated. The consensus sequence models were inspected for any unfavorable strain in the structures, represented by unfavorable regions of the Ramachandran plot or any obvious cavities in the hydrophobic core. Moreover, the sequences were checked for the existence of residues already known to be involved in conserved intra-domain interaction patterns – such as conserved charge clusters and hydrogen-bonding patterns – as well as for exposed hydrophobic side chains known to decrease expression levels. Nevertheless, differences between the natural subclasses became apparent in the models. Empirical results had suggested that certain natural framework types display more favorable stabilities and expression levels than others. These differences are already existing in the original natural human germ-line sequences. The intrinsic differences in terms of biophysical properties for each subtype were then experimentally explored in a systematic fashion, first on single domains, then on scFv fragments [13]. The experiments confirmed the observed trends, showing that VH 3 displays the highest thermodynamic stability and soluble expression level, when expressed as an individual domain, among all VH subtypes. In contrast, VH 2, VH 4, and VH 6 display the least favorable properties in terms of stability, folding yield, and the tendency to aggregate. For the VL domains, members of the Vκ subtypes showed slightly higher stabilities and expression yields than the Vλ subtypes, but the behavior was much more homogenous overall. In order to trace back these differences to the structural level, the model structures of the different subtypes were compared with each other. Several structural features can be invoked to explain the extraordinary stability of VH 3 domains in comparison with the even-numbered VH subtypes. First, differences in the hydrogen-bonding networks have an influence on the thermodynamic stabilities. Long-range interactions involving several residues are concentrated in a charge cluster at the base of VH domains to establish a complex interaction network. In VH 3 domains the ionic and hydrogen-bonding interactions within this charge cluster are very well satisfied, whereas fewer interactions are observed at the analogous positions in other subtypes (Figure 3b). Corresponding to the subclass, different hydrogen-bonding networks are formed in the charge cluster of VH domains. Some of these networks are less extended and contain a smaller number of interactions than in VH 3. Additionally, based on the residues at three different positions in the first β-strand of the VH domains, the domains can be classified into four different structural subtypes with respect to their conformations in the first framework region [80]. Mutations bringing together incompatible residues and thus “mixing” subtypes have previously been shown to have a large unfavorable effect on the stability of the whole scFv [81]. Also, clear differences can be observed for hydrophobic core packing of the family subtypes. The upper core region of VH 3 is densely packed, whereas cavities can be identified in VH 4, VH 1a, and VH 5 on the basis of structural alignments (Figure 3c). In the lower core, two of the stable domains have an aromatic residue, while the others do not.

23

24 Engineering Proteins for Stability and Efficient Folding

Finally, a comparison of the Ramachandran plots showed additional non-glycine residues with positive φ-angles and residues with higher secondary structure propensity at certain positions for the even-numbered subtypes compared with the odd-numbered ones. The immediate question was, therefore, whether the results of this structural trouble-shooting could be used to project favorable properties of VH 3 domains onto the less stable subtypes and thereby add another step to the optimization of antibody sequences while maintaining the structural diversity of the immune system. Instead of using the stable VH 3 framework exclusively, resulting in a loss of diversity, some point mutations might be enough to correct some of the shortcomings of the less stable domains. Based on these comparisons, the mutation of six residues in a scFv containing a VH 6 framework led to an overall increase in thermodynamic stability of 20 kJ mol−1 and a fourfold increase in soluble expression yield [73], indeed bringing this framework to the level of VH 3. The effects of the single mutations on stability were almost fully additive, while the effects of folding efficiency (soluble expression yield) were only qualitatively additive. The most dramatic effects on thermodynamic stability were obtained by mutations removing an unsatisfied H-bond donor in the hydrophobic core and introducing glycines at positions with positive -angles. Individually, all mutations, except the one in the hydrophobic core, led to slight increases in soluble expression yield. Interestingly, one mutation that removed a hydrophilic, solvent-exposed glutamine residue by a replacement with hydrophobic valine on the basis of higher secondary structure propensity also significantly increased the expression yield, possibly since valine secures this stretch to be in β-sheet structure. In antibodies, disulfide bond engineering has also been investigated. Optimized disulfide bonds engineered between VH and VL indeed significantly increased the half-life of an Fv fragment at 37 ◦ C [82]. This strategy was subsequently extended to interchain disulfides in generic framework positions [83]. Even more stable proteins can be obtained by combining the single-chain Fv approach with the engineered disulfide [84, 85]. It should be pointed out, however, that the additional inter-domain disulfide significantly reduces the yield of folded protein when produced in the bacterial periplasm, and such proteins have to be prepared by in vitro refolding. Therefore, the additional disulfides are not ideal for antibody libraries; instead, optimized frameworks have shown the greatest promise for combining diversity with stability. In summary, the consensus concept provides a convenient tool for proceeding with large steps in sequence space and a high probability of accumulating features that are favorable for higher stability. In many cases, structural analysis can serve as a trouble-shooting tool to identify shortcomings that might have been created by the consensus-building process or, as in the case of the antibodies described here, that are inherent to the natural sequence. In addition, it serves to rank the potential mutations identified by the consensus approach, keeping the mutational load on the target molecule to a minimum. Experimental data are important not only to validate these results but also to give important hints for future design approaches.

4 The Selection and Evolution Approach 25

Many entries in the table of experimental antibody stability data linked to mutations have come from directed evolution experiments. By combining semi-rational and rational approaches with experimental data, optimization of biophysical properties can be achieved in an iterative fashion. This interplay of experiment and structural analysis can therefore be an effective way to probe the vast sequence space in a systematic manner in order to find the valleys of free energy.

4 The Selection and Evolution Approach 4.1 Principles

The previous section may have led to the impression that mutations enhancing the biophysical properties of proteins can rapidly be identified. In the highlighted case of antibody domains, only the wealth of structural data and empirical measurements available allowed predictions with a high probability of success. Some of the empirical data used successfully for structure-based engineering have come, paradoxically, from directed evolution experiments (see Section 3.3). Despite the rapidly increasing amount of structural data and the better understanding of folding mechanisms of proteins, the effects of introduced mutations still cannot be predicted with a high degree of accuracy in most cases. The main reason is that even slight alterations in the primary sequence can lead to profound conformational changes in tertiary or quaternary structure; consequently, structural predictions have to be very accurate and usually must be backed up by empirical data. Because rational engineering uses site-directed mutagenesis followed by biophysical investigation to probe the effects of specific amino acid substitutions, it becomes very labor-intensive if many mutants have to be checked individually and if no additional hints are available. Additionally, as soon as small synergistic effects of several mutations need to be checked, the combinatorial explosion rapidly exceeds the sample number that can be handled efficiently. More importantly, the restriction to certain target residues automatically excludes alternative solutions to a given problem that may not have been obvious by the initial analysis. For example, affinity improvement of a protein to its binding partner is often achieved more efficiently by slight spatial adjustments of residues directly involved in binding, rather than by substitution of these residues. This kind of spatial adjustment can be caused by mutation of residues whose location in the native structure is further away from the actual binding site (so-called “second-sphere mutations”) [86]. Today, it is almost impossible to predict this kind of mutation by rational means. When it comes to stability engineering of proteins, the problems associated with rational design procedures are even intensified. First, because the energetic contributions of single-site mutations are usually small, the need to sample multiple mutants – in which synergistic effects of several mutations are combined – is more acute. Second, the factors and principles responsible for the overall stability of a

26 Engineering Proteins for Stability and Efficient Folding

native protein are still far from being completely understood, and a multitude of different forces and interactions contribute to it. A complete analysis will have to consider not only the interaction network of the native protein but also the effects on the denatured state. Additionally, potential aggregation pathways will have to be considered. Even simple substitutions like the ones discussed in the previous sections often make contributions to the entropy as well as the enthalpy of the folded and unfolded states, including the solvent in either state. Third, because rational approaches always rely on the current theoretical knowledge, new principles underlying protein stability will rarely be uncovered. In any case, rational engineering requires a clear definition of the problem by the investigator in order to find a solution. Ironically, in the field of protein engineering, the exact definition of the problem is often the problem itself. Thus, to overcome these limitations, an experimental setup is needed that allows the creation of a vast number of variants of a given protein and that subsequently can identify “superior” molecules that best fulfill a desired property. Nature samples the vast sequence space by the strategy of Darwinian evolution, a cyclic iteration of randomization and selection. Nature thereby adapts proteins to fulfill a function under the given environmental conditions. Recent developments in molecular biology have made it possible to mimic Darwinian evolution in a reasonable time in vitro. Not only has this “evolutionary approach” become the most powerful method to date for engineering proteins towards a desired property, but it also provides new insights into the mechanisms and principles that are responsible for this property. However, as has been illustrated in the case study on antibodies, such experiments can be used not only to solve a particular problem but also to gather information about which residues tend to become enriched in particular positions. This again provides a database for rational engineering. Many different selection and evolution strategies have been developed in recent years, but all of them have several features in common that reflect the principles of Darwinian evolution. The starting point is the generation of a genetic library of mutants derived from the wild-type sequence of the protein under examination. Several methods exist to create sequence diversity. In error-prone PCR, the error rate of polymerases is increased by performing the PCR reaction in the presence of deoxynucleotide analogues or in the presence of other metal ions. By using bacterial mutator strains, which are characterized by deficient DNA repair systems, random mutations are introduced during DNA amplification in the bacterial cell, albeit usually at a lower frequency [87]. In DNA shuffling, which mimics the natural process of sexual recombination, genes are randomly fragmented by nuclease digestion and reassembled by a PCR reaction in which homologous fragments act as primers for each other [88] (Figure 4). The staggered extension process [89] is another possibility to obtain recombined genes in vitro. Here, the polymerase-catalyzed extension of template sequences is extremely abbreviated, and repeated cycles of denaturation and extension lead to several template switches – thereby recombining elements from different genes – before the extension finally yields full-length products. Additionally, techniques are available to focus the diversity to certain regions on the whole gene, such as the use of degenerate primers in PCR or the “doping” of a

4 The Selection and Evolution Approach 27

Fig. 4 Methods to create genetic diversity by biochemical means. On the left, errorprone PCR (see text) is depicted schematically. Two types of mutations are shown: favorable ones (open squares) and unfavorable ones (closed circles). In successive cycles of PCR, more mutations of each are introduced, and usually molecules will contain some of either type. Thus, the beneficial effect of the favorable mutations can be completely obscured by the presence of unfavorable ones, if the error rate is too high.

Therefore, this method is most successful if it is not used at too high an error rate. On the right, DNA shuffling according to Stemmer [88] is shown. A short DNAse digestion breaks up the DNA into small pieces, and PCR is used to reassemble the gene. Thereby, mutations are “crossed” and genes with largely favorable mutations can be obtained that can be enriched by selection. Nevertheless, successful evolution experiments have been carried out with either method.

shuffling reaction with degenerate primers. Depending on the problem and the sequence under investigation, each of these methods has its advantages and disadvantages. In any case, the generated library should obey two major criteria: it must be diverse enough to contain individual sequences with beneficial mutations, and it should be of high enough quality to reduce the experimental “noise” (such as sequences with stop codons or frameshifts) in the subsequent selection experiment [90]. A primary prerequisite of any selection system is the coupling of the genotype (the gene sequence) and phenotype (the respective protein displaying the properties) of any individual library member. Briefly, this can be done by two strategies. Either the gene and the protein need to be compartmentalized in cells or artificial compartments, such that the “improved” phenotype stays connected to the altered

28 Engineering Proteins for Stability and Efficient Folding

Fig. 5 Two principal strategies to link phenotype and genotype are depicted. The genetic material must be connected in a unique way to the protein, which defines the phenotype, such that the gene encoding the valuable mutation can be selectively amplified. A collection of two mutant proteins and mutant genes is shown (light gray and dark gray). (a) This connection can be realized through a direct link (e.g., in phage display or ribosome display), as shown on the top. In this case, all assemblies can freely diffuse in the same volume and the selection must filter out those proteins with the desired function, e.g., by binding to a ligand. (b) Alternatively, gene and protein

must be in the same compartment. In nature, this is realized in cells, and microbial cells can be manipulated so that each takes up one variant of a mutant collection. The key is to identify the improved phenotype. This can be done by screening (individual assays on cells) or selection (giving cells with the desired property of the protein a growth advantage). Rather than natural cells, water-in-oil emulsions can be used to create artificial compartments of small water droplets in an oil phase. Usually, the emulsion must be broken and the proteins must be selected by binding to identify the one with the desired phenotype. For details, see text.

gene, or the protein has to be physically coupled to the gene, such that they can be isolated as a particle containing both gene and protein (Figure 5). By selecting the proteins displaying the desired properties, the linked gene sequence can be inherited and amplified subsequently. The various selection technologies differ mainly in the way this physical linkage or compartmentalization is achieved. This will be discussed in more detail in the following section. The library must then be screened or subjected to selection for a certain function, and a defined selection pressure can be applied to direct the selection towards a molecular quantity of interest. It is necessary at this point to discriminate between “screening” and “selection.” Screening methods examine individual members of a library for a given property (e.g., catalytic activity or solubility). For certain properties, screening is often the only way to go. The number of mutants to be screened in a reasonable time depends on the versatility of the screening method. Despite constant progress in automation and miniaturization, even the best screens to date usually do not allow assessment of more than 106 variants in a reasonable time. In contrast, selection methods force the single library members to compete with each other, and members that best fulfill the specified criteria are enriched. Often, the selection

4 The Selection and Evolution Approach 29

is based on the binding of particular variants to an immobilized ligand. Note that the binding is only a “surrogate quantity” of the real property to be improved. The basis for a successful enrichment is an efficient counter-selection of variants that do not possess properties fulfilling these criteria. Especially during the selection of proteins for higher stability, an efficient counter-selection, in addition to the correct choice of the applied selection pressure, is one of the major experimental challenges [91]. The basic rule of screening and selection technology describes the importance of assigning the correct selection pressure. This has been succinctly phrased: “You only get what you screen for!” [92]. An analogous statement can be made for selection. Even though the rule sounds plausible, it is often difficult to translate this statement into an experimental setup, because an additional complication is introduced by the fact that an explicit selection pressure towards just one property is impossible to realize. Depending on the selection procedure used, more than one molecular property will have an influence on the enrichment process, including, for example, affinity or catalytic activity, thermodynamic stability, folding efficiency, and toxicity of the respective molecule. The outcome of the selection experiment will therefore always be a “compromise solution” with respect to the weighting of many properties in a given experimental setup. Assigning the right selection pressure therefore means biasing the selection towards a certain property, rather than exclusively altering this property. For these reasons we will discuss the application of the various selection technologies with an emphasis on the available selection pressures for stability engineering. Some of the described methods and examples will be based on mere screening of library members, but the main focus will be on the selection for favorable protein variants. Choosing the appropriate selection pressure is just one side of the coin. The proper adjustment of its strength is another important factor. If the selection pressure on the system is too low, molecules with the desired properties will be lost in the background noise of the experiment – i.e., they are not enriched. On the other hand, if the selection pressure is too stringent, even the best variants might not pass the “survival threshold.” The genes of variants that survive the applied selection pressures are subsequently amplified and subjected to another so-called “round of selection.” This either can be performed in the absence of further mutagenesis to simply enrich the best members of the initial library (which is constant) or, to completely mimic the Darwinian principle, the selected members can be subjected to alternating rounds of randomization or recombination before subsequent selection, leading to an adaptation of the library from round to round. However, as most of the mutations will be non-beneficial, care has to be taken that the mutation rate is low enough to allow successful enrichment of improved members and not to extinguish the whole population. We use the terms “combinatorial selection” for a process in the absence of such mutations and “evolutionary selection” for a process that includes such mutations. It has been pointed out that the method of DNA family shuffling is in some respects similar to the consensus approach because it combines gene fragments

30 Engineering Proteins for Stability and Efficient Folding

Fig. 6 Common display systems used for selection for enhanced biophysical properties. In in vitro display technologies (a), the proteins are produced by in vitro translation, and protein production does not rely on a host organism. The displayed proteins are linked to RNA either by stabilized ternary complexes (ribosome display, upper panel)

or by a covalent puromycin cross-linker (mRNA display, lower panel), thereby establishing the genotype-phenotype linkage. (b) In contrast, partial in vitro display systems rely on cells to produce the displayed protein, but selections can be performed in vitro. In phage display (upper panel), the protein is displayed on the

4 The Selection and Evolution Approach 31

from homologous sources in a random fashion. For mere statistical reasons, the probability of replacing a residue with a consensus residue by gene shuffling is also higher than replacing it with a non-consensus residue [93]. There is, however, a crucial difference: when recombining genes in a random fashion, a selection or screening step is needed subsequently to identify the “fittest” members of the resulting collection. Rather than being based on theoretical assumptions, gene-shuffling methods mimic the natural process of evolution by identifying the members exhibiting a desired property by explicitly subjecting them to selective pressure for this desired property. In practice, however, even with the largest of libraries, a full “re-equilibration” of residues will not take place. In contrast, the consensus approach is based on the explicit assumption that the statistical preferences in a given (often limited or biased) set of sequences indeed reflect the energetic preferences. However, there may be other reasons that certain sequences are prominent.

4.2 Screening and Selection Technologies Available for Improving Biophysical Properties

As described above, any selection technology relies on four major steps: (1) generation of a genetic library, (2) establishment of the link between genotype and phenotype upon translation into protein, (3) subsequent screening or selection under defined conditions, and (4) re-amplification of selected members. Depending on how these steps are performed, selection technologies can be subdivided into in vitro methods, partial in vitro methods, and in vivo methods. While in vivo systems rely on a host organism to express the protein and to carry the respective genetic information, all in vitro display technologies have in common that the protein production and the selection process are performed entirely in vitro, i.e., that the protein is obtained by cell-free translation. In the partial in vitro methods, the genetic information is introduced in cells, where protein production occurs, while the selection process is performed in vitro. All techniques differ in the way the physical linkage between the protein and its genetic information is established. The principles of linking genotype and phenotype are shown in Figure 6.

←-----------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------Fig. 6 (Continued) surface of filamentous phages, usually fused to the CT domain of the minor coat protein g3p. The three domains (N1, N2, and CT) of the minor coat protein g3p are depicted. The phage particle also carries the gene encoding the displayed protein. In other partial in vitro technologies, the proteins are displayed on the surface of the expressing host cell itself, which are either bacterial cells (bacterial surface display) or yeast cells (yeast surface display)

(lower panel) that also harbor the respective plasmid DNA. (c) in vivo screening or selection systems use the properties of fused reporter proteins for screening (upper panel) or split proteins for intracellular selection (lower panel) in which cellular growth and therefore amplification of the genetic material occurs only if the two protein halves are reconstituted upon interaction of the fused library protein with its target.

32 Engineering Proteins for Stability and Efficient Folding

4.2.1 In Vitro Display Technologies in vitro display technologies started off with the selection of peptides [94] and were then made efficient enough to select for functional proteins [95]. A number of technologies have been developed, which will be described briefly: ribosome display [95, 96], RNA-peptide fusions [97, 98], and water-in-oil emulsions [99, 100]. Water-in-oil emulsions constitute artificial compartments, rather than a physical link, and thereby create a phenotype-genotype coupling and can also use a selection step for interaction with a target. In ribosome display, the genetic library, usually in the form of a PCR product, is directly used to produce mRNA by in vitro transcription. This mRNA contains a ribosome-binding site for the subsequent translation of the protein and several features that stabilize it against degradation. The encoded protein variants are expressed in a cell-free translation reaction by a stoichiometric number of ribosomes. The essential linkage of the translated proteins to their respective mRNA molecule is achieved by eliminating the stop codon and stabilizing the ribosomal complexes, which prevents the release of the translated protein from the ribosome. In the related method of mRNA-peptide fusions, additional steps are used after this stoichiometric translation to covalently couple a linker between the end of the mRNA and the protein. The ribosome is then removed and the complexes are purified. These complexes are then used to bind to an immobilized target. In all these methods, after the selection process the re-amplification of genetic material can be performed solely by biochemical means, using reverse transcriptase to first synthesize RNA-DNA hybrids, which are then subsequently amplified in a PCR reaction. The resulting DNA then serves as a template for the production of mRNA used for the next round of selection (Figure 7). In vitro display technologies offer several advantages over in vivo selection systems. First, much larger library sizes are accessible because the creation of large molecular diversity on the genetic level does not pose a great challenge, and diversities up to 1013 are easily achieved. For in vivo systems, the critical step that limits the size of the displayed library is the transformation efficiency of the respective host organism. Only a small fraction of the initially generated pool will actually enter the cells, and, depending on the host organism, library sizes of about 107 variants in yeast or up to 1011 variants in bacteria can be achieved. In contrast, for in vitro selection systems such a transformation step is not necessary and the diversity of the library is defined by the number of different RNA molecules added to the cell-free translation reaction or by the number of functional ribosomes, whichever number is smaller. For a library of 1014 , a 10-mL translation reaction is required. A second advantage is that, because translation and protein folding take place in vitro, reagents can be added during protein synthesis, which can either promote protein folding or minimize aggregation of the displayed proteins (e.g., chaperones), or substances that exert certain selection pressures can be added. in vitro translation encompasses an additional advantage: many proteins are not compatible with in vivo selection systems because the wild type or at least some of the mutants are toxic to the host organism, undergo severe degradation, or cannot be expressed functionally in the respective cellular environment. Last but not least, a major advantage

Fig. 7 Selection cycle of ribosome display (left panel) and phage display (right panel). For ribosome display, all necessary steps are performed in vitro, whereas for phage display protein translation, folding, and phage assembly take place in the bacterial host cell. Other selection criteria can be applied to the protein of interest (see Figure 8) before selecting for binding to a target, thereby introducing selection pressures for higher stability. It has to be guaranteed, however, that the selection conditions are not too harsh to destroy the linkage of genotype and phe-

notype. Importantly, the genetic pool of library members in phage display remains constant in the course of several selection rounds, whereas in ribosome display, the genetic pool undergoes continuous modifications during the in vitro amplification steps by the limited accuracy of the polymerases, resulting in a true Darwinian evolution. In addition, this pool can deliberately and conveniently be altered by introducing additional mutations or recombination events such as DNA shuffling.

4 The Selection and Evolution Approach 33

34 Engineering Proteins for Stability and Efficient Folding

results from the amplification process after the selection has been performed. Because all enzymes used to convert and amplify the genetic material possess an intrinsic error rate, the genetic pool is never constant and is continuously modified from round to round. Therefore, even without special measures to increase the error rate, usually some “evolution” is observed. The error rates can be additionally increased by performing error-prone PCR or by recombining favorable mutations by means of DNA shuffling as explained in the previous section. 4.2.2 Partial in vitro Display Technologies We define partial in vitro display technologies as those methods in which host organisms are employed to carry and amplify the genetic information and to produce the proteins from this genetic library, usually encoded on plasmids. The selection step, however, is performed in vitro. Therefore, selection pressures similar to those in the in vitro methods can be applied. The array of available techniques includes phage display [101], bacterial surface display [102], and surface display on yeast [103]. Phage display is still the most popular selection technique, and due to its robustness, most of the studies that aim for enhanced biophysical properties of proteins have utilized this technique. Originally, phage display was applied as a method for the identification and selection of peptides or proteins binding to a specific target [101]. More recently, several developments paved the way for its application as a tool for studying protein folding, and as a result, it is now used for the selection of proteins with improved biophysical properties. We will briefly review the main principles of phage display in its most common format. Variants of the protein of interest are fused to the minor coat protein g3p of the filamentous bacteriophage, which consists of three domains (namely, N1, N2, and CT) that are connected by glycine-rich linkers. The fusion protein is usually encoded on a phagemid vector, expressed in the E. coli host and assembled. Typically, the other proteins necessary to produce an intact phage particle are provided by the helper phage. The assembled phages are secreted by the cell, which does not lyse, and the phages can be collected. While the protein is displayed on the exterior of the phage (typically as an N-terminal fusion to either N1 or CT), the gene of interest encoding a library member is packed into the phage particle upon its assembly in the bacterial host. The phages can then be subjected to the selection process. Phages compete with each other for binding to an immobilized target, using the displayed library members for interaction, and can thereby be captured. Optionally, before capturing, the phages can be subjected to harsh environmental conditions. The captured phages are subsequently amplified by re-infecting bacteria, which then produce phages for a new round of selection (Figure 7). In the ideal case, only phages that specifically recognize the target would be amplified. Due to reasons such as nonspecific “sticking” of phages to the surface, the efficiency of selection is in practice at most a 1000- to 10 000-fold enrichment over nonspecific molecules and can even be very much lower. Several aspects are important when phage display is used as a means of evolving proteins with improved biophysical properties. The correct folding of the fusion

4 The Selection and Evolution Approach 35

protein during phage morphogenesis is an important parameter determining the frequency of incorporation of correctly folded proteins displayed on the phage. Folding intermediates that have a strong aggregation tendency lead to aggregation of a particular library member during the assembly process, and rather than the g3p fusion, the g3p wild-type protein from the helper phage will be incorporated. Despite this inherent selection pressure for the folding efficiency of displayed proteins, the enrichment of improved variants is slow [104], and the utility of phage display to study and improve folding of a given protein was initially not obvious [105]. Combined with a selection for protein functionality, it nevertheless sets the basis for selecting proteins according to their folding properties, as will be discussed in Section 4.3.2. Since the selection procedure is performed in vitro, a large variety of external selection pressures can be applied to the phage particles and be combined with functional selection. If function is omitted as a direct indicator for the native state, other criteria that directly correlate with the native state have to be translated into a selectable feature. This opens the door to select for proteins, which do not bind a ligand, albeit with the caveat that it is not assured that the native state is being selected. Such general selection approaches for physical properties take advantage of the fact that unfolded proteins are more susceptible to proteolytic cleavage than are compactly folded ones [106]. One variation on this theme makes use of the modular nature of the g3p protein, thereby linking phage infectivity directly to the proteolytic susceptibility of the target protein. This system has been called “Proside” (protein stability increase by directed evolution) [107] and will be discussed in Section 4.4.4. 4.2.3 In Vivo Selection Technologies In vivo selection technologies such as the yeast two-hybrid system [108, 109] or other split protein complementation assays [110, 111] are valuable tools for studying protein-protein interactions in living cells. They commonly employ reporter systems, in which a covalent link between the protein of interest and another so-called reporter protein is established. Cell growth or a colorimetric reaction is dependent on either a specific protein-protein interaction or the solubility of a critical component. The reporter protein transduces certain properties of the host protein, most importantly its native fold and its ability to interact and/or its solubility or its resistance to cellular proteases, to a screenable or selectable feature of the fused reporter protein itself [112]. Examples of reporter proteins are transcription factors [108], critical metabolic enzymes [111], or proteins that can easily be assayed [113]. Several facts, however, limit the applicability of in vivo systems for stability engineering. First, as the viability of the host organism has to be guaranteed, external selection pressures cannot easily be applied. Thermophilic organisms represent in some cases an interesting alternative, but conditions allowing selections are hard to establish. Additionally, the host environment as well as the fused reporter proteins possibly perturb the characteristics of the target protein. Furthermore, control experiments must ensure that growth is really dependent on the interaction

36 Engineering Proteins for Stability and Efficient Folding

of interest and that mutations in the host have not “short-circuited” the selection strategy. 4.3 Selection for Enhanced Biophysical Properties

As has been explained in the Introduction, desirable biophysical parameters include solubility, stability, and folding efficiency. Even though the various selection methods and conditions usually do not improve one of these properties exclusively, they are often biased towards one of them. We will therefore discuss different ways to exert selection pressure on the system with respect to the property that is likely to be changed. 4.3.1 Selection for Solubility Solubility, correctly defined as the maximal concentration of the native protein that can be kept in solution, is usually not a property to be selected, as most proteins – with the exception of membrane proteins – are sufficiently soluble. The word “solubility” in the context of screening is often inaccurately used to refer to “soluble expression yield,” and thus it usually mirrors the efficiency of folding in the cell. Even though soluble expression yield is not necessarily equivalent to the folding properties and even less to the stability of a protein, a correlation can often be observed, and soluble expression yield is comparatively easy to screen for. In some cases, a function for a given protein cannot be assigned, or it is difficult and laborious to screen for, and then this property becomes especially important. The fluorescence yield of bacterial colonies expressing proteins or protein domains fused to the green fluorescent protein (GFP) correlates over a wide range with the soluble expression yield of the fused domain [114]. Screening for fluorescence intensity is a versatile method, and, combined with directed evolution, has the potential to select variants of proteins that are less aggregation-prone than their progenitors [115, 116]. In a similar setup, reporter proteins can be used as selection markers instead, such as by fusing the protein of interest to chloramphenicol acetyltransferase [117]. Assays exploiting protein-protein interactions can also be used for this purpose and they have the additional advantage that the fused reporter sequence can be much smaller and thereby minimize the risk of a perturbing influence of the reporter itself on the domain of interest [113]. To completely exclude such perturbing influences, reporter systems might possibly be established that do not need any fusion at all and exploit instead the stress response of the expression host cell. As the overexpression of proteins often activates particular stress response genes by the accumulation of insoluble aggregates, the respective gene promoters can be employed to activate reporter genes instead [118]. In special cases, reporter systems can also be combined with other selection methods to select for folding properties. Many intrinsically stable proteins contain permissive sites in loops, into which polypeptide chains of variable length can be inserted without loss of function of the host protein. The higher the number of residues inserted, the larger the entropic cost of ordering these residues will be.

4 The Selection and Evolution Approach 37

Consequently, the overall stability of the host protein should decrease. However, the entropic cost of inserting a folded sequence will be lower than that of an unfolded sequence of the same length. The probability of the host protein reaching the native (functional) state should thus correlate directly with the ability of the inserted sequence to fold into a compact structure. If host proteins with various inserted sequences are displayed on phages and if the host protein allows a selection for “foldedness” by means of binding to a target, folded sequence insertions are enriched. This system has been termed “loop entropy reduction phage display selection” [119]. While attractive in theory, it was found experimentally that mostly sequences that keep the hybrid protein soluble are enriched. Protein solubility, as used here, can also be interpreted as a lower degree of exposed hydrophobic residues, and folded proteins usually display fewer hydrophobic residues on their surface than do unfolded proteins. Conversely, the exposure of hydrophobic residues is usually a sign of non-native states. Display technologies can thus, for example, use the interaction with hydrophobic surfaces to select against more hydrophobic, i.e., less “folded,” proteins [120]. In summary, protein solubility can reflect in many cases the folding properties of proteins and thus be used as a selection criterion. Moreover, soluble expression in vivo is often a major requirement for the large-scale production and the convenient in vitro handling of proteins. In any case, solubility should never be confused with protein stability. Even though both properties might correlate in some cases, the governing principles are often of a different nature. 4.3.2 Selection for Protein Display Rates The in vivo folding efficiency of proteins represents an intrinsic selection pressure when using phage display. Because correct folding is a prerequisite for both incorporation into the phage coat and binding to the target, the subsequent selection of phages that are able to bind to the target is partly influenced by the folding properties of the displayed molecule. However, this does not automatically imply that the selection is driven towards superior folding properties. As mentioned above, the enrichment factors of proteins with superb folding behavior over the poorly folding members are low [104]. Moreover, even though the binding of a given protein to its target is a very direct way to monitor and screen for its proper folding, the selection criterion “folding” is not decoupled from the selection criterion “binding affinity.” If the enrichment factor for one property (folding) is low, the selection is more likely to be driven towards the other (affinity). As a result, selected members will represent a compromise between folding properties, which only have to be sufficient under the given experimental conditions, and the binding affinity to their target. By randomizing exclusively the residues that build the hydrophobic core of the IgG binding domain of peptostreptococcal protein L, which are not involved in ligand binding but determine the stability and folding kinetics of this small protein, Gu et al. [121] could unambiguously demonstrate the utility of phage display for studying the stability and folding efficiency of proteins. A more extensive randomization effort with subsequent characterization of the selected variants pointed out

38 Engineering Proteins for Stability and Efficient Folding

some important aspects of selection for folding [122]. First, it showed that folding kinetics is not the critical parameter for the selection but rather the overall stability of the mutants, a tendency which could be confirmed by more advanced selection approaches (see Section 4.4.2). Second, the authors demonstrated the utility of selection methods to identify certain residues that are important for the folding mechanism. Third, the selected pools were highly diverse and the thermodynamic stabilities of all variants were lower compared to the wild type. In fact, most of the selected proteins denatured just above room temperature. These results illustrate the principle that any selection – be it natural or in the test tube – simply continues until the minimum requirements are met, in this case, functionality at the selection temperature. Consequently, this suggests that additional, more stringent selection pressures are necessary to really accomplish a directed evolution for improved properties. 4.3.3 Selection on the Basis of Cellular Quality Control Additional selection pressure can be provided by the host organism itself. For example, the secretory quality-control system of eukaryotic cells discriminates proteins according to their folding behavior in an efficient way. This is based on mechanisms that lead to retention of misfolded proteins in the endoplasmic reticulum (ER), followed by degradation of these proteins. In yeast, the Golgi complex can reroute misfolded proteins, which have escaped ER retention, to the vacuole for degradation, thereby constituting an additional important quality-control pathway [14]. In combination with yeast surface display, these mechanisms can be employed to bias functional selections towards enhanced folding efficiency. Several studies have shown that the surface display rate of proteins strongly correlates with their ther-mostability and their soluble secretion efficiency [123, 124]. As an example, by making use of elevated temperatures during expression as an additional selection pressure, improved T-cell receptor fragments, whose thermostability exceeded by far the expression temperatures, could be obtained [125]. Even though the low transformation efficiency of yeast restricts the accessible library size, one obvious advantage of the method is the applicability to glycosylated eukaryotic proteins, which generally are not amenable to yeast two-hybrid or phage display methodologies [126]. 4.4 Selection for Increased Stability 4.4.1 General Strategies The key to all selections for stability is to introduce a threshold that separates the molecule with desired properties from the starting molecules. If the population initially lacks functionality, while only a few members are above the selection threshold, the selection system can distinguish between these slight energetic differences. This is the starting situation if the target protein is initially of very low stability and should be brought to “average” properties.

4 The Selection and Evolution Approach 39

If, however, the starting protein is already of considerable stability, but should be brought to even higher stability, it can be advantageous to intentionally destabilize it prior to selection in order to find stabilizing mutations that reconstitute its functionality. This principle can be applied to very different kinds of proteins, provided that mutations are known that destabilize the protein to an extent that will subsequently allow the selection for alternative stabilizing mutations (Figure 8a). Because the effects of independent stabilizing mutations are often additive, the deliberately introduced destabilizing mutations can be reverted in the context of the additional newly selected ones, thereby rendering the molecule far more stable than the original one (Figure 8b). In principle, all reagents and conditions known to destabilize proteins can decrease the number of library members populating the folded state and can therefore be used to exert increasing selection pressure on the system. Increased temperature and denaturing agents are obvious methods that are useful for selecting proteins of higher thermodynamic stability. The major problem is presented by the compatibility of the conditions with the selection method used. 4.4.2 Protein Destabilization An example of the stabilization of a naturally unstable domain by using phage display was a selection performed with the prodomain of the protease subtilisin BPN [127]. At room temperature the prodomain folds into a stable conformation only upon binding to subtilisin. By randomizing positions that are not directly in contact with subtilisin and subsequent selection on subtilisin, a mutant was selected that showed an increase of Gunfolding by 25 kJ mol−1 , from −8 kJ mol−1 to 17 kJ mol−1 , despite the fact that the library size was comparatively small. Intriguingly, the predominant energetic contribution was mediated by a selected disulfide bond. Previously, a similar strategy had been used to select for thermodynamically favored β-turns of the B1 domain of protein G [128]. Based on considerations described above and on the fact that the replacement of amino acids on the surface of proteins would be predicted to have only moderate effects on stability, the authors reasoned that such a selection could be successful only for proteins of marginal stability. Most of the substitutions would then lead to a positive free energy of folding, and thus the molecule would fail to fold into a functional form at all. In fact, selected turn sequences showed clear sequence preferences only if less stable host proteins were used to accept the turn. In this case, the selected sequences either resembled the wild-type turn or reflected the statistical preferences of turn sequences in the databases and stabilized the protein by 12–20 kJ mol−1 , compared to random sequences. Moreover, increased temperature was used as an additional selection pressure during the phage selection. As discussed in the previous section, the inability of the wild-type target protein to fold is the prerequisite for performing an efficient positive selection for functionality by additional mutations. The internal disulfide bond of immunoglobulin domains significantly contributes to the stability of antibodies [129]. The dramatic loss of free energy of folding upon their removal usually renders antibodies nonfunctional. By first destabilizing a scFv antibody fragment through replacement

Fig. 8 Principle of selection for proteins with improved stability. (a) By random mutagenesis of a given sequence, many mutations are obtained. Some of these mutations are favorable for the stability of the native protein and others are unfavorable. As a result, a diverse pool of proteins with a distribution of different energies is created. Upon exposure of the pool to an external selection pressure, only mutants with stabilities exceeding the selection threshold can be recovered and amplied. However, the selection threshold has to be set high enough to allow an efficient selection of “improved”

members. (b) Destabilization of the wild-type sequence prior to selection allows reducing the necessary selection threshold. Thereby, alternative mutations that stabilize the native fold of the protein can be identited and the selection process becomes more efficient. Moreover, additional stability gains can be achieved by removing the deliberately introduced destabilizing mutations after the selection. The initially lost energy is therefore regained, resulting in mutants of higher stability than the wild-type protein. Adapted ¨ et al. [76]. from Worn

40 Engineering Proteins for Stability and Efficient Folding

4 The Selection and Evolution Approach 41

of cysteines with other residues in both variable domains separately, a completely disulfide-free antibody was identified after several cycles of functional selection and recombination by DNA shuffling [17]. One globally stabilizing mutation was found to compensate for the initial stability loss. This study illustrates some important features of how several parameters exert influence on different stages of the selection process. Interestingly, the stabilizing mutation was already selected during the first rounds, showing that the loss of thermodynamic stability was the primary problem that had to be overcome. Because the stability gain of this mutation was large enough to shift the free energy of folding above the required threshold, all successive rounds did not affect protein stability but led to a fine-tuning with respect to the improvement of folding yield [17]. To illustrate the principle of additivity, reintroduction of the disulfide bridge into the selected variant yielded a scFv antibody of very high stability and superior expression yields [84]. 4.4.3 Selections Based on Elevated Temperature in vivo selection systems using thermophilic expression hosts have been employed for the stabilization of enzymes [130]. However, a rather large set of requirements must be met to apply such a selection system. In order to use stability of the enzyme at elevated temperatures as the selection criterion, its enzymatic activity must be vital to the thermophilic organism, and the corresponding gene of the thermophilic host has to be deleted. Randomized versions of a mesophilic enzyme can then be screened by means of metabolic selection. Although the approach is very powerful, the utility of these systems is restricted to special cases. To evolve enzymes with altered thermal stability, conventional screening for enzymatic activity in vitro after randomization or recombination of the respective gene and subsequent expression of the enzyme is still the most widely used method. An activity screen rapid and sensitive enough to identify slightly improved members from a vast pool of mutants is the key feature of this directed evolution approach [131]. However, individual screens have to be developed for each specific class of enzymes. Even though screening limits the explorable sequence space considerably compared to selection, mesophilic enzymes such as subtilisin E [132] and p-nitrobenzyl esterase [133] could be converted into mutants functionally equivalent to thermophilic enzymes by only a few rounds of directed evolution. Similar to observations discussed before, only very few mutations were necessary to increase the melting temperatures by more than 14 degrees. An important feature of screening compared to selection is that the much smaller library size is partially compensated for by directly measuring the quantity of interest: enzymatic activity at elevated temperature. In contrast, most selection systems use a surrogate measure where “false positives” can lead to the phenotype by mechanisms different from the ones desired. Elevated temperatures not only can be used in screening but also can be combined with selection technologies. While ribosome display is not an option – because low temperatures are essential to keep the ternary complexes of RNA, ribosome, and nascent polypeptide intact and thus ensure the coupling of genotype and phenotype – phage display has proved to be a very suitable method for harsh conditions

42 Engineering Proteins for Stability and Efficient Folding

due to the robustness of the phage particles. Nevertheless, future improvements of in vitro technologies may allow their application under more stringent conditions. As of today, in the case of in vitro technologies, it is vital to destabilize the protein first (see Section 4.4.2); then, very significant stability improvements can be selected [134]. The upper temperature limit for selections using filamentous M13 phages is approximately 60 ◦ C [135]. Above this temperature, re-infection titers of the phages are severely decreased, presumably due to the irreversible heat denaturation of phage coat proteins. Phages displaying the protein of interest can be incubated up to this temperature, and proteins still able to function can be selected. It should be noted that the exposure of phages to higher temperature after phage assembly exerts a somewhat different stress on the proteins than in the methods described before. While in the previous examples the functionality of the proteins was influenced mainly by in vivo folding efficiency and the protein stability during the panning procedure, it is the irreversible unfolding reaction at a given temperature that now becomes an additional parameter. The fraction of unfolded molecules will therefore reflect the rate of unfolding and thus kinetic stability. 4.4.4 Selections Based on Destabilizing Agents The use of protein destabilizing agents for biasing the selection pressure towards stability is limited to in vitro and partial in vitro display methods. Like temperature, the concentrations of denaturing agents can be controlled precisely and can be varied from round to round, allowing a gradual increase of stringency. Even though phages are quite resistant to denaturing agents [136], one should be aware of possible general problems when selections are based on the chemical denaturation properties of the displayed proteins. However, if chemical denaturation is combined with a selection for binding, high concentrations of denaturant can prevent binding to the target, even if the protein is not yet unfolded: as the forces governing ligand binding are very similar to those responsible for protein stability, both are disturbed by chemical denaturants. Conversely, because chemical denaturation is in fact often reversible, removal or dilution of the denaturant prior to ligand binding will often result in refolding on the phage and thus release of selection pressure. Moreover, ionic denaturants such as guanidinium chloride weaken electrostatic interactions and strengthen hydrophobic ones. This might impair the selection of mutations that introduce additional ionic interactions on the protein surface [59] and may favor additional hydrophobic interactions, which is not necessarily desired. If the protein is known to be stabilized by disulfide bridges, a strategy similar to the one used by Proba et al. [17] described above may be applicable. To identify globally stabilizing mutations, which compensate for the stability loss upon removal of disulfide bridges, selection can be performed in the presence of reducing agents such as DTT. Thus, the disulfide-forming cysteines do not have to be removed in advance. However, one should be aware of the fact that some disulfide bridges in proteins, once formed, are often hard to reduce, especially if they are buried within the protein core. Thus, it is advantageous to add the reducing agent at a time

4 The Selection and Evolution Approach 43

when the protein is not yet folded. Jermutus et al. [134] stabilized a scFv antibody fragment by using ribosome display. In contrast to the phage display method, the synthesis of the targeted protein occurs in vitro, which makes the polypeptide chain accessible to reagents during its synthesis on the ribosome and before folding has taken place. DTT was added during translation of the scFv, and its concentration was continuously increased from round to round. When using ribosome display, the population undergoes slight changes due to mutations occurring during PCR, resulting in an iterative adjustment of the selected variants towards tolerating the increasingly stringent conditions. 4.4.5 Selection for Proteolytic Stability In each of the examples cited above, selection for stability was based on a functional selection. Therefore, only those proteins for which a specific function can be assigned could be targeted; in addition, this function has to be screenable or, better yet, selectable. It would be highly advantageous to completely uncouple function from stability in the selection process to extend the range of problems to which this can be applied. A more general approach would thus be an invaluable tool for engineering any given protein and for selecting stable folds from a pool of de novo designed proteins. For functional selections, an additional problem arises from the fact that after a certain stability threshold is reached, allowing enough proteins to populate the folded state, the selection is likely to run towards improved binding properties instead. On the other hand, certain mutations might be stabilizing but might result in slight structural rearrangements that affect functionality in a negative way. Thus, it will not be possible to recover these mutations [106]. A general approach that has proved to be powerful for stability engineering is based on the concept that compactly folded proteins are much more resistant to proteolysis than are partly folded or unfolded proteins [137]. Several variations on this theme have been applied to phage display selections. In principle, it is only necessary that the phages displaying proteins, which can be cleaved by the protease, can be efficiently separated from the phages displaying protease-resistant proteins. This can be achieved in a physical way by providing the displayed protein with an N-terminal tag sequence allowing capture of only phages with non-cleaved proteins [106]. Alternatively, the ability of phages to re-infect bacteria can be directly linked to the protease resistance of the protein of interest [107, 136]. This second alternative takes advantage of the modular structure of the protein domains responsible for phage infection. If the displayed domains are inserted between the carboxy-terminal domain of gp3 and the amino-terminal domains N1 and N2, which are required for phage infectivity, proteolytic cleavage of the displayed domain renders the phage noninfectious. This was inspired by the so-called selectively infective phage method (SIP), where it was shown that the g3p domains could be interrupted by additional domains and even an interacting pair [138]. By employing a phage system derived from the SIP technique, which lacks any wild-type g3p, it is assured that infectivity is completely abolished upon proteolytic cleavage [107] (Figure 9). Alternatively, phages can be engineered to make the remaining wild-type g3p itself susceptible to the proteolytic attack [136]. The principle of protease selection can also be

44 Engineering Proteins for Stability and Efficient Folding

Fig. 9 Strategies for selecting for improved protein stability and folding by phage display. (a) Methods utilizing the binding to a given target (affinity selection) as a means for selecting members with improved folding behavior and higher stability from a protein library. Several types of selection pressure can be applied in order to recover mutants with enhanced stability. Destabilizing mutations are deliberately introduced (top) to render the protein “nonfunctional,” and alternative stabilizing mutations are identified by selecting variants whose functionality is regained. Alternatively (bottom), elevated temperatures or denaturing agents can be used to render most of the protein variants “nonfunctional,” allowing the selection of simply the “fittest” members. (b) The resistance of the target protein to a protease can be combined with the ability of phages

to re-infect bacteria. The protein library is inserted between the C-terminal domain and the N-terminal domain in all copies of the g3p protein (cloned into the phage genome), which are needed for reinfection of bacterial cells. By proteolytic cleavage of the inserted protein, these domains are cut off and phage infectivity is lost. In alternative approaches, an N-terminal tag sequence is fused to the protein of interest instead of the N-terminal g3p domains. Upon proteolytic cleavage, the tag is lost. In contrast, phages presenting proteins that are resistant to proteolytic cleavage can subsequently be captured on an affinity matrix binding to the tag sequence. In order to further increase the selection threshold, the shown selection strategies can also be combined.

5 Conclusions

applied to in vitro selection techniques in which the displayed protein is freely accessible [120]. Even though proteolysis seems at first glance to be less correlated to the stability of proteins than temperature, it is suitable for optimizing packing of the hydrophobic core [139] as well as for optimizing electrostatic interactions on the protein surface [59]. One reason for the strength of this approach is based on the fact that proteolytic cleavage is an irreversible reaction, which is not necessarily the case for denaturation by temperature or denaturing agents. Furthermore, protease resistance monitors the flexibility of the polypeptide chain rather than complete denaturation. It is therefore capable not only of selecting against completely unfolded variants but also of detecting local unfolding events. Because sites of local unfolding often initiate the global unfolding process, it can be advantageous to remove such sites. Nevertheless, the effective cleavage of flexible parts of the protein restricts the method to proteins that do not have extended flexible regions in the native state. Furthermore, a selection against the primary recognition sequence of the protease is clearly a possible outcome of such experiments. None of the described methods can stand completely on its own. Many methods can easily be combined to increase the stringency of a selection. A combination of temperature stress and increasing amounts of denaturing agents may, for example, lead to higher flexibility in certain regions of the protein, thereby increasing the sensitivity of the subsequent proteolytic attack. Additionally, temperature and the concentration of denaturing agents allow a much tighter control of the selection pressure than do increasing concentrations of protease and thus open the possibility of a well-controlled gradual increase of selection stringency.

5 Conclusions

Several different approaches are now available for engineering proteins for enhanced biophysical properties. In many cases, few and specific mutations are sufficient to provide proteins of marginal stability with considerably stabilizing features. Thus, the challenge for protein engineering is to identify these positions in a given sequence among the vast number of possible changes and to correctly alter them. Each of the applied techniques has its own merits and bottlenecks. Instead of playing them off against each other, the future challenge will be to find ways to synergistically use them to improve a given molecular property. Evolutionary methods have the advantage of being much less biased by theoretical assumptions or working hypotheses. Additionally, proteins stable in new environments may be evolved, e.g., proteins that fulfill a given function in nonaqueous solutions or high concentrations of detergents. Because the biophysical principles in such environments are of a different nature, rational design has to rely on a much smaller empirical dataset. While rational approaches are likely to become more important as more structural and experimental data become available, notably also those from selection experiments, the large number of variations

45

46 Engineering Proteins for Stability and Efficient Folding

that are potentially able to improve the biophysical properties of a protein often still exceeds the experimentally accessible number. Moreover, because rational engineering has to rely on the available dataset, mutations that lie off the beaten track will rarely be identified. Nevertheless, selection experiments often identify the same “key” mutations in proteins. It is then useful to exploit this information and directly introduce such mutations. A “rational” analysis can also help to recombine important “key” mutations in selected clones and to reduce the effects of a selection-neutral genetic drift. Another combination of rational and combinatorial methods is the creation of “smart” libraries of variants. Library design that is based on such structural considerations and principles will therefore allow more accurate focusing of the selection on specific regions of interest and thereby increase the chances for success. Acknowledgements

The authors thank Drs. Daniela R¨othlisberger, Casim Sarkar and Annemarie Honegger for critical reading of the manuscript. B. S. was the recipient of a Kekule fellowship from the Fonds der Chemischen Industrie. References 1 TAVERNA, D. M. & GOLDSTEIN, R. A. (2002). Why are proteins marginally stable?. Proteins 46, 105–109. 2 ZAVODSZKY, P., KARDOS, J., SVINGOR & PETSKO, G. A. (1998). Adjustment of conformational flexibility is a key event in the thermal adaptation of proteins. Proc. Natl. Acad. Sci. U.S.A. 95, 7406–7411. 3 ARNOLD, F. H., GIVER, L., GERSHENSON, A., ZHAO, H. & MIYAZAKI, K. (1999). Directed evolution of mesophilic enzymes into their thermophilic counterparts. Ann. N.Y. Acad. Sci. 870, 400–403. 4 GOVINDARAJAN, S. & GOLDSTEIN, R. A. (1995). Searching for Foldable Protein Structures Using Optimized Energy Functions. Biopolymers 36, 43–51. 5 DILL, K. A. (1990). Dominant forces in protein folding. Biochemistry 29, 7133–7155. 6 PAPPENBERGER, G., SCHURIG, H. & JAENICKE, R. (1997). Disruption of an ionic network leads to accelerated thermal denaturation of D-glyceraldehyde-3-phosphate

7

8

9

10

11

dehydrogenase from the hyperthermophilic bacterium Thermotoga maritima. J. Mol. Biol. 274, 676–683. STERNER, R. & LIEBL, W. (2001). Thermophilic adaptation of proteins. Crit. Rev. Biochem. Mol. Biol. 36, 39– 106. ZLOTNICK, A. & STRAY, S. J. (2003). How does your virus grow? Understanding and interfering with virus assembly. Trends Biotechnol. 21, 536–542. JASWAL, S. S., SOHL, J. L., DAVIS, J. H. & AGARD, D. A. (2002). Energetic landscape of alpha-lytic protease optimizes longevity through kinetic stability. Nature 415, 343–346. FERSHT, A. R., BYCROFT, M., HOROVITZ, A., KELLIS, J. T., Jr., MATOUSCHEK, A. & SERRANO, L. (1991). Pathway and stability of protein folding. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 332, 171–176. J¨AGER, M. & PLU¨ CKTHUN, A. (1999). Domain interactions in antibody Fv and scFv fragments: effects on unfolding kinetics and equilibria. FEBS Lett. 462, 307–312.

References 12 WILLUDA, J., HONEGGER, A., WAIBEL, R., SCHUBIGER, P. A., STAHEL, R., ZANGEMEISTER-WITTKE, U. & PLU¨ CKTHUN, A. (1999). High thermal stability is essential for tumor targeting of antibody fragments: engineering of a humanized anti-epithelial glycoprotein-2 (epithelial cell adhesion molecule) single-chain Fv fragment. Cancer Res. 59, 5758– 5767. 13 EWERT, S., HUBER, T., HONEGGER, A. & PLU¨ CKTHUN, A. (2003). Biophysical properties of human antibody variable domains. J. Mol. Biol. 325, 531–553. 14 ELLGAARD, L., MOLINARI, M. & HELENIUS, A. (1999). Setting the standards: quality control in the secretory pathway. Science 286, 1882–1888. 15 MARTINEAU, P. & BETTON, J. M. (1999). in vitro folding and thermodynamic stability of an antibody fragment selected in vivo for high expression levels in Escherichia coli cytoplasm. J. Mol. Biol. 292, 921–929. 16 SCHULER, B. & SECKLER, R. (1998). P22 tailspike folding mutants revisited: effects on the thermodynamic stability of the isolated beta-helix domain. J. Mol. Biol. 281, 227–234. 17 PROBA, K., W¨ORN, A., HONEGGER, A. & PLU¨ CKTHUN, A. (1998). Antibody scFv fragments without disulfide bonds made by molecular evolution. J. Mol. Biol. 275, 245–253. 18 KNAPPIK, A. & PLU¨ CKTHUN, A. (1995). Engineered turns of a recombinant antibody improve its in vivo folding. Protein Eng. 8, 81–89. 19 W¨ORN, A. & PLU¨ CKTHUN, A. (1999). Different equilibrium stability behavior of ScFv fragments: identification, classification, and improvement by protein engineering. Biochemistry 38, 8739–8750. 20 STEIPE, B., SCHILLER, B., PLU¨ CKTHUN, A. & STEINBACHER, S. (1994). Sequence statistics reliably predict stabilizing mutations in a protein domain. J. Mol. Biol. 240, 188–192. 21 LEHMANN, M. & WYSS, M. (2001). Engineering proteins for thermostability: the use of sequence alignments versus rational design and

22

23

24

25

26

27

28

29

30

directed evolution. Curr. Opin. Biotechnol. 12, 371–375. KNAPPIK, A., GE, L., HONEGGER, A., PACK, P., FISCHER, M., WELLNHOFER, G., HOESS, A., WOLLE, J., PLU¨ CKTHUN, A. & VIRNEKA¨ S, B. (2000). Fully synthetic human combinatorial antibody libraries (HuCAL) based on modular consensus frameworks and CDRs randomized with trinucleotides. J. Mol. Biol. 296, 57–86. MOSAVI, L. K., MINOR, D. L., Jr. & PENG, Z. Y. (2002). Consensus-derived structural determinants of the ankyrin repeat motif. Proc. Natl. Acad. Sci. U.S.A. 99, 16029–16034. WANG, Q., BUCKLE, A. M., FOSTER, N. W., JOHNSON, C. M. & FERSHT, A. R. (1999). Design of highly stable functional GroEL minichaperones. Protein Sci. 8, 2186–2193. LEHMANN, M., LOCH, C., MIDDENDORF, A., STUDER, D., LASSEN, S. F., PASAMONTES, L., van LOON, A. P. & WYSS, M. (2002). The consensus concept for thermostability engineering of proteins: further proof of concept. Protein Eng. 15, 403–411. BINZ, H. K., STUMPP, M. T., FORRER, P., AMSTUTZ, P. & PLU¨ CKTHUN, A. 39 Engineering Proteins for Stability and Efficient Folding (2003). Designing repeat proteins: well-expressed, soluble and stable proteins from combinatorial libraries of consensus ankyrin repeat proteins. J. Mol. Biol. 332, 489–503. MAIN, E. R., XIONG, Y., COCCO, M. J., D’ANDREA, L. & REGAN, L. (2003). Design of stable alpha-helical arrays from an idealized TPR motif. Structure (Camb) 11, 497–508. STUMPP, M. T., FORRER, P., BINZ, H. K. & PLU¨ CKTHUN, A. (2003). Designing repeat proteins: modular leucine-rich repeat protein libraries based on the mammalian ribonuclease inhibitor family. J. Mol. Biol. 332, 471–487. FORRER, P., BINZ, H. K., STUMPP, M. T. & PLU¨ CKTHUN, A. (2004). Consensus design of repeat proteins. Chembiochem 5, 183–189. KOHL, A., BINZ, H. K., FORRER, P., STUMPP, M. T., PLU¨ CKTHUN, A. & GRU¨ TTER, M. G. (2003). Designed to be

47

48 Engineering Proteins for Stability and Efficient Folding

31

32

33

34

35

36

37

38

39

40

41

stable: crystal structure of a consensus ankyrin repeat protein. Proc. Natl. Acad. Sci. U.S.A. 100, 1700–1705. LEVITT, M., GERSTEIN, M., HUANG, E., SUBBIAH, S. & TSAI, J. (1997). Protein folding: the endgame. Annu. Rev. Biochem. 66, 549–579. BOHM, G. & JAENICKE, R. (1994). Relevance of sequence statistics for the properties of extremophilic proteins. Int. J. Pept. Protein Res. 43, 97–106. JAENICKE, R. & BOHM, G. (1998). The stability of proteins in extreme environments. Curr. Opin. Struct. Biol. 8, 738–748. VIEILLE, C. & ZEIKUS, G. J. (2001). Hyperthermophilic enzymes: sources, uses, and molecular mechanisms for thermostability. Microbiol. Mol. Biol. Rev. 65, 1–43. van den BURG, B. & EIJSINK, V. G. (2002). Selection of mutations for increased protein stability. Curr. Opin. Biotechnol. 13, 333–337. MATSUMURA, M., SIGNOR, G. & MATTHEWS, B. W. (1989). Substantial increase of protein stability by multiple disulphide bonds. Nature 342, 291–293. PERRY, L. J. & WETZEL, R. (1984). Disulfide bond engineered into T4 lysozyme: stabilization of the protein toward thermal inactivation. Science 226, 555–557. PACE, C. N., GRIMSLEY, G. R., THOMSON, J. A. & BARNETT, B. J. (1988). Conformational stability and activity of ribonuclease T1 with zero, one, and two intact disulfide bonds. J. Biol. Chem. 263, 11820–11825. ZHANG, T., BERTELSEN, E. & ALBER, T. (1994). Entropic effects of disulphide bonds on protein stability. Nat. Struct. Biol. 1, 434–438. DOIG, A. J. & WILLIAMS, D. H. (1991). Is the hydrophobic effect stabilizing or destabilizing in proteins? The contribution of disulphide bonds to protein stability. J. Mol. Biol. 217, 389–398. MARTENSSON, L. G., KARLSSON, M. & CARLSSON, U. (2002). Dramatic stabilization of the native state of human carbonic anhydrase II by an engineered

42

43

44

45

46

47

48

49

50

51

disulfide bond. Biochemistry 41, 15867–15875. IVENS, A., MAYANS, O., SZADKOWSKI, H., JURGENS, C., WILMANNS, M. & KIRSCHNER, K. (2002). Stabilization of a (βα)8 -barrel protein by an engineered disulfide bridge. Eur. J. Biochem. 269, 1145–1153. BETZ, S. F. & PIELAK, G. J. (1992). Introduction of a disulfide bond into cytochrome c stabilizes a compact denatured state. Biochemistry 31, 12337–12344. IWAI, H. & PLU¨ CKTHUN, A. (1999). Circular beta-lactamase: stability enhancement by cyclizing the backbone. FEBS Lett. 459, 166–172. SCOTT, C. P., ABEL-SANTOS, E., WALL, M., WAHNON, D. C. & BENKOVIC, S. J. (1999). Production of cyclic peptides and proteins in vivo. Proc. Natl. Acad. Sci. U.S.A. 96, 13638–13643. CAMARERO, J. A., FUSHMAN, D., SATO, S., GIRIAT, I., COWBURN, D., RALEIGH, D. P. & MUIR, T. W. (2001). Rescuing a destabilized protein fold through backbone cyclization. J. Mol. Biol. 308, 1045–1062. NAGI, A. D. & REGAN, L. (1997). An inverse correlation between loop length and stability in a four-helix-bundle protein. Fold. Des. 2, 67–75. RUSSELL, R. J., FERGUSON, J. M., HOUGH, D. W., DANSON, M. J. & TAYLOR, G. L. (1997). The crystal structure of citrate synthase from the hyperthermophilic archaeon pyrococcus furiosus at 1.9 Å resolution. Biochemistry 36, 9983–9994. MACEDO-RIBEIRO, S., DARIMONT, B., STERNER, R. & HUBER, R. (1996). Small structural changes account for the high thermostability of 1[4Fe-4S] ferredoxin from the hyperthermophilic bacterium Thermotoga maritima. Structure 4, 1291–1301. MATTHEWS, B. W., NICHOLSON, H. & BECKTEL, W. J. (1987). Enhanced protein thermostability from site-directed mutations that decrease the entropy of unfolding. Proc. Natl. Acad. Sci. U.S.A. 84, 6663–6667. SRIPRAPUNDH, D., VIEILLE, C. & ZEIKUS, J. G. (2000). Molecular determinants of xylose isomerase thermal stability and

References

52

53

54

55

56

57

58

59

60

61

activity: analysis of thermozymes by site-directed mutagenesis. Protein Eng. 13, 259–265. PACE, C. N. (1992). Contribution of the hydrophobic effect to globular protein stability. J. Mol. Biol. 226, 29–35. LIM, W. A., HODEL, A., SAUER, R. T. & RICHARDS, F. M. (1994). The crystal structure of a mutant protein with altered but improved hydrophobic core packing. Proc. Natl. Acad. Sci. U.S.A. 91, 423–427. VENTURA, S., VEGA, M. C., LACROIX, E., ANGRAND, I., SPAGNOLO, L. & SERRANO, L. (2002). Conformational strain in the hydrophobic core and its implications for protein folding and design. Nat. Struct. Biol. 9, 485–493. OHMURA, T., UEDA, T., OOTSUKA, K., SAITO, M. & IMOTO, T. (2001). Stabilization of hen egg white lysozyme by a cavity-filling mutation. Protein Sci. 10, 313–320. CHEN, J. & STITES, W. E. (2001). Higher-order packing interactions in triple and quadruple mutants of staphylococcal nuclease. Biochemistry 40, 14012–14019. HENDSCH, Z. S. & TIDOR, B. (1994). Do salt bridges stabilize proteins? A continuum electrostatic analysis. Protein Sci. 3, 211–226. PACE, C. N. (2001). Polar group burial contributes more to protein stability than nonpolar group burial. Biochemistry 40, 310–313. MARTIN, A., SIEBER, V. & SCHMID, F. X. (2001). In-vitro selection of highly stabilized protein variants with optimized surface. J. Mol. Biol. 309, 717–726. LOLADZE, V. V., IBARRA-MOLERO, B., SANCHEZ-RUIZ, J. M. & MAKHATADZE, G. I. (1999). Engineering a thermostable protein via optimization of charge-charge interactions on the protein surface. Biochemistry 38, 16419–16423. SHOEMAKER, K. R., KIM, P. S., YORK, E. J., STEWART, J. M. & BALDWIN, R. L. (1987). Tests of the helix dipole model for stabilization of alpha-helices. Nature 326, 563–567.

62 PACE, C. N., SHIRLEY, B. A., MCNUTT, M. & GAJIWALA, K. (1996). Forces contributing to the conformational stability of proteins. FASEB J. 10, 75–83. 63 LOLADZE, V. V., ERMOLENKO, D. N. & MAKHATADZE, G. I. (2002). Thermodynamic consequences of burial of polar and non-polar amino acid residues in the protein interior. J. Mol. Biol. 320, 343–357. 64 MCDONALD, I. K. & THORNTON, J. M. (1994). Satisfying hydrogen bonding potential in proteins. J. Mol. Biol. 238, 777–793. 65 HENNIG, M., DARIMONT, B., STERNER, R., KIRSCHNER, K. & JANSONIUS, J. N. (1995). 2.0 Å structure of indole-3-glycerol phosphate synthase from the hyperthermophile Sulfolobus solfataricus: possible determinants of protein stability. Structure 3, 1295– 1306. 66 RAMACHANDRAN, G. N., RAMAKRISHNAN, C. & SASISEKHARAN, V. (1963). Stereochemistry of polypeptide chain configurations. J. Mol. Biol. 7, 95–99. 67 KIMURA, S., KANAYA, S. & NAKAMURA, H. (1992). Thermostabilization of Escherichia coli ribonuclease HI by replacing left-handed helical Lys95 with Gly or Asn. J. Biol. Chem. 267, 22014–22017. 68 TAKANO, K., YAMAGATA, Y. & YUTANI, K. (2001). Role of amino acid residues in left-handed helical conformation for the conformational stability of a protein. Proteins 45, 274–280. 69 STITES, W. E., MEEKER, A. K. & SHORTLE, D. (1994). Evidence for strained interactions between side-chains and the polypeptide backbone. J. Mol. Biol. 235, 27–32. 70 VEGA, M. C., MARTINEZ, J. C. & SERRANO, L. (2000). Thermodynamic and structural characterization of Asn and Ala residues in the disallowed II region of the Ramachandran plot. Protein Sci. 9, 2322–2328. 71 GUNASEKARAN, K., RAMAKRISHNAN, C. & BALARAM, P. (1996). Disallowed Ramachandran conformations of amino acid residues in protein structures. J. Mol. Biol. 264, 191–198.

49

50 Engineering Proteins for Stability and Efficient Folding

72 PAL, D. & CHAKRABARTI, P. (2002). On residues in the disallowed region of the Ramachandran map. Biopolymers 63, 195–206. 73 EWERT, S., HONEGGER, A. & PLU¨ CKTHUN, A. (2003). Structure-based improvement of the biophysical properties of immunoglobulin VH domains with a generalizable approach. Biochemistry 42, 1517–1528. 74 NIEBA, L., HONEGGER, A., KREBBER, C. & PLU¨ CKTHUN, A. (1997). Disrupting the hydrophobic patches at the antibody variable/constant domain interface: improved in vivo folding and physical characterization of an engineered scFv fragment. Protein Eng. 10, 435–444. 75 BRANDTS, J. F., HU, C. Q., LIN, L. N. & MOS, M. T. (1989). A simple model for proteins with interacting domains. Applications to scanning calorimetry data. Biochemistry 28, 8588–8596. 76 W¨ORN, A. & PLU¨ CKTHUN, A. (2001). Stability engineering of antibody single-chain Fv fragments. J. Mol. Biol. 305, 989–1010. 77 GOKHALE, R. S., AGARWALLA, S., FRANCIS, V. S., SANTI, D. V. & BALARAM, P. (1994). Thermal stabilization of thymidylate synthase by engineering two disulfide bridges across the dimer interface. J. Mol. Biol. 235, 89–94. 78 HUDSON, P. J. (1999). Recombinant antibody constructs in cancer therapy. Curr. Opin. Immunol. 11, 548–557. 79 PLU¨ CKTHUN, A., KREBBER, A., KREBBER, C., HORN, U., KNU¨ PFER, U., WENDEROTH, R., NIEBA, L., PROBA, K. & RIESENBERG, D. (1996). Producing antibodies in Escherichia coli: From PCR to fermentation. In Antibody Engineering (MCCAFFERTY, J., HOOGEN-BOOM, H. R. & CHISWELL, D. J., eds.), pp. 203–252. IRL Press, Oxford. 80 HONEGGER, A. & PLU¨ CKTHUN, A. (2001). The influence of the buried glutamine or glutamate residue in position 6 on the structure of immunoglobulin variable domains. J. Mol. Biol. 309, 687–699. 81 JUNG, S., SPINELLI, S., SCHIMMELE, B., HONEGGER, A., PUGLIESE, L., CAMBILLAU, C. & PLU¨ CKTHUN, A. (2001). The importance of framework residues H6, H7 and H10 in antibody heavy chains:

82

83

84

85

86

87

88

89

90

91

experimental evidence for a new structural subclassification of antibody VH domains. J. Mol. Biol. 309, 701–716. GLOCKSHUBER, R., MALIA, M., PFITZINGER, I. & PLU¨ CKTHUN, A. (1990). A comparison of strategies to stabilize immunoglobulin Fv-fragments. Biochemistry 29, 1362–1367. JUNG, S. H., PASTAN, I. & LEE, B. (1994). Design of interchain disulfide bonds in the framework region of the Fv fragment of the monoclonal antibody B3. Proteins 19, 35–47. W¨ORN, A. & PLU¨ CKTHUN, A. (1998). Mutual stabilization of VL and VH in single-chain antibody fragments, investigated with mutants engineered for stability. Biochemistry 37, 13120–13127. YOUNG, N. M., MACKENZIE, C. R., NARANG, S. A., OOMEN, R. P. & BAENZIGER, J. E. (1995). Thermal stabilization of a single-chain Fv antibody fragment by introduction of a disulphide bond. FEBS Lett. 377, 135–139. ARKIN, M. R. & WELLS, J. A. (1998). Probing the importance of second sphere residues in an esterolytic antibody by phage display. J. Mol. Biol. 284, 1083–1094. LOW, N. M., HOLLIGER, P. H. & WINTER, G. (1996). Mimicking somatic hypermutation: affinity maturation of antibodies displayed on bacteriophage using a bacterial mutator strain. J. Mol. Biol. 260, 359–368. STEMMER, W. P. (1994). Rapid evolution of a protein in vitro by DNA shuffling. Nature 370, 389–391. ZHAO, H., GIVER, L., SHAO, Z., AFFHOLTER, J. A. & ARNOLD, F. H. (1998). Molecular evolution by staggered extension process (StEP) in vitro recombination. Nat. Biotechnol. 16, 258–261. AMSTUTZ, P., FORRER, P., ZAHND, C. & PLU¨ CKTHUN, A. (2001). in vitro display technologies: novel developments and applications. Curr. Opin. Biotechnol. 12, 400–405. FORRER, P., JUNG, S. & PLU¨ CKTHUN, A. (1999). Beyond binding: using phage

References

92

93

94

95

96

97

98

99

100

101

display to select for structure, folding and enzymatic activity in proteins. Curr. Opin. Struct. Biol. 9, 514–520. ZHAO, H. & ARNOLD, F. H. (1997). Combinatorial protein design: strategies for screening protein libraries. Curr. Opin. Struct. Biol. 7, 480–485. LEHMANN, M., PASAMONTES, L., LASSEN, S. F. & WYSS, M. (2000). The consensus concept for thermostability engineering of proteins. Biochim. Biophys. Acta 1543, 408–415. MATTHEAKIS, L. C., BHATT, R. R. & DOWER, W. J. (1994). An in vitro polysome display system for identifying ligands from very large peptide libraries. Proc. Natl. Acad. Sci. U.S.A. 91, 9022–9026. HANES, J. & PLU¨ CKTHUN, A. (1997). in vitro selection and evolution of functional proteins by using ribosome display. Proc. Natl. Acad. Sci. U.S.A. 94, 4937–4942. HE, M. & TAUSSIG, M. J. (1997). Antibody-ribosome-mRNA (ARM) complexes as efficient selection particles for in vitro display and evolution of antibody combining sites. Nucleic Acids Res 25, 5132–5134. ROBERTS, R. W. & SZOSTAK, J. W. (1997). RNA-peptide fusions for the in vitro selection of peptides and proteins. Proc. Natl. Acad. Sci. U.S.A. 94, 12297–12302. KURZ, M., GU, K., AL-GAWARI, A. & LOHSE, P. A. (2001). cDNA – protein fusions: covalent protein-gene conjugates for the in vitro selection of peptides and proteins. ChemBioChem 2, 666–672. TAWFIK, D. S. & GRIFFITHS, A. D. (1998). Man-made cell-like compartments for molecular evolution. Nat. Biotechnol. 16, 652–656. DOI, N. & YANAGAWA, H. (1999). STABLE: protein-DNA fusion system for screening of combinatorial protein libraries in vitro. FEBS Lett. 457, 227–230. SMITH, G. P. (1985). Filamentous fusion phage: novel expression vectors that display cloned antigens on the virion surface. Science 228, 1315–1317.

102 GEORGIOU, G., STATHOPOULOS, C., DAUGHERTY, P. S., NAYAK, A. R., IVERSON, B. L. & CURTISS, R., 3rd. (1997). Display of heterologous proteins on the surface of microorganisms: from the screening of combinatorial libraries to live recombinant vaccines. Nat. Biotechnol. 15, 29–34. 103 BODER, E. T. & WITTRUP, K. D. (1997). Yeast surface display for screening combinatorial polypeptide libraries. Nat. Biotechnol. 15, 553–557. 104 JUNG, S. & PLU¨ CKTHUN, A. (1997). Improving in vivo folding and stability of a single-chain Fv antibody fragment by loop grafting. Protein Eng. 10, 959– 966. 105 HOESS, R. H. (2001). Protein design and phage display. Chem. Rev. 101, 3205–3218. 106 FINUCANE, M. D., TUNA, M., LEES, J. H. & WOOLFSON, D. N. (1999). Core-directed protein design. I. An experimental method for selecting stable proteins from combinatorial libraries. Biochemistry 38, 11604–11612. 107 SIEBER, V., PLU¨ CKTHUN, A. & SCHMID, F. X. (1998). Selecting proteins with improved stability by a phage-based method. Nat. Biotechnol. 16, 955–960. 108 FIELDS, S. & SONG, O. (1989). A novel genetic system to detect protein-protein interactions. Nature 340, 245–246. 109 BAI, C. & ELLEDGE, S. J. (1996). Gene identification using the yeast two-hybrid system. Methods Enzymol. 273, 331–347. 110 JOHNSSON, N. & VARSHAVSKY, A. (1994). Split ubiquitin as a sensor of protein interactions in vivo. Proc. Natl. Acad. Sci. U.S.A. 91, 10340–10344. 111 MICHNICK, S. W. (2001). Exploring protein interactions by interaction-induced folding of proteins from complementary peptide fragments. Curr. Opin. Struct. Biol. 11, 472–477. 112 WALDO, G. S. (2003). Genetic screens and directed evolution for protein solubility. Curr. Opin. Chem. Biol. 7, 33–38. 113 WIGLEY, W. C., STIDHAM, R. D., SMITH, N. M., HUNT, J. F. & THOMAS, P. J. (2001). Protein solubility and folding monitored in vivo by structural

51

52 Engineering Proteins for Stability and Efficient Folding

114

115

116

117

118

119

120

121

122

123

complementation of a genetic marker protein. Nat. Biotechnol. 19, 131–136. WALDO, G. S., STANDISH, B. M., BERENDZEN, J. & TERWILLIGER, T. C. (1999). Rapid protein-folding assay using green fluorescent protein. Nat. Biotechnol. 17, 691–695. PEDELACQ, J. D., PILTCH, E., LIONG, E. C., BERENDZEN, J., KIM, C. Y., RHO, B. S., PARK, M. S., TERWILLIGER, T. C. & WALDO, G. S. (2002). Engineering soluble proteins for structural genomics. Nat. Biotechnol. 20, 927–932. WURTH, C., GUIMARD, N. K. & HECHT, M. H. (2002). Mutations that reduce aggregation of the Alzheimer’s Aβ42 peptide: an unbiased search for the sequence determinants of Aβ amyloidogenesis. J. Mol. Biol. 319, 1279–1290. MAXWELL, K. L., MITTERMAIER, A. K., FORMAN-KAY, J. D. & DAVIDSON, A. R. (1999). A simple in vivo assay for increased protein solubility. Protein Sci. 8, 1908–1911. LESLEY, S. A., GRAZIANO, J., CHO, C. Y., KNUTH, M. W. & KLOCK, H. E. (2002). Gene expression response to misfolded protein as a screen for soluble recombinant protein. Protein Eng. 15, 153–160. MINARD, P., SCALLEY-KIM, M., WATTERS, A. & BAKER, D. (2001). A “loop entropy reduction” phage-display selection for folded amino acid sequences. Protein Sci. 10, 129–134. MATSUURA, T. & PLU¨ CKTHUN, A. (2003). Selection based on the folding properties of proteins with ribosome display. FEBS Lett. 539, 24–28. GU, H., YI, Q., BRAY, S. T., RIDDLE, D. S., SHIAU, A. K. & BAKER, D. (1995). A phage display system for studying the sequence determinants of protein folding. Protein Sci. 4, 1108–1117. KIM, D. E., GU, H. & BAKER, D. (1998). The sequences of small proteins are not extensively optimized for rapid folding by natural selection. Proc. Natl. Acad. Sci. U.S.A. 95, 4982–4986. KOWALSKI, J. M., PAREKH, R. N., MAO, J. & WITTRUP, K. D. (1998). Protein folding stability can determine the efficiency of

124

125

126

127

128

129

130

131

132

133

escape from endoplasmic reticulum quality control. J. Biol. Chem. 273, 19453–19458. SHUSTA, E. V., KIEKE, M. C., PARKE, E., KRANZ, D. M. & WITTRUP, K. D. (1999). Yeast polypeptide fusion surface display levels predict thermal stability and soluble secretion efficiency. J. Mol. Biol. 292, 949–956. SHUSTA, E. V., HOLLER, P. D., KIEKE, M. C., KRANZ, D. M. & WITTRUP, K. D. (2000). Directed evolution of a stable scaffold for T-cell receptor engineering. Nat. Biotechnol. 18, 754–759. BODER, E. T. & WITTRUP, K. D. (2000). Yeast surface display for directed evolution of protein expression, affinity, and stability. Methods Enzymol. 328, 430–444. RUAN, B., HOSKINS, J., WANG, L. & BRYAN, P. N. (1998). Stabilizing the subtilisin BPN pro-domain by phage display selection: how restrictive is the amino acid code for maximum protein stability?. Protein Sci. 7, 2345–2353. ZHOU, H. X., HOESS, R. H. & DEGRADO, W. F. (1996). in vitro evolution of thermodynamically stable turns. Nat. Struct. Biol. 3, 446–451. GOTO, Y. & HAMAGUCHI, K. (1979). The role of the intrachain disulfide bond in the conformation and stability of the constant fragment of the immunoglobulin light chain. J. Biochem. (Tokyo). 86, 1433–1441. AKANUMA, S., YAMAGISHI, A., TANAKA, N. & OSHIMA, T. (1999). Further improvement of the thermal stability of a partially stabilized Bacillus subtilis 3-isopropylmalate dehydrogenase variant by random and site-directed mutagenesis. Eur. J. Biochem. 260, 499–504. FARINAS, E. T., BULTER, T. & ARNOLD, F. H. (2001). Directed enzyme evolution. Curr. Opin. Biotechnol. 12, 545–551. ZHAO, H. & ARNOLD, F. H. (1999). Directed evolution converts subtilisin E into a functional equivalent of thermitase. Protein Eng. 12, 47–53. GIVER, L., GERSHENSON, A., FRESKGARD, P. O. & ARNOLD, F. H. (1998). Directed evolution of a thermostable esterase.

References Proc. Natl. Acad. Sci. U.S.A. 95, 12809–12813. 134 JERMUTUS, L., HONEGGER, A., SCHWESINGER, F., HANES, J. & PLU¨ CKTHUN, A. (2001). Tailoring in vitro evolution for protein affinity or stability. Proc. Natl. Acad. Sci. U.S.A. 98, 75–80. 135 JUNG, S., HONEGGER, A. & PLU¨ CKTHUN, A. (1999). Selection for improved protein stability by phage display. J. Mol. Biol. 294, 163–180. 136 KRISTENSEN, P. & WINTER, G. (1998). Proteolytic selection for protein folding using filamentous bacteriophages. Fold. Des. 3, 321–328.

137 FONTANA, A., POLVERINO DE LAURETO, P., De FILIPPIS, V., SCARAMELLA, E. & ZAMBONIN, M. (1997). Probing the partly folded states of proteins by limited proteolysis. Fold. Des. 2, R17–26. 138 KREBBER, C., SPADA, S., DESPLANCQ, D. & PLU¨ CKTHUN, A. (1995). Co-selection of cognate antibody-antigen pairs by selectively-infective phages. FEBS Lett. 377, 227–231. 139 FINUCANE, M. D. & WOOLFSON, D. N. (1999). Core-directed protein design. II. Rescue of a multiply mutated and destabilized variant of ubiquitin. Biochemistry 38, 11613–11623.

53

1

Protein-based Artiﬁcial Enzymes Ben Duckworth and Mark D. Distefano

University of Minnesota, Minneapolis, USA

Originally published in: Artiﬁcial Enzymes. Edited by Ronald Breslow. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31165-1

1 Introduction

The design of catalysts that rival the speciﬁcity and speed of natural enzymes is a challenging objective. Various approaches have been employed to create such molecules. Polypeptides are a logical choice of possible framework for catalyst design because they provide a simple means for generating complex structures. The catalyst structure can be changed by altering the corresponding gene and expressing the desired protein in bacteria. However, chemical methods can also be incorporated into such an approach to greatly expand the range of possible reactions that can be studied. In such a chemogenetic strategy, a small molecule-based catalyst is coupled to a protein scaffold devoid of catalytic activity. The small molecule provides intrinsic reactivity while the protein component controls speciﬁcity; the protein can also be used to tune or augment the reactivity of the small molecule catalyst. Such an approach provides enormous ﬂexibility in the design of new catalytic materials. Early efforts in this ﬁeld focused on using chemical modiﬁcation to alter the type of reaction catalyzed by existing enzymes or to change their substrate speciﬁcity. Thiosubtilisin [1, 2], selenosubtilisin [3] and ﬂavopapain [4] are all examples of proteins that manifest altered reactivity resulting from chemical modiﬁcation. Other work with sta-phylococcal nuclease [5] and subtilisin [6, 7] employed chemical modiﬁcation to modulate substrate selectivity. While that pioneering work served as the foundation for protein-based catalyst design, these efforts were, notably, based on protein frameworks that possessed native catalytic activity. Such proteins already contained highly evolved substrate binding sites and/or catalytic machinery. The present chapter focuses on the design of protein-based catalysts starting with polypeptides that have no intrinsic catalytic activity. While this is an enormous challenge when compared with approaches starting with existing enzymes, it opens Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Protein-based Artiﬁcial Enzymes

up a plethora of exciting possibilities; if it could be accomplished, the ability to create catalysts with enzyme-like properties would have dramatic effects that would extend well beyond the ﬁeld of chemistry. Several recent articles provide a more comprehensive review of the ﬁeld of protein design, including work with existing enzymes [8–19]. 2 Artiﬁcial Nucleases Based on DNA and RNA Binding Proteins 2.1 Introduction

The process of cleaving the phosphodiester bond of RNA and DNA is a vital and widespread phenomenon in nature. Scission of the nucleic acid backbone plays crucial roles in myriad in vivo and in vitro processes, including mutational repair, cell death, cloning, and sequencing. Much attention has been given to the discovery and development of nucleases, both natural and artiﬁcial, which might beneﬁt the ﬁelds of biotechnology and pharmacology [20]. Several chemical agents possess nuclease activity. However, such compounds have low afﬁnity for oligonucleotides and may not be optimally positioned for attack and scission of the backbone. Therefore, if a chemical nuclease could be incorporated into a DNA or RNA binding protein, substrate binding and nuclease activity would be enhanced. 2.2 Artiﬁcial Nucleases from Native Protein Scaffolds

To create an artiﬁcial nuclease, Sigman and co-worker enhanced the nuclease speciﬁcity of 1,10-phenanthroline(OP)-copper by covalently linking the moiety to the Escherichia coli trp repressor (TrpR) [21]. OP is a robust chemical nuclease that cleaves the phosphodiester bonds of nucleic acids. A copper-oxo species attacks the C-1H of the deoxyribose in the minor grove, yielding 3 - and 5 -phosphomonoesters, free base, and 5-methylene furanone (5-MF) (Figure 1) [22, 23]. This makes it possible to convert DNA-binding proteins into nucleases by conjugation with OPcopper. Since most DNA/RNA binding proteins bind the major groove, it is crucial for the OP-copper group to interact solely with the minor groove to avoid disruption of key binding interactions. To conjugate E. coli trp repressor with OP-copper, the four lysine amino groups were ﬁrst converted into sulfhydryl groups by reaction with 2-iminothiolane hydrochloride in the presence of the corepressor L-tryptophan (Figure 2) [21, 24, 25]. The thiols were then alkylated with 5-(iodoacetamido)-1,10-phenanthroline (IOP, Figure 2-1), yielding the TrpR-OP conjugate. DNase I footprinting experiments indicated that the DNA-binding ability of TrpR was not jeopardized after modiﬁcation with OP. Early experiments involved the incubation of trpEDCBA operator with OP-TrpR copper complex for 20 h, which resulted in double-strand scission as well as single-strand nicks in 50% of the labeled strands [21].

2 Artiﬁcial Nucleases Based on DNA and RNA Binding Proteins

Fig. 1 Shortened chemical mechanism for DNA cleavage by 1, 10-phenanthroline-copper complexes.

Fig. 2 (A) Procedure for the attachment of 1,10-phenanthroline to proteins via lysine derivatization. (B) 5-(iodoacetamido)-1, 10-phenanthroline (IOP, 1) and 5-(iodoacetylglycylamido)-1, 10-phenanthroline (IAOP, 2).

3

4 Protein-based Artiﬁcial Enzymes

2.3 OP Nuclease Design by Mutagenesis and Chemical Modiﬁcation

An additional approach to artiﬁcial nuclease design is cysteine mutagenesis followed by conjugation with the desired catalytic group. This avoids the need to convert amino acid residues into reactive thiols, while allowing precise control over the placement of the reagent within the DNA-binding protein. Sigman and coworkers applied this strategy to create an artiﬁcial nuclease using the bacteriophage λCro protein [26]. When complexed with DNA, the C-terminal arm binds within the minor groove and is close to the C-1 hydrogen of the deoxyribose on either DNA strand [26–30]. Therefore, an alanine close to the C-terminus was mutated to a cysteine and derivatized with IOP. After incubation with the Cro A66C-OP conjugate, 40% of the 17 base pair OR-3 operator site was cleaved within 10 min [11]. Thus, placement of the phenanthroline group close to the DNA substrate ensured efﬁcient nuclease activity. Recently, the carboxy terminal domain of NarL, NarLC , was modiﬁed with 1,10phe-nanthroline. NarL is a response regulatory protein of E. coli and binds to a heptameric consensus sequence [31]. Two residues of the C-terminal domain, which were mutated and conjugated with IOP (NarLC K201C-IOP, NarLC K211CIOP), showed high site-speciﬁc cleavage efﬁciency of the top strand (Figure 3). When the mutated NarLC was modiﬁed with IAOP (Figure 2-2), a similar DNA

Fig. 3 Model of the NarLC -DNA complex showing the locations of amino acids 201 and 211. Two molecules of NarLC are positioned above the major groove of the DNA. Color scheme: Protein secondary structure (green), DNA (white), residue 211 (pink), residue 201 (orange).

2 Artiﬁcial Nucleases Based on DNA and RNA Binding Proteins

cleavage pattern was observed on the bottom strand. This IAOP conjugate contains a 4 Å longer linker arm than does the previous IOP conjugate, which allows the OP-Cu complex to extend and cleave the adjacent bases of the DNA substrate. These artiﬁcial cleavage enzymes could ultimately be used to identify the number, position and orientation of NarL monomers on several promoters and may be extended to other regulatory proteins. These results highlight the promise of chemical nucleases as tools for biological investigation. 2.4 Additional Applications for OP Conjugates

Other research groups have also created artiﬁcial nucleases by OP conjugation. Johnson and co-workers created an artiﬁcial nuclease, which aided in the understanding of DNA conformational changes upon protein binding [32]. E. coli Fis (factor for inversion stimulation) was chosen as the protein scaffold and binds to DNA with a low sequence speciﬁcity. X-Ray crystallography and electrophoretic mobility studies indicated signiﬁcant bending of the DNA upon Fis binding [33, 34]. To aid in the study of DNA-Fis complexes, OP-Cu was conjugated to four separate mutated cysteines. Of these four conjugates, N98C-OP and N78C-OP displayed nuclease activity. Scission patterns from these two nucleases not only conﬁrmed earlier studies of DNA bending, but also revealed that the DNA wrapped around the Fis dimer. The E. coli catabolite gene activator protein (CAP) binds to a 22 base-pair, twofold symmetric DNA recognition site with high afﬁnity (K A = 4 × 1010 M−1 ) [35]. Ebright and co-workers successfully converted CAP into a site-speciﬁc DNA cleavage enzyme. In their ﬁrst attempt, OP was conjugated to the only solvent accessible cysteine (C178 ) of CAP. The catalytic group was placed at this position so that intra-CAP interactions would not be disrupted, CAP-DNA contacts would not be comprised, and OP would be in close proximity for a favorable attack on the phosphodiester bond. The binding constant for this conjugate was 1 × 108 M−1 , indicating that the DNA binding ability of CAP was not signiﬁcantly lost due to conjugation with OP. The CAP178-OP conjugate cleaved the DNA site at four adjacent nucleotides (Figure 4).

Fig. 4 DNA cleavage sites produced by the CAP178–1, 10-phenanthroline conjugate. Longer arrows indicate sites where greater cleavage efﬁciency occurred.

5

6 Protein-based Artiﬁcial Enzymes

Earlier work indicated that in the speciﬁc CAP-DNA complex the DNA was bent 90◦ away from the protein but was not distorted in the nonspeciﬁc complex. (CAP bound to its noncognate DNA site) [3, 36]. This DNA bending phenomenon was exploited to construct a new CAP-OP nuclease capable of single site cleavage [37]. The crystal structure of the speciﬁc CAP-DNA complex revealed that amino acids 24–26 and 89–91 of CAP were close to the DNA substrate [38]. With this in mind, residue 26 was mutated to a cysteine and modiﬁed with IOP. While the catalytic group neighbors the DNA in the speciﬁc complex, OP is too far from the substrate in the nonspeciﬁc complex, and thus cannot cleave the DNA. Ultimately, conjugation of OP at position 26 yielded an artiﬁcial nuclease that proceeded to ∼90% cleavage with no detectable nonspeciﬁc cleavage at distal sites.

2.5 A Fe-EDTA Artiﬁcial Nuclease

An alternative DNA-cleaving agent was used to identify the σ subunit DNA contact sites of E. coli RNA polymerase [39]. Iron-EDTA protein conjugates have previously been shown to cleave DNA by producing hydroxyl radicals, which ultimately attack the deoxyribose backbone [40]. To create this artiﬁcial nuclease, Minchin and co-workers mutated several residues within the σ subunit to cysteines. These residues were conjugated with (S)-1-[p-(bromoacetamido)benzyl]-EDTA (BABE) (Figure 5). By identifying the cleavage products of several related promoters, this Fe-BABE conjugate conﬁrmed the location of the σ subunit-promoter DNA contact sites.

2.6 Concluding Remarks

Through direct conjugation of chemical nucleases to DNA binding proteins, several groups have successfully created artiﬁcial nuclease from proteins with no

Fig. 5 Procedure for the derivatization of proteins with (S)-1[p-(bromoacetamido)benzyl]-EDTA (BABE).

3 Catalysts Based on Hollow Lipid-binding Proteins 7

native nuclease ability. These nucleases can utilize the favorable protein–DNA binding elements and can position the hydrolytic metal close to the DNA backbone for efﬁcient cleavage [41]. These nucleases have provided useful biological insights and a better understanding of the molecular details involved in protein–DNA interactions.

3 Catalysts Based on Hollow Lipid-binding Proteins 3.1 Lipid-binding Proteins

Lipid-binding proteins are a class of molecules found in eukaryotic cells involved in the transport of fatty acids and other types of hydrophobic compounds. The protein structure consists of two orthogonal planes of β-sheets that form a cup-shaped cavity that is capped off with a helix-turn-helix element. Considerable structural data, obtained from X-ray and NMR analysis, exists for this family of proteins [42, 43]. Of particular interest is the presence of a large, solvent sequestered, cavity whose overall volume varies between 500 and 1000 A3 depending on the exact identity of the protein; with the exception of certain membrane-bound channel proteins, there are few other examples of macromolecules that possess such a cavity. This structural feature serves as the ligand binding site in the protein cavity for a diverse range of molecules. Such a large, solvent sequestered, binding site also provides a useful scaffold for the design of catalysts. Formally, this cavity can be viewed as a protein equivalent to the cyclodextrin template used in much of the pioneering work of Breslow and co-workers in their development of enzyme mimics [18]. 3.2 Initial Work

In their initial work Distefano and co-workers used the protein ALBP (adipocyte lipid-binding protein) for their protein scaffold. This protein contains a unique cysteine residue a position 117 that can be selectively modiﬁed using reagents that capitalize on the unique reactivity of the thiol side chain. Kuang and co-workers developed a reagent, TP-PX (6-1) that contained an activated disulﬁde suitable for protein derivatization. This molecule was used to incorporate a PX moiety into ALBP at position 117, resulting in the formation of a construct denoted ALBP-PX (Figure 6, 6-2) [44]. This semisynthetic biocatalyst aminated reductively various α-keto acids (7-2) to ami-no acids (7-4) with 0 to 94% ee (Figure 7). Because these reactions were performed in the absence of any additional amine source, only single turnovers were obtained. The reaction rates were not, however, signiﬁcantly faster than those involving free pyridoxamine (7-1b). This suggested that the protein

8 Protein-based Artiﬁcial Enzymes

Fig. 6 Derivatization reagents for the preparation of pyridoxamine conjugates of fatty acid binding proteins.

cavity functions as a chiral environment that controls the facial selectivity of the protonation of the aldimine intermediate without forming speciﬁc interactions with the bound pyridoxamine cofactor, which could accelerate the reaction, as conﬁrmed by a X-ray crystal structure [45]. Modeling of the Schiff base complexes with several amino acids indicated that one face of the putative aldimine intermediate was protected against the approach of the solvent or buffer molecules that must be the proton source for the reaction given the lack of suitable functional groups within the cavity. This structural data provided a rationale for explaining the enantioselectivity observed in the ALBP-PX system. 3.3 Exploiting the Advantage of a Protein-based Scaffold

The next step in developing these protein-based catalysts was to take advantage of the synthetic ﬂexibility provided by a protein scaffold. Since the protein is encoded by a DNA sequence, changes can be made in the gene that will result in modiﬁcations in the protein. Mutagenesis and other genetic methods can produce alterations in the protein framework much more rapidly than the organic synthesis

Fig. 7 Single turnover, reductive amination, reactions promoted by fatty acid binding protein–pyridoxamine conjugates.

3 Catalysts Based on Hollow Lipid-binding Proteins 9

required to make changes in synthetic scaffolds. Initially, the position of catalyst attachment within the cavity was varied to probe its effect on reaction rate and selectivity. For these experiments, a different fatty acid binding protein (IFABP) was employed. The wild-type IFABP contains no cysteine residues in its primary sequence and is, hence, a useful template for the introduction of single cysteine residues at different positions [46]. Cysteine residues were introduced at several positions, including V60 , L72 , and A104 , using site-directed mutagenesis. Figure 8 gives the locations of these mutations within the protein structure. The three different mutant proteins described above were puriﬁed and the corresponding conjugates were prepared using the TP-PX reagent (6-1). The conjugates were then evaluated in single turnover conditions (Figure 7) with several α-keto acids to produce amino acids in enantiomerically enriched form. This collection of catalysts exhibited various differences in reactivity and selectivity. Compared to ALBP-PX, IFABP-PX60 reacted at least 9.4-fold more rapidly, while IFABP-PX72 displayed opposite enantioselectivity, and IFABP-PX104 showed a clear selectivity preference for unbranched substrates [47]. From these experiments, the position of cofactor attachment is clearly an important parameter in modulating the reactivity and speciﬁcity of these biocatalysts. Moreover, these results underscore the utility of site-directed mutagenesis and the power of using a protein-based scaffold for catalyst development.

Fig. 8 Stereo view showing positions 60, 72, 104 and 117 in IFABP that have been used for the attachment of pyridoxamine and other catalytic groups. From top to bottom: A104, Y117, L72 and V60. Color scheme: Protein secondary structure (green), carbon (white), oxygen (red), nitrogen (blue).

10 Protein-based Artiﬁcial Enzymes

Fig. 9 Multiple turnover reactions that produce chiral amino acids from achiral keto acids.

3.4 Catalytic Turnover with Rate Acceleration

Due to its intriguingly more rapid reaction rate than ALBP-PX and free pyridoxamine, IFABP-PX60 was subsequently studied further [48]. Under single turnover conditions, it converted α-keto glutarate (Figure 9, 9-1) into glutamic acid (7-4) 62-fold faster than free pyridoxamine (7-1b); this was determined by evaluating the extent of conversion at much shorter reaction times. This more rapid reaction rate under single turnover conditions suggested that this construct might be capable of catalytic transamination. Using tyrosine or phenylalanine (9-2) as the amine source to recycle the cofactor from the pyridoxal form (7-3a) back to the pyridoxamine form (7-1), L-glutamate (9-3) was formed with an enantiomeric purity of 93% ee. As many as 50 turnovers were observed with long reaction times (14 days). Both kcat and K M for the reaction were determined by kinetic analysis (Table 1). Comparison with similar parameters obtained from reactions with free pyridoxamine indicated that IFABP-PX60 catalyzed transamination some 200 times more efﬁciently. Analysis of the speciﬁc kinetic constants kcat and K M indicated that the observed rate acceleration was due mostly to an increase in substrate binding Table 1 Kinetic constants and catalytic efﬁciencies for semi-synthetic transaminases based

on fatty acid binding proteins

PX MPX IFABP-PX60 IFABP-MPX60 hsIFABP-PX60 IFABP-PxK38 IFABP-MPxK38 IFABP-PxK51 IFABP-MPxK51 IFABP-Px126 IFABP-MPx126 IFABP-Px126/14 IFABP-MPx126/14

KM (mM)

kcat (h−1 )

kcat /KM (h−1 mM−1 )

73 38.7 1.8 6.8 10.2 0.81 13.7 0.24 8.9 5.5 40 6.0 48

0.032 0.031 0.29 0.23 0.22 0.44 1.12 0.44 0.52 0.18 1.10 0.11 0.78

4.4 × 10−4 8.0 × 10−4 0.16 0.034 0.022 0.54 0.08 1.83 0.06 0.03 0.03 0.02 0.02

3 Catalysts Based on Hollow Lipid-binding Proteins 11

(50-fold), with a smaller effect on the maximal rate (4-fold). While this is an impressive result, the absolute magnitude of kcat /K M (0.02 s−1 M−1 ) makes it clear that this catalyst is still quite primitive compared to natural enzyme systems that occasionally operate with catalytic efﬁciencies near the diffusion limit. 3.5 Modulation of Cofactor Reactivity with Metal Ions

Model studies with pyridoxamine and related analogs have shown that transamination reactions promoted by these catalysts can be accelerated by the addition of metal ions, including Zn(II), Cu(II), Ni(II) and Al(III). Hypothetically, these cations accomplish this by stabilizing the formation of Schiff base intermediates and by increasing the acidity of the protons that must be removed in the reaction. Interestingly, the addition of metal ions to reactions catalyzed by ALBP- and IFABP-pyridoxamine conjugates resulted in both positive and negative reaction rate perturbations [49]. IFABP-PX104 reacted 4.7-fold faster in the presence of Cu(II) whereas IFABP-PX60 reacted 4.4-fold slower – both rate effects were accompanied by a decrease in reaction enan-tioselectivity. Little change was observed for the reaction catalyzed by IFABP-PX72. UV/Vis spectroscopy experiments that monitored the formation of metal-aldimine intermediates suggested that IFABP-PX60 and IFABP-PX104 but not IFABP-PX72 formed a complex with Cu(II). Thus, metal ions may be used to increase semisynthetic enzyme efﬁciency, although this did not occur in all cases. These studies also noted that the reactions rates were sensitive to changes in buffer; the use of imidazole resulted in a lower rate than with similar reactions performed in HEPES buffer. These observations raise the possibility that buffer molecules may actually enter the protein cavity and directly participate in the reaction. This makes sense because several proton transfers must occur in a complete transamination reaction and yet there are no functional groups within the cavity that could do this. 3.6 Chemogenetic Approach

In the catalytic mechanism of pyridoxamine-based reactions, the pyridine nitrogen atom must be protonated; this protonation gives the pyridine ring a net positive charge that increases the acidity of the nearby benzylic protons and serves to stabilize a developing negative charge. One way protonation can be effectively enforced is by introducing a permanent positive charge via N-methylation. In some model systems, an N-methylated species accelerates the transamination rate up to 20-fold. In contrast, after reconstitution with N-methylpyridoxamine, alanine aminotransferase lost its activity almost completely (>99.8%). Therefore, N-quaternization may have both positive and negative inﬂuences on the reaction rate. To allow a similar modiﬁcation to be made in these fatty acid binding protein systems, a new reagent containing N-methylpyridoxamine and an activated disulﬁde (TP-MPX, 6-3) was prepared. A conjugate, IFABP-MPX60 (6-4) was prepared [50] and, under

12 Protein-based Artiﬁcial Enzymes

catalytic conditions, signiﬁcant reaction (2 turnovers, 41% ee) was observed using α-keto glutarate and phenylalanine as substrates. Kinetic analysis of the reaction showed that the MPX-containing conjugate gave higher K M (3.8-fold higher than the earlier PX construct) but similar kcat . This indicated that N-methylation had no positive effects on the reaction catalyzed by conjugates based on IFABP-V60 C. However, the MPX cofactor proved to be considerably more useful in subsequent experiments with IFABP variants containing lysine mutations (see below for further discussion of those proteins). In those studies, efforts were made to increase their catalytic efﬁciency by employing a genetic method to enforce N-protonation. Taking a cue from the crystal structure of AATase (aspartate amino transferase), an enzyme that catalyzes transamination using a pyridoxamine cofactor, carboxylatecontaining amino acid residues were introduced close to the pyridine nitrogen locus in two IFABP mutant proteins. It was thought, based on functional studies of AATase, that the positioning of anionic side chains near the pyridine nitrogen center would enforce protonation via an ion pairing mechanism; the presence of the proximal negative charge would favor the protonated pyridine. Several carboxylatecontaining IFABP mutants were prepared for this purpose. Unfortunately, none of these mutants were sufﬁciently stable for conjugate preparation – they precipitated during puriﬁcation, and efforts to refold the denatured protein were unsuccessful. However, in separate experiments, the TP-MPX reagent (6-3) was used to prepare MPX-containing constructs instead (see 6-4 for a generic structure). Interestingly, these conjugates showed enhanced catalytic activity compared to their PX progenitors [51]. Kinetic studies indicated that the kcat (1.12 h−1 for IFABP-MPxK38 and 0.52 h−1 for IFABP-MPxK51) and turnover numbers (12.2 turnovers by IFABPMPxK38 and 5.7 by IFABP-MPx51 in 24 h) observed with these constructs under standard conditions are the highest achieved in this system (see Table 1 for a summary of kinetic parameters). The success of these constructs, prepared using a combination of chemical modiﬁcation of the catalyst structure and genetic manipulation of the protein scaffold, highlights the enormous power and ﬂexibility of this chemogenetic approach for catalyst development. 3.7 Adding Functional Groups within the Cavity

A major goal of research involving these protein scaffolds was to determine whether the ﬂexibility of using a protein scaffold could be fully capitalized upon by using rational design in concert with knowledge of chemical mechanism to improve catalytic efﬁciency. Taking a cue from Nature, based on biochemical experiments with AATase, lysine residues were introduced into the protein cavity to enable Schiff base formation and to serve as general acids and bases in the reaction cycle. Figure 10 shows these functions in an abbreviated transamination mechanism. Based on the crystal structure of IFABP, molecular modeling was employed to identify possible positions where lysine residues could be introduced to perform the functions noted above. Mutants L38 K,V60 C and E51 K,V60 C were prepared and used to create pyridoxamine conjugates [52]. Figure 11 gives a model of IFABP-PXK51.

3 Catalysts Based on Hollow Lipid-binding Proteins 13

Fig. 10 Abbreviated mechanism for transamination reactions promoted by pyridoxal-based catalysts.

The resulting assemblies IFABP-PxK38 and IFABP-PxK51 showed improved kcat and K M . The overall catalytic efﬁciency (kcat /K M ) of IFABP-PxK51 increased 4200-fold compared to unliganded pyridoxamine phosphate and was 12-fold greater than for IFABP-PX60 while maintaining comparable enantioselectivity (83–94% ee). The principal effect on the kinetic constants for the reactions catalyzed by these mutants was on K M . Each conjugate showed a signiﬁcant decrease in

Fig. 11 Stereo representation of a model of IFABP-PXK51. The overall protein structure is shown with C60 and the pyridoxal linked via a disulﬁde bond. The pyridoxal aldehyde group is bonded through a Schiff base to K51 . Color scheme: Protein secondary structure (green), carbon (white), oxygen (red), nitrogen (blue), sulfur (orange).

14 Protein-based Artiﬁcial Enzymes

K M (2.2- and 7.5-fold) and a small increase in kcat (1.5-fold). UV/Vis spectroscopy, ﬂuorescence and electron-spray mass spectrometry veriﬁed the catalytic function (Schiff base formation) of the introduced lysine residue in the reaction process. Signiﬁcantly, in a study of rate versus pH, IFABP-PX60 gave a rate that decreased monotonically with increasing pH while the lysine mutants exhibited a bell-shaped proﬁle with a maximum rate near pH 7.5. Taken together, these results provide compelling evidence that the lysines participate directly in the reaction and that features of enzymatic processes, including covalent catalysis, can be mimicked successfully. Computer modeling together with mutagenesis was also used to identify existing residues in the protein cavity that contributed to the substrate speciﬁcity of the IFABP-PX60 catalyst. R126 and Y14 were identiﬁed as two important residues that interacted with the γ -carboxylate group of α-keto glutarate when bound to the enzyme via a Schiff base with the PX moiety (Figure 12). Mutants IFABPV60 C,R126 M and -V60 C,R126 M,Y14 F were prepared and pyridoxamine was attached to each mutant [53]. Of particular note, IFABP-PxM126 had a K M three-fold higher than IFABP-PX, while the second mutation at position 14 had no signiﬁcant effect. The kcat s for both conjugates were 2-fold lower than for the original IFABP-PX60. N-methylated pyridoxamine conjugates were also introduced into these two mutants. Compared to the original IFABP-MPX60 conjugate described above, the

Fig. 12 Putative binding site for the sidechain carboxylate of the substrate "-keto glutarate in IFABP-PX60. Present are Y14 (upper left), R126 (upper right) and the Schiff base complex between "-keto glutarate and the pyridoxamine catalyst. (lower

center). Dashed lines indicate putative hydrogen bonds between the (-carboxylate of "-keto glutarate and the side-chain phenol of Y14 and the guanidinium group from R126 . Color scheme: Carbon (white), oxygen (red), nitrogen (blue), sulfur (orange).

3 Catalysts Based on Hollow Lipid-binding Proteins 15

mutations in positions 126 and 14 increased K M as much as seven-fold. IFABPMPxM126 gave a particularly signiﬁcant result, manifesting a 5-fold higher kcat . While neither the experiments with the lysine mutants nor those with the carboxylate binding site mutants produced catalysts that rival the efﬁciency of natural enzymes, incremental improvements were observed with many of the conjugates. Moreover, these advances came from constructs that combined mutations in the protein scaffold with synthetic alterations in cofactor structure, thus highlighting the power of the chemogenetic approach. 3.8 Scaffold Redesign

While site-directed mutagenesis for the incorporation of new functional groups into the protein cavity is the most obvious type of genetic modiﬁcation that can be performed with these systems, more global changes can be made in the scaffold using similar methods. For example, as noted above, the IFABP architecture consists of a β-barrel structure caped off with a α-helical lid. To examine whether entry or exit of substrates or products into or out of this closed cavity was limiting catalytic turnover, a helixless (hs) IFABP mutant was prepared by deleting 17 helical residues (15–31) from the N-terminus and replacing them with a dipeptide linker (Ser-Gly) using site-directed mutagenesis [54]. A previous NMR study showed that this mutant preserved the original β-sheet secondary structure without the α-helical lid and was still relatively stable [55]. The structures of IFABP and hsIFABP are compared in Figure 13; except for the absence of the α-helical lid, the overall protein fold is largely intact. Studies that investigated reactions with this conjugate (hsIFABP-PX60) revealed that in 24 h, approximately two turnovers were observed with a selectivity of 93% ee. A kinetics analysis indicated that kcat s were comparable for both IFABP-PX60 (0.20 h−1 ) and hsIFABP-PX60 (0.22 h−1 ), while the former exhibited a 4-fold increase in K M [56]. After introducing the MPX cofactor into the cavity of helixless protein, 4.9 turnovers in 24 h with 35% ee was observed for the production of Glu under the same conditions. Taken together, these results suggested that removal of the α-helical lid did not affect the maximal rate and enantioselectivity of the respective constructs. However, deletion of this structural element did decrease the substrate binding afﬁnity, as evidenced by an increase in K M . Thus, it appears that while the hsIFABP scaffold was not useful in increasing the efﬁciency of PX-promoted transamination, these experiments did show that stable constructs could be prepared based on the helixless scaffold. This may have greater utility in reactions with larger cofactor catalysts or substrates. 3.9 Hydrolytic Reactions

Fatty acid binding proteins have also been used as scaffolds for constructing hydrolytic catalysts. Phenanthroline was attached to Cys117 of ALBP using

16 Protein-based Artiﬁcial Enzymes

Fig. 13 Comparison of the structures of IFABP and the helixless variant hsIFABP. Top: Stereo view of IFABP. Bottom: Stereo view of hsIFABP.

iodoacetamido 1,10-phenanthroline and metallated to produce a coordinated Cu(II) ion encapsulated within a chiral protein cavity (Figure 14). The resulting conjugate, ALBP-Phen-Cu(II) was able to promote the hydrolysis of several unactivated amino acid esters under mild conditions (pH 6.1, 25 ◦ C) at rates 32- to 280-fold above the background rate in buffered aqueous solution; these reactions also showed modest stereoselectivity [57]. In 24 h incubations, 0.70–7.6 turnovers were obtained with enantiomeric excesses ranging from 31 to 86% ee. ALBPPhen-Cu(II) could also catalyze the hydrolysis of an activated amide (picolinic

Fig. 14 Reaction used to prepare ALBP-Phen.

3 Catalysts Based on Hollow Lipid-binding Proteins 17

acid methyl nitroanilide, PMNA), under slightly more vigorous conditions (37 ◦ C) and after longer incubation times. The rate of amide hydrolysis was 1.6 × 104 fold higher than the background rate [57]. However, kcat obtained with ALBPPhen-Cu(II) was still signiﬁcantly lower than that obtained with a related Cu(II) bipyridine complex [58]. This rate decrease may reﬂect a non-optimal, perhaps nonplanar, conformation for PMNA binding within the ALBP cavity. Of related interest, an X-ray crystal structure of ALBP-Phen was obtained. Inspection of this structure showed that the protein could not accommodate the phenanthroline and PMNA within a planar conformation without signiﬁcant distortion of the protein backbone [45]. This may account for the lower than expected rate of PMNA hydrolysis promoted by ALBP-Phen-Cu(II) compared with non-proteinaceous model complexes. In analogy to the above experiments with pyridoxamine complexes, the 1,10phenan-throline ligand was introduced at several alternative sites (positions 60, 72 and 104, see Figure 8) using the protein scaffold IFABP [59]. Using alanine isopropyl ester as a substrate, IFABP-Phen60 catalyzed ester hydrolysis with less selectivity than ALBP-Phen, while Phen72 promoted the same reaction with higher selectivity. In contrast, hydrolysis of tyrosine methyl ester was catalyzed with higher selectivity by Phen60 and more rapidly by Phen104. These results indicated that the rate enhancement and substrate selectivity of hydrolysis reactions catalyzed by phenanthroline conjugates depended largely on the orientation and environment of the metal ligand within the protein cavity.

3.10 A Flavin-containing Conjugate

Reactions catalyzed by a ﬂavin analog incorporated into IFABP have also been studied by alkylating a Cys residue within the cavity of the helixless variant (hsIFABP) [60]. The conjugate hsIFABP-FL catalyzed the oxidation of several dihydronicotinamides. These experiments were performed mainly to compare results obtained with ﬂavin-IFABP conjugates with similar results acquired with ﬂavopapains; these latter proteins were among the ﬁrst semisynthetic enzymes produced. Interestingly, while hsIFABP-FL and ﬂavopapain gave comparable rate accelerations (kcat /K M ) for dihydronicotinamide oxidation, hsIFABP-FL manifested much higher K M and kcat than did ﬂavopapain. The low K M observed with ﬂavopapain indicates that it primarily accelerates the reaction rate by enhancing substrate binding, whereas the higher kcat obtained with hsIFABP-FL suggests that its major mode of rate acceleration involves enhancing ﬂavin reactivity. Molecular modeling of hsIFABP-FL indicates that Gln27 is close to N(3)H and O(4) of the ﬂavin (Figure 15). Interaction between the carboxamide of Gln27 and the ﬂavin is likely to alter the redox potential of the isoalloxazine and hence augment its reactivity. Taken together, these results with ﬂavopapain and hsIFABP-FL highlight the diverse ways in which protein scaffolds can be used to modulate catalyst activity.

18 Protein-based Artiﬁcial Enzymes

Fig. 15 Hydrogen bonding to the ﬂavin in hsIFABP-FL.

3.11 Some Limitations

While the examples described above highlight the impressive results that can be achieved with protein as scaffolds for catalyst design, this approach is not without problems. Firstly, there is the issue of cavity size. It would be useful to incorporate large metal–ligand systems into this protein system to generate catalysts for a plethora of interesting reactions. Unfortunately, the enclosed cavity of fatty acid binding proteins (and even the somewhat open cavity of hsIFABP) limits the size of what can be introduced into the protein interior. Efforts to incorporate a heme group or a Mn(III)-salen complex into hsIFABP were not successful. While it was possible to attach these moieties to the protein scaffold under denaturing conditions and refold them, the resulting conjugates were unstable and underwent aggregation and precipitation upon storage. In future work, it will be necessary to identify other protein structures that can serve as effective scaffolds but that are not subject to these size limitations. Secondly, work with fatty acid binding proteins is limited to aqueous systems. This creates two types of problems. The ﬁrst is that catalytic systems that are sensitive to water cannot be employed. The Mn(III)salen complex described above underwent Schiff base hydrolysis and subsequent ligand decomposition upon prolonged storage in aqueous buffer. While the rate of this hydrolytic degradation can be decreased by employing mixtures of water and organic solvents, fatty acid binding proteins have limited solubility in such solvents. In addition, the inability to work in such solvents also limits the substrate concentrations that can be used and also precludes the use of these catalysts with many types of hydrophobic substrates. However, given the extensive interest and the large body of literature concerning the use of enzymes in organic media, it should be possible to identify other types of proteins that can serve as scaffolds for catalyst design in such solvent systems.

4 Myoglobin as a Starting Point for Oxidase Design 4.1 Artiﬁcial Metalloproteins and Myoglobin

Metalloproteins represent a major faction of scaffold proteins in artiﬁcial enzyme design. The introduction of a wide range of metals and redox cofactors into protein scaffolds can greatly increase both the diversity of enzyme catalysis and the

4 Myoglobin as a Starting Point for Oxidase Design

application of these artiﬁcial metalloproteins. To sequester artiﬁcial redox cofactors into protein scaffolds, which would otherwise not bind these chemical catalysts, one may either use non-covalent or covalent attachment [61]. Myoglobin is a wellstudied protein that contains a non-covalently bound heme for oxygen transport. The large heme-binding site provides substantial room to accommodate other ligand systems, and for these reasons serves as a useful starting point for catalyst design. 4.2 Non-covalent Attachment of a Redox Center

Watanabe and co-workers pursued an approach involving the non-covalent placement of Mn (III) and Cr (III) salophen complexes into apo-myoglobin [61]. In this artiﬁcial metalloprotein, two residues required mutation to improve the binding afﬁnity of the cofactor and to allow increased access of substrates. The sulfoxidation activity of this Cr(III)-salophen myoglobin complex was studied and showed a sixfold rate increase over free Cr(III)-salophen in solution. The enantioselectivity of free salophen was 0%, while that of the myoglobin catalyst was 13%. While the ee is low for this artiﬁcial metalloprotein, Wantabe and co-workers were successful in demonstrating that asymmetric reactions can be accomplished using chiral protein cavities. 4.3 Dual Anchoring Strategy

To increase the enantioselectivity of these myoglobin metalloenzymes, Lu and coworkers have successfully utilized a covalent linkage approach [62]. In an earlier attempt a Mn(III)-salen complex was incorporated into apo-myoglobin by mutating residue 103 to cysteine, followed by modiﬁcation with a methane thiosulfonate derivative of Mn(III)(salen) (Figure 16). This catalyst showed sulfoxidation activity; however, the ee was only 12%. As such a low ee might be a result of the ability of the bound ligand to exist in multiple conformations within the protein cavity, it was hypothesized that the rotational freedom of the salen complex could be limited if it was anchored at two separate locations, unlike the previous single-point attachment conjugate. The Mn(III)(salen) derivative was modeled into myoglobin to identify two positions that would yield the best ﬁt into the heme pocket (Figure 17). In

Fig. 16 A Mn(III)(salen) derivatizing reagent used to prepare proteinMn(III)(salen) conjugates.

19

20 Protein-based Artiﬁcial Enzymes

Fig. 17 Structure of myoglobin complexed with a heme group. The two residues chosen for mutation, Y103 and L72 , are shown. Left: Side view of myoglobin. Right: Top view of myoglobin. Color scheme: Carbon (white), oxygen (red), nitrogen (blue), iron (pink).

addition to an increase in the rate of sulfoxidation of the substrate thioanisole, the enantioselectivity increased to 51% ee. These results indicate that the dual anchoring strategy may be applied to the design of other artiﬁcial metalloenzymes to obtain better enantioselective control. It may also be possible in future work to modulate substrate speciﬁcity as well. 5 Antibodies as Scaffolds for Catalyst Design 5.1 Antibodies as Speciﬁcity Elements

The immune system of higher eukaryotes produces antibody molecules that recognize a broad range of small molecules. By immunizing animals with speciﬁc antigens, produced by conjugating organic compounds to carrier proteins, antibodies that speciﬁcally recognize the original small molecule can be generated. This allows protein scaffolds that have binding sites tailored for a speciﬁc organic molecule to be produced. Figure 18 shows the hapten binding site of MOPC 315; an impressive number of speciﬁc contacts between the antibody and the nitrophenyl amide containing ligand can be seen. While the immunoglobulin-derived scaffolds are quite large, the ability to create high-afﬁnity substrate binding sites without numerous iterations of design makes this an attractive approach for catalyst design. 5.2 Incorporation of an Imidazole Functional Group into an Antibody for Catalysis

The capability of antibodies to provide tailor-made binding sites for catalyst design has been exploited for the preparation of hydrolytic catalysts. The antibody MOPC 315 binds substituted 2,4-dinitrophenyl-containing compounds with association

5 Antibodies as Scaffolds for Catalyst Design 21

Fig. 18 Stereo view of the structure of the antibody MOPC 315 complexed with an amide-containing hapten. Only the region near the hapten binding site is shown. The hapten and Y34 are shown as larger sticks with the remaining residues in the bind-

ing site as smaller sticks. Residues shown include Y34 , W93 and W98 (from VL ) and W333 , H335 , Y339 , R350 , K359 , Y399 , Y401 and S405 (VH ). Color scheme: Carbon (white), oxygen (red) and nitrogen (blue).

constants ranging from 5 × 104 to 1 × 106 M−1 [63]. Imidazole was incorporated into this antibody via a thiol group introduced by chemically modifying K52H in the active site via a disulﬁde linkage (Figure 19). Using a series of coumarin esters as substrates, multiple (>10) hydrolytic turnovers were observed with no loss of activity. The catalytic efﬁciency, kcat /K M , for this reaction was over 103 times higher than for a model reaction employing 4-methylimidazole.

Fig. 19 Introduction of imidazole into the ligand binding site of MOPC 315 through chemical modiﬁcation and site-directed mutagenesis.

22 Protein-based Artiﬁcial Enzymes

5.3 Comparison of Imidazole-containing Antibodies Produced by Chemical Modiﬁcation and Site-directed Mutagenesis

Since an imidazole group can be incorporated into a protein through site-directed mutagenesis, such an approach was used to prepare a MOPC 315 mutant containing a histidine residue at the corresponding site. In brief, a hybrid Fv fragment of MOPC 315 was constructed by reconstituting a recombinant variable light chain (VL ) produced in E. coli with a variable heavy chain (VH ) from the antibody [64]. Imidazole was introduced into the combining site by substituting Y34 of VL with His using site-directed mutagenesis (see Y34 in Figure 18). This His mutant Fv catalyzed the hydrolysis of the 7-hydroxycoumarin esters of 5-(2,4-dinitrophenyl)aminopentanoic acid 90 000-fold faster than the reaction in the presence of 4-methyl imidazole at pH 7.8. Using that substrate, Fv(Y34HL ) turned over at least eleven times and retained 44% of its activity. The loss of activity probably resulted from the accumulation of the inhibitory reaction product. The mutant Fv bound ε-2, 4-DNP-L-lysine only six-fold less tightly than wild-type protein, suggesting that the substituted Tyr residue is not involved in the DNP recognition process. Compared to the imidazole-catalytic antibody generated by tethering an imidazole chemically, a sixteen-fold greater rate increase was observed for the reactions catalyzed by His mutant Fv protein under similar conditions. This rate enhancement is likely due to the fewer degrees of the freedom of the imidazole moiety inside the combining site (cf. 19-1 and 19-2). While these results alone indicate that the construct prepared by mutagenesis functioned more efﬁciently than the one obtained from chemical modiﬁcation, this particular example does not exemplify the most powerful feature of chemical modiﬁcation, i.e., the ability to incorporate non-natural functionality or spacers into complex protein structures. Considerable use will probably be made of this strategy in future catalyst design.

6 Conclusions

The examples described in this chapter reveal the breadth of methods now used to create protein-based catalysts. Structural analysis via X-ray and NMR techniques has proved to be critical for providing atomic structures that serve as the starting point for design. Computer modeling has become a powerful method in both the planning of new designs and in the interpretation of experimental results. Recombinant DNA techniques allow chemists to make mutations or more global changes in protein scaffolds. Together, these tools have allowed researchers to assemble new protein-based catalysts that can be used to probe biological systems or catalyze useful chemical transformations. While these catalysts do not in general rival the efﬁciency of natural enzymes, impressive rate accelerations have been obtained in some cases. New methods including phage display [65] and antibody production promise to provide greater access to ligand-speciﬁc scaffold development.

References

Peptide/protein ligation techniques offer the possibility of increasing the scope of functionality that can be incorporated into protein structures [66]. As the understanding of how enzymes promote chemical reactions with high efﬁciency increases, so too will the ability to design more efﬁcient protein-based catalysts. Given the enormous power obtained from combining chemical and genetic methods it is likely that the ﬁeld of protein-based artiﬁcial enzyme design will continue to grow in the near future.

References 1 K. E. NEET, D. E. KOSHLAND, Jr., Proc. Natl. Acad, Sci. U.S.A. 1966, 56, 1606–1611. 2 L. POLGAR, M. L. BENDER, J. Am. Chem. Soc. 1966, 88, 3153–3154. 3 Z. P. WU, D. HILVERT, J. Am. Chem. Soc. 1989, 111, 4513–4514. 4 H. L. LEVINE, Y. NAKAGAWA, E. T. KAISER, Biochem. Biophys. Res. Commun. 1977, 76, 64–70. 5 D. R. COREY, P. G. SCHULTZ, Science 1987, 238, 1401–1403. 6 P. BERGLUND, M. R. STABILE, M. GOLD, J. B. JONES, Bioorg. Med. Chem. Lett. 1996, 6, 2507–2512. 7 P. BERGLUND, G. DESANTIS, M. R. STABILE, X. SHANG, M. GOLD, R. R. BOTT, T. P. GRAYCAR, T. H. LAU, C. MITCHINSON, J. B. JONES, J. Am. Chem. Soc. 1997, 119, 5265–5266. 8 E. T. KAISER, D. S. LAWRENCE, Science 1984, 226, 505–511. 9 E. T. KAISER, D. S. LAWRENCE, S. E. ROKITA, Annu. Rev. Biochem. 1985, 54, 565–595. 10 C. F. MEARES, T. G. WENSEL, Acc. Chem. Res. 1984, 17, 202–209. 11 D. S. SIGMAN, T. W. BRUICE, A. MAZUMDER, C. L. SUTTON, Acc. Chem. Res. 1993, 26, 98–104. 12 I. M. BELL, D. HILVERT, Perspect. Supramol. Chem. 1994, 1, 73–88. 13 M. D. DISTEFANO, H. KUANG, D. QI, A. MAZHARY, Curr. Opin. Struct. Biol. 1998, 8, 459–465. 14 J. B. JONES, G. DESANTIS, Acc. Chem. Res. 1999, 32, 99–107. 15 D. HARING, P. SCHREIER, Naturwissenschaften 1999, 86, 307–312. 16 G. DESANTIS, J. B. JONES, Curr. Opin. Biotecnol. 1999, 10, 324–330.

17 C.-M. TANN, D. QI, M. D. DISTEFANO, Curr. Opin. Chem. Biol. 2001, 5, 696–704. 18 R. BRESLOW, S. D. DONG, Chem. Rev. 1998, 98, 1997–2011. 19 D. QI, C. M. TANN, D. HARING, M. DISTEFANO, Chem. Rev. 2001, 101, 3081–3111. 20 M. A. ZENKOVA 2004, Artiﬁcial Nucleases, Springer, Germany. 21 C. H. CHEN, D. S. SIGMAN, Science 1987, 237, 1197–1201. 22 M. M. MEIJLER, I. ZELENKO, D. S. SIGMAN, J. Am. Chem. Soc. 1997, 119, 1135–1136. 23 T. E. GOYNE, D. S. SIGMAN, J. Am. Chem. Soc. 1987, 109, 2846–2848. 24 R. JUE, J. M. LAMBERT, L. R. PIERCE, R. R. TRAUT, Biochemistry 1978, 17, 5399– 5406. 25 D. N. ARVIDSON, C. BRUCE, R. P. GUNSALUS, J. Biol. Chem. 1986, 261, 238–243. 26 T. W. BRUICE, J. G. WISE, D. S. E. ROSSER, D. S. SIGMAN, J. Am. Chem. Soc. 1991, 113, 5446–5447. 27 D. H. OHLENDORF, W. F. ANDERSON, R. G. FISHER, Y. TAKEDA, B. W. MATTHEWS, Nature 1982, 298, 718–723. 28 R. G. BRENNAN, Y. TAKEDA, J. KIM, W. F. ANDERSON, B. W. MATTHEWS, J. Mol. Biol. 1986, 188, 115–118. 29 R. G. BRENNAN, S. L. RODERICK, Y. TAKEDA, B. M. MATTHEWS, Proc. Natl. Acad. Sci. U.S.A., 1990, 87, 8165–8169. 30 P. LEIGHTON, P. LU, Biochemistry 1987, 26, 7262–7271. 31 G. XIAO, D. L. COLE, R. P. GUNSALUS, D. S. SIGMAN, Protein Sci. 2002, 11, 2427–2436. 32 C. Q. PAN, J. A. FENG, S. E. FINKEL, R. LANDGRAF, D. SIGMAN, R. C. JOHNSON,

23

24 Protein-based Artiﬁcial Enzymes

33

34

35

36

37 38 39

40 41 42

43

44

45

46

47

Proc. Natl. Acad. Sci. U.S.A. 1994, 91, 1721–1725. D. KOSTREWA, J. GRANZIN, C. KOCH, H. W. CHOE, S. RAGHUNATHAN, W. WOLF, J. LABAHN, R. KAHMANN, W. SAENGER, Nature 1991, 349, 178–180. H. S. YUAN, S. E. FINKEL, J. A. FENG, M. KACZOR-GRZESKOWIAK, R. C. JOHNSON, R. E. DICKERSON, Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 9558–9562. R. H. EBRIGHT, Y. W. EBRIGHT, P. S. PENDERGRAST, A. GUNASEKERA, Proc. Natl. Acad. Sci. U.S.A. 1990, 87, 2882–2886. R. H. EBRIGHT, Y. W. EBRIGHT, A. GUNASEKERA, Nucl. Acids Res. 1989, 17, 10295–10305. P. S. PENDERGRAST, Y. W. EBRIGHT, R. H. EBRIGHT, Science 1994, 265, 959–962. S. C. SCHULTZ, G. C. SHIELDS, T. A. STEITZ, Science 253, 1001–1007. J. A. BOWN, J. T. OWENS, C. F. MEARES, N. FUJITA, A. ISHIHAMA, S. J. W. BUSBY, S. D. MINCHIN, J. Biol. Chem. 1999, 274, 2263–2270. T. D. TULLINS, B. A. DOMBROSKI, Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 5469–5473. C. LIU, M. WANG, T. ZHANG, H. SUN, Coord. Chem. Rev. 2004, 248, 147–168. L. BANASZAK, N. WINTER, Z. XU, D. A. BERNLOHR, S. COWAN, T. A. JONER, Adv. Protein Chem. 1994, 45, 89–151. M. E. HODSDON, J. W. PONDER, D. P. CISTOLA, J. Mol. Biol. 1996, 264, 585–602. H. KUANG, M. L. BROWN, R. R. DAVIES, E. C. YOUNG, M. D. DISTEFANO, J. Am. Chem. Soc. 1996, 118, 10702–10706. J. J. ORY, A. MAZHARY, H. KUANG, R. R. DAVIES, M. D. DISTEFANO, L. J. BANASZAK, Protein Eng. 1998, 11, 253–261. H. KUANG, R. R. DAVIES, M. D. DISTEFANO, Bioorg. Med. Chem. Lett. 1997, 7, 2055–2060. H. KUANG, M. D. DISTEFANO, Protein Eng. 1997, 10, 25.

48 H. KUANG, M. D. DISTEFANO, J. Am. Chem. Soc. 1998, 120, 1072–1073. 49 D. QI, H. KUANG, M. D. DISTEFANO, Bioorg. Med. Chem. Lett. 1998, 8, 875–880. 50 H. KUANG, D. HARING, D. QI, A. MAZHARY, M. D. DISTEFANO, Bioorg. Med. Chem. Lett. 2000, 10, 2091–2095. 51 D. HARING, H. KUANG, L. J. BANASZAK, M. D. DISTEFANO, Protein Eng. 2002, 15, 603–610. 52 D. HARING, M. D. DISTEFANO, Bioconj. Chem. 2001, 12, 385–390. 53 D. HARING, M. D. DISTEFANO, Bioorg. Med. Chem. 2001, 9, 2461–2466. 54 K. KIM, D. P. CISTOLA, C. FRIEDEN, Biochemistry 1996, 35, 7553–7558. 55 R. A. STEELE, D. A. EMMERT, J. KAO, M. E. HODSDON, C. FRIEDEN, D. P. CISTOLA, Protein Sci. 1998, 7, 1332–1339. 56 D. QI 2003, Chemistry PhD Thesis, University of Minnesota, 140. 57 R. R. DAVIES, M. D. DISTEFANO, J. Am. Chem.Soc. 1997, 119, 11643–11652. 58 L. M. SAYRE, K. V. REDDY, A. R. JACOBSON, W. TANG, Inorg. Chem. 1992, 31, 935–937. 59 R. R. DAVIES, H. KUANG, D. QI, A. MAZHARY, E. MAYAAN, M. D. DISTEFANO, Bioorg. Med. Chem. Lett. 1999, 9, 79–84. 60 C-M. TANN 2003, Chemistry PhD Thesis, University of Minnesota, 201. 61 M. OHASHI, T. KOSHIYAMA, T. UENO, M. YANASE, H. FUJJI, Y. WATANABE, Angew. Chem. Int. Ed. 2003, 42, 1005–1008. 62 J. R. CAREY, S. K. MA, T. D. PFISTER, D. K. GARNER, H. K. KIM, J. A. ABRAMITE, Z. WANG, Z. GUO, Y. LU, J. Am. Chem. Soc. 2004, 126, 10812–10813. 63 S. J. POLLACK, P. G. SCHULTZ, J. Am. Chem. Soc. 1989, 111, 1929–1931. 64 E. BALDWIN, P. G. SCHULTZ, Science 1989, 245, 1104–1107. 65 R. H. HOESS, Chem. Rev. 2001, 101, 3205–3218. 66 P. E. DAWSON, T. W. MUIR, I. CLARK-LEWIS, S. B. H. KENT, Science 1994, 266, 776–779.

1

Force Generation in Molecular Motors George Oster University of California, Berkeley, USA

Hongyun Wang University of California, Santa Cruz, USA

Originally published in: Molecular Motors. Edited by Manfred Schliwa. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-52730594-0

1 Introduction

Imagine living in a world where a Richter 9 earthquake raged continuously. In such an environment, engines would be unnecessary. You would not need to even pedal your bicycle: you would simply attach a ratchet to the wheel preventing it from going backwards and shake yourself forwards! At the scale of proteins, Brownian motion is even more furious, and proteins evolved to take advantage of this enormous supply of energy. Feynman showed that the familiar mechanical ratchet could not work in an isothermal environment, lest it violate the Second Law of Thermodynamics [1], and so motor proteins must employ a different strategy to convert random thermal fluctuations into a directed force: they use chemical energy via intermolecular forces to capture ‘favorable’ configurations. The way in which proteins do this is dictated by three factors: their size, the strength and range of intermolecular bonds at physiological conditions and the magnitude of the Brownian fluctuations that constitute their thermal environment. These determine the energy, length and time scales on which protein motors can operate. Roughly speaking, motor proteins trap thermal fluctuations in two ways: by biasing diffusion of small, angstrom-sized, steps (‘small ratchets’), and by rectifying nanometer-sized or larger, diffusional displacements (‘big ratchets’). For reasons that will become clear, biasing a sequence of small Brownian fluctuations is generally called a power stroke, while rectifying a large thermal displacement is called a Brownian ratchet. Said another way, Brownian ratchets move down free energy landscapes in steps much larger than kB T, while power strokes move in steps

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Force Generation in Molecular Motors

comparable to or smaller than kB T1) . The distinction is imprecise, but useful, since it delineates two extremes in the general mechanism by which proteins use intermo-lecular attractions to convert chemical energy into mechanical work.2) There are only a few motors that can be regarded as being pure power stroke motors or pure ratchets; most protein motors employ a combination of the two strategies. However, these ‘thoroughbreds’ are good illustrations of the principles. In fact, evolution has designed one protein that joins both ratchet and power stroke motors into one remarkable device: F 0 F 1 ATPase, or ATP synthase, the machine that manufactures the fuel that powers many other protein machines. We will use this protein as our running example. Our discussion will be mostly qualitative and heuristic. However, it is important to realize that the explanatory cartoons we use are supported by extensive calculations. Omitting them is akin to leaving out the ‘Materials and Methods’ section in an experimental paper: assertions without authority are simply opinions. The supporting quantitative arguments are, perforce, contained in the citations.

2 A Brief Description of ATP Synthase Structure

In order to discuss the workings of the F0 and F1 motors we give a brief account of their structure, which is summarized in Fig. 1. ATP synthase comprises two rotary motors acting in opposition, each operating on an entirely different physical mechanism. The F0 motor is contained in the membrane-bound portion and employs as its energy source a transmembrane electromotive force. The F1 motor is contained in the soluble portion and is driven by the hydrolysis of ATP. The F1 motor is constructed of a coiled-coil shaft (the γ -subunit) surrounded by a hexamer composed of alternating α and β subunits. Nucleotide binding sites nestle in the cleft between subunits; however, only three of the sites are catalytically active, the other three bind, but do not hydrolyze, ATP. The catalytic sites are contained mostly in the β-subunit, with a few—but crucial—residues contributed by the α subunit. The catalytic sites hydrolyze sequentially and drive the rotation of the γ shaft. Movies showing the motion of the hexamer and the rotation of the γ -shaft can be downloaded from a web site given in [1, 2]. The F0 motor is composed of two portions. Between 10 and 14 c-subunits (depending on the species and/or conditions) are assembled into a cylinder attached to the γ -shaft and ε-subunit. It interfaces with a second transmembrane assembly consisting of the a and b subunits. By convention, the γ -cn -ε assemblage is called the ‘rotor’, and the remainder of the protein (the α 3 β 3 hexamer and the δ-b2 -a complex) comprises the ‘stator’, although each rotates in the opposite direction. 1) The quantity kB T measures the thermal energy of Brownian motion; its value is 1 kB T ∼ 4.1 pN nm = 4.1 × 10−21 J at 298 K (25 ◦ C).

2) We have included a short Appendix with several simple examples that illustrate the difference between a ‘power stroke’ and ‘Brownian ratchet’.

3 The F1 Motor: A Power Stroke 3

Fig. 1 The structure of ATP synthase. The panel on the left shows a composite based on the pdb coordinates of the known subunits [23]; the right-hand panel is the corresponding structure represented schemat-

ically showing the relative rotation of the F0 and F1 motors and the direction of ion flow through the a-cn subunit interface. Subscripts denote the subunit stoichiometries.

3 The F1 Motor: A Power Stroke

The study of protein ATPases has led to a few generalizations that help us understand the mechanism by which these molecular motors convert the energy residing in the nucleotide γ -phosphate bond into a directed mechanical force. Ĺ At physiological conditions, the free energy of hydrolysis of one ATP is ∼ 20–24 kB T; of this, about 8–9 kB T is enthalpic, the balance being entropic. Ĺ Almost all nucleotide binding sites are nestled in the cleft between protein subunits. The nucleotide is grasped by loops emanating from a parallel β-sheet. Ĺ In many motors, the force-generating step is associated with the binding of nucleotide to the catalytic site. We propose that this is true of all ATPase motors. In particular, for the F1 motor this is the only way to reconcile all of the biochemical and mechanical measurements with its high mechanical efficiency. Ĺ After the force has been generated by ATP binding, an ATP is tightly bound in the catalytic site. The role of hydrolysis is to break the tightly bound ATP into two products and weaken the binding so the products can be released and the force-generating cycle can be repeated.

In the F1 motor, the ATP binding site lies asymmetrically in the cleft between the β and α subunits, the majority of the catalytic sites residing in the β subunit. The power stroke is accomplished by a hinge bending motion that swings the top of the β subunit down towards the bottom portion. The bending motion of the β subunit can be measured by the motion of helices B and C that emanate from the β sheet of

4 Force Generation in Molecular Motors

Fig. 2 The F1 power strokes are accomplished by the hinge bending motions of $ subunits, which are driven by ATP binding to the catalytic sites. (a) During the hinge bending motion, the top part of $ rotates about 30◦ toward the bottom part. This rotation closes the angle between helices B

and C. (b) The hinge bending motion of each $ subunit pushes on the off-axis section of the ( shaft. The rotation of the y shaft is driven by the coordinated hinge bending motions of all three $ subunits [1, 2, 5].

the catalytic site, as shown in the ribbon diagram in Fig. 2a. The bending of the (3 subunit by about 30◦ rotates the central γ shaft by pushing on its off-axis section, much like turning a crankshaft (Fig. 2b). The energy for the power stroke derives from the nucleotide hydrolysis cycle, which consists of four major steps: Binding

Hydrolysis

1

2 ADP Release

CS + ATP CS • ATP PI Release

3

CS • ADP + PI

CS • ADP • PI

CS + ADP

4

(1)

The nucleotide binding step 1 is the force-generating step and should more properly be expressed as a sequence of binding steps: CS + ATP CS • ATP . . . CS • ATP

(2)

Binding Transition

Here the progression from weak to tight binding is symbolized by the increasing size of the bonding symbol indicating the progressive annealing of 15–20 hydrogen bonds (and some hydrophobic interactions at the sugar end). The mechanism driving the hinge bending motion of the β is illustrated schematically in Fig. 3. Immediately after it diffuses into an open catalytic site, the ATP is bound only weakly. The catalytic site wraps around the ATP by forming more bonds which tightens the catalytic site and pulls down the top part of β toward the bottom part. The bonds between the ATP and the catalytic site are formed more or less sequentially, the formation of each bond corresponding to a small drop in binding

3 The F1 Motor: A Power Stroke 5

Fig. 3 The ATP binding transition from weak to tight proceeds as the catalytic site grasps the ATP in a grip of hydrogen bonds (the ‘binding zipper’). As binding progresses, the catalytic site closes up and pulls the top part of the $ subunit toward the bottom part. In this way, the binding free energy is converted to a power stroke

with nearly a constant force. During the power stroke some of the binding energy is stored in the elastic deformation of the p-sheet, which acts ike a spring. This energy is released during the unbending motion to aid product release and return the subunit to its open state.

free energy, that drives a small fraction of the hinge bending motion [1, 3, 4]. When all the bonds have been formed, the ATP is tightly bound in the catalytic site. The overall process from weak to tight binding is called the ‘binding transition’. During the binding transition, the ATP binding free energy is utilized efficiently to generate a bending motion with a nearly constant torque about the hinge region near the β-sheet. The binding transition has two important features [1, 4, 5]: Ĺ The binding free energy decreases (the binding becomes stronger) nearly monotonically and smoothly during the binding transition. This drives the hinge bending motion of the β subunit and consequently the γ shaft rotation in the hydrolysis cycle. In the reverse synthesis cycle, the unbending of the β subunit is driven by the γ shaft rotation in the opposite direction, powered by the F0 motor. When the top of the β subunit is forced up and away from the bottom portion, the

6 Force Generation in Molecular Motors

binding free energy increases (the binding becomes weaker). This progressively releases the nucleotide from the catalytic site [6]. Ĺ By the end of the binding transition, approximately 6–10 kB T of elastic energy is stored in the β-sheet whose loops grasp the nucleotide. Note that the catalytic site should be flexible but not elastic, lest it dissipate too much of the binding free energy by elastic recoil. To summarize, the power stroke is driven by progressive capturing of relatively small Brownian motions that anneal the nucleotide into the catalytic site. Product release is accomplished by using the free energy of hydrolysis to weaken the product binding so that thermal fluctuations can knock them out of the catalytic site. Thus Brownian motion drives both the power and exhaust strokes.

4 The F0 Motor: A Brownian Ratchet

The F0 motor is driven by the ion-motive force, µc across a bacterial or mitochondrial inner membrane. The ion-motive force consists of an ion concentration gradient and an electrical potential difference. In most organisms, the ion is a proton. However, much information has been gleaned from anaerobic bacteria whose F0 is driven by a sodium ion motive force. In either case, the transmembrane chemical potential difference in millivolts is given by: µc + = 2.3

kB T e

log10 [c + ] p − log10 [c + ]c +ψ

[mV ]

(3)

pH

where e is the electronic charge, [c+ ]p the ion (sodium or proton) concentration of the periplasm expressed in molars, [c+ ]c the ion concentration of the cytoplasm and ψ is the electrical potential difference across the membrane. Equation 3 is a thermodynamic relationship that implies that a concentration difference is equivalent to an electrical potential at equilibrium. Since motors operate out of equilibrium, this turns out not to be true in general. However, Eq. 3 points out the need for a mechanism for transforming both an entropic and electrical potential into mechanical work. The basic principle that accomplishes this can be illustrated by the ‘toy’ model shown in Fig. 4. 4.1 A Pure Brownian Ratchet

First, consider a rod with a linear array of negative charges (e. g. a DNA strand, etc.) that can freely diffuse through a hydrophilic pore embedded in a membrane as shown in Fig. 4a. If a potential, , is imposed across the membrane, then the rod will be propelled to the left and can perform work against a load force, FL . This can

4 The F0 Motor: A Brownian Ratchet

Fig. 4 Principle of the F0 motor. (a) A pure power stroke. A rod of negative electrica charges passes through a polar transmembrane pore. An electrical potential, imposed across the membrane drives the line of charges to the left. (b) A pure Brownian ratchet. The charged rod passes through a hydrophobic membrane pore. A high concentration of counterion charges (Na+ or H+ ) on the right can bind to the negative sites and neutralize them so that they can pass through the pore. A second positive ion on the left (not shown) neutralizes the membrane potential. The low concentration of counterions on the left ensures that

the bound charges dissociate but do not quickly rebind, so that the bare site cannot re-enter the pore. The attractive bonds between the rod charge and the aqueous solvent molecules rectify the rod’s diffusion. We call this a Brownian ratchet. (c) The power stroke and ratchet can be combined using an L-shaped ‘stator’. An aqueous input channel (colored white) connects via a polar channel (to the right) to the output reservoir The body of the stator is hydrophobic so that ions cannot leak across the membrane. With this design, the motor has both power stroke and ratchet components.

7

8 Force Generation in Molecular Motors

be considered as nearly a pure power stroke. The motion of the rod can be viewed as being driven by a driving potential, φ, tilted to the left in Fig. 4a. The slope of the driving potential gives the motor force driving the rod to the left: FM = −dφ/dx. The rod will stall when the motor force equals the load force: FM = FL . Because the rod is very small, its motion is stochastic (indicated in Fig. 4 by the Gaussian force labeled kB T). In fact, the net drift of the rod driven by the potential is deeply buried in its random motion: at any time the rod is only a tiny bit more likely to move to the left than to the right, and the mean square of the instantaneous velocity is several orders of magnitude larger than the net drift velocity. So the net movement down the potential gradient can only be detected by looking at correlations over many steps. This is illustrated by example 3 in the Appendix. 4.2 A Pure Power Stroke

Next, consider the situation in Fig. 4b where the rod passes through a hydrophobic pore. Moreover, there is a difference in the concentration of a positive counterion (e. g. sodium or protons) between the two sides of the membrane. The counterion can bind to and neutralize the negative charges (binding sites) on the rod. A difference in the concentration of a second ion (e.g. potassium), which cannot bind to the sites on the rod, neutralizes the membrane potential. A bare binding site is negatively charged and cannot move into the hydrophobic pore, for that would entail shedding its hydration shell at a considerable energetic cost.3) Because of the high concentration on the right side, the binding sites will be largely neutralized and so the rod can diffuse freely to the left through the pore. However, once a neutralized site has emerged from the pore to the left, the bound positive charge will quickly dissociate and leave the binding site unoccupied. Because of the low concentration on the left side, the binding site is likely to remain unoccupied, which prevents it from moving back into the pore. Thus diffusion to the right is rectified by the ion concentration difference, and the rod moves stochastically to the left, driven by a pure Brownian ratchet. The driving potential, φ, in this case looks like a staircase. When the concentration on the left is much lower than that on the right, each step of the staircase potential is much larger than kB T, so that reverse steps are unlikely. This is illustrated by example 4 in the Appendix. The load force, FL , opposing the motion has the effect of ‘tilting’ the driving potential so that the rod must diffuse uphill, against the load force. The two driving forces can be combined as shown in Fig. 4c. Here the membrane is horizontal and so the motor must be augmented by a fixed ‘stator’ assembly. The stator body is hydrophobic with two exceptions. First, there is a large aqueous channel that permits the ions from the high concentration reservoir to access the binding sites. Second, there is a polar channel connecting the aqueous channel 3) The energy cost of moving a negative charge into the pore is approximately G ∼ 45 kB T [22].

4 The F0 Motor: A Brownian Ratchet

Fig. 5 Operation of the F0 motor [7] (a) Simplified geometry of the sodium driven F0 motor showing the path of ions through the stator. This is the same arrangement as in Fig. 4c, but with the charge array wrapped around a cylinder that is free to rotate in the plane of the membrane with respect to the stator. In addition, a ‘blocking charge’ has been added to the stator to prevent the leakage of charge from the high concentration reservoir (periplasm) to the low concentration reservoir (cytoplasm). (b) Free energy diagram of one rotor site as it passes through the rotor–stator interface. Step 1 → 2, the rotor diffuses to the left, bringing the empty (negatively charged) site into the attractive field of the positive stator charge. 2 → 3, once the site is captured,

the membrane potential biases the thermal escape of the site to the left (by tilting the potential and lowering the left edge). 3 → 4, the site quickly picks up an ion from the input channel neutralizing the rotor. 4 → 5, an occupied site being nearly electrically neutral can pass through the dielectric barrier. If the occupied site diffuses to the right, the ion quickly dissociates back into the input channel as it approaches the stator charge. 5 → 6, upon exiting the stator the site quickly loses its ion. The empty (charged) site binds solvent and cannot pass backwards into the low dielectric of the stator. The cycle decreases the free energy of the system by an amount equal to the electromotive force.

to the low concentration reservoir. Since the aqueous channel is isopotential with the high concentration reservoir, this arrangement converts the transmembrane potential drop from vertical to horizontal. In this way the membrane potential and the ion concentration difference act in tandem to move the rod to the left, the former driving a power stroke and the latter driving a Brownian ratchet. The actual F0 motor works somewhat differently from the idealized version in Fig. 4c. Fig. 5a shows two more modifications that are required to make the arrangement resemble the sodium driven F0 motor of the bacterium P. modestum [7, 8]. The linear array of charged binding sites is first wrapped around a cylinder. This cylinder consists of 10–14 c-subunits, depending on the organisms and/or conditions. Second, a ‘blocking charge’ (R232) is present on the stator that prevents leakage of ions between the two reservoirs that would dissipate the ion gradient unproductively. The presence of this blocking charge alters the picture of the rotation of the rotor with respect to the stator substantially. Fig. 5b shows the potential experienced by one binding site of the F0 motor. The presence of the blocking charge creates an electrostatic potential well that attracts the rotor binding site as soon as it diffuses into the polar channel. Once trapped in the well, the rotor

9

10 Force Generation in Molecular Motors

depends on Brownian fluctuations to escape. However, the membrane potential biases escape by lowering the left side of the electrostatic well so that the rotor charge is much more likely to escape to the left than to the right. Once it escapes into the aqueous input channel, it is quickly neutralized by the positive ions so that it can move freely across the hydrophobic barrier. Note that the membrane potential only biases the Brownian fluctuations to the left, but does not actually drive a power stroke as in Fig. 4a. In summary, the F0 motor qualifies as a Brownian ratchet since it rectifies large thermal fluctuations (or long diffusions). The energy for rectification derives from the ion concentration gradient via the short range interactions between ions and the binding sites (binding and unbinding). It also uses the membrane potential to bias thermal fluctuations that release the rotor site from the attraction of the stator blocking charge: it takes less energy to hop out to the left than to the right. This might be thought of as a partial power stroke, so we see that the classification into power stroke and ratchet is largely a question of definition. An interesting class of ratchet motors is those that use the principle of trapping Brownian fluctuations during their assembly to perform a ‘1-shot’ motor task. Examples include the acrosome of Limulus and Thyone, the spasmoneme of Vorticella [9], and the polymerizaton of actin that propels la-mellipodial protrusion and certain intracellular parasites, such as Listeria and Shigella [10, 11]. An important corollary of the ratchet principle, and one that dramatically distinguishes molecular motors from other machines, is that using thermal fluctuations to go uphill in free energy amounts cools off the immediate environment. If the ratchet shown in Fig. 4b is coupled to work against a conservative load force, then the process is endothermic: heat is absorbed from surrounding fluid to increase the potential energy of the external agent that exerts the load force. By contrast, if the motor shown in Fig. 4a is coupled to work against a viscous load, then the process is exothermic: energy from the electrical potential goes to produce the drift velocity of the rod, which in turn is converted to heat by viscous friction. These microscopic thermal energy transactions lead to some surprising properties of molecular motors. For example, it is possible for the motor to perform more work on a viscous load than the free energy it derives from a reaction cycle! These counterintuitive properties are discussed in more detail in the references [5, 12, 13].

5 Coupling and Coordination of Motors

Most ATPase motors have more than one catalytic site. During the motor operation, each catalytic site hydrolyzes ATP and contributes to force generation. These catalytic sites generally do not operate independently, but act in concert with other catalytic sites. Two heads of a kinesin dimer are coordinated with each other to generate unidirectional motion and to ensure processivity. ATP synthase has three catalytic sites. AAA motors are hexamers of ATPases, sometimes stacks of two hexamers. The chaperonin Groel is a stacked pair of heptameric rings, each with seven

5 Coupling and Coordination of Motors

catalytic sites and the portal protein may even be a dodecameric ring of 12 ATPases. Generally, the catalytic sites must act in concert, either firing sequentially, as in F1 , or as in Groel, simultaneously in each ring, but alternating between the rings. This coordination is necessary for the proper operation of the motors, but how is it accomplished? In all cases, motor ATPase catalytic sites are too far apart to communicate in any other way than via elastic strain through the intervening protein structure. Although the details of strain coordination differ, a clue can be found in the correlation between the ADP release at one catalytic site and the ATP binding at another. In F1 , the strain-induced release of ADP arises from two possible sources. First, the γ shaft is asymmetric, so that at every rotational position it strains each catalytic site differently. Second, the intrinsic asymmetry of the protein structure allows the catalytic site to radiate strain differently to the two neighbor sites. The catalytic sites are located in the cleft between adjacent α and β subunits, with the majority of the binding residues in the β subunit [14, 15] This asymmetry allows ATP binding at one catalytic site to propagate a conformational change to the β portion of next catalytic site in the direction of motor rotation, but propagates a different conformational change to the β portion of the previous catalytic site. Thus, one catalytic site can affect the reactions on two neighboring catalytic sites differently. The conformational change directed to the β portion of the next catalytic site can lower the free energy barrier for ADP dissociation, readying it for the next hydrolysis cycle. Indeed this may be the primary structural feature determining the direction of rotation. Because of the unequal symmetry between the F0 and F1 motors, elastic coupling plays an additional role. The F1 motor has three catalytic sites and rotates in three major 120◦ steps, each with a brief pause at 90◦ . On the other hand, the F0 motor has a rotational symmetry that varies between 10 and 14, depending on the source. Therefore, there is no unique ‘stoichiometry’ between the two rotational motions. This symmetry mismatch is not a problem for the motors since the γ -shaft that couples them is torsionally flexible. In synthesis, this allows the stochastic F0 motor to deliver a smooth torque to F1 , which minimizes dissipation as the nucleotide is unzipped from the catalytic site [1, 16]. Thus elastic coupling between subunits of a protein motor provides the means for both coordinating the catalytic cycles and buffering the independent Brownian motions of the subunits. In walking motors the determination of directionality depends on asymmetric structural features of the heads that alternate depending on whether the head is leading or trailing. However, the situation is more complicated since each head has two binding partners: nucleotide and track. One proposal is that strain is generated by binding of the forward head to the track, triggers release of product from that head [17]. Also strain is relayed to the rear head via the connecting structure to lower the energy barrier for a particular step in the reaction cycle (for example, hydrolysis, or product release). This particular reaction step, in turn, triggers the release of rear head from the track [18]. Thus, binding to and release from the track are correlated with the relative positions of the heads. The alternating phases of the two hydrolysis cycles generate unidirectional motion [19].

11

12 Force Generation in Molecular Motors

6 Measures of Efficiency

We have discussed the molecular principles by which protein motors convert chemical bond energy into mechanical work. However, while the general principles may be the same for all motors, the detailed mechanisms are quite diverse. Therefore, it is frequently useful to determine the efficiency of a motor to provide clues as to its mechanism. The most common mechanical measurement performed on protein motors is to vary the load (the force resisting the motion) and measure the speed. In general, two kinds of load experiments are carried out on protein motors in order to determine their load–velocity behavior. In one type, a load is applied to resist the motor’s progress using a laser trap or the elastic stylus of an atomic force microscope. In this case the motor is working against a conservative force (i.e. derivable from a potential energy function) that depends only on the displacement of the motor, f = −∂φ/∂ξ ). A second, and generally more experimentally convenient method, is to vary the viscous resistance of the fluid environment of the motor. This is a dissipative force that depends on the motor velocity. The information gleaned from the two kinds of measurements yield different information [1, 12, 13]. The thermodynamic efficiency, ηTD is generally defined as the ratio of the work done by the motor to the energy input: ηT D =

f ·δ −G

(4)

where G is the free energy drop in one reaction cycle (e.g. from hydrolyzing one ATP, or passing one proton through the motor), and f ·δ. is the reversible work done by the motor against the conservative load force, f , in one reaction cycle. Here δ is the average distance covered per reaction cycle, sometimes called the ‘step size’. f ·δ is the energy output from the motor because it goes to increasing the potential energy of the external agent that exerts the conservative force. Thus, the thermodynamic efficiency is the ratio of energy output to energy input and it measures the energy conversion efficiency when the motor is operating reversibly i.e. ‘infinitely slowly’. For a motor working against a constant force (e.g. a laser trap force clamp), Eq. (4) can be generalized to the steady state [1, 13] by defining an efficiency: η≡

f ·v −G·r

(5)

Here r is the average reaction rate (e.g. hydrolysis cycles per second or average proton flux) and (v) is the average velocity. Strictly speaking, Eq. (5) is not a thermodynamic quantity since the steady state need not be the equilibrium state. Nevertheless, it is a well-defined quantity that is less than unity.

6 Measures of Efficiency

In general, the average step size, δ depends on the load force, f . When δ is independent of f we say the motor is tightly coupled. In that case, each reaction cycle is, on average, coupled to a fixed displacement (one motor step) regardless of the load force. A tightly coupled motor has two properties: Ĺ When the motor is stalled the chemical reaction is also stopped. Ĺ Increasing the load beyond the stall force drives the motor in the opposite direction and also reverses the chemical reaction.

For a tightly coupled motor, one can show that the stall force is given by f stall = −G/δ. At stall, the thermodynamic efficiency is 100%. When the motor is operating close to stall, the thermodynamic efficiency approaches 100%, regardless of the motor mechanism. Conversely, a high thermodynamic efficiency suggests only that the motor motion and the chemical reaction are tightly coupled [12]. Equation (5) applies only to the situation where the motor is working against a constant load. For macroscopic motors, inertia is important and the effect of Brownian fluctuations is negligible. Therefore, they tend to move at roughly a constant velocity (at least on short time scales), and so the friction force on the motor is approximately constant. In this case, the friction force can be treated effectively as a conservative force and the efficiency given by Eq. (5) is well defined. Thus, for a macroscopic motor, we generally do not have to worry whether it is working against a conservative force or a friction force. That is, for a macroscopic motor moving with roughly a constant velocity, both a viscous drag and a conservative load simply oppose the motor motion and they have approximately the same effect on the motor. For a protein motor, the situation is very different. Because protein motors are very small, the effect of inertia is negligible and the motor is driven mostly by the random Brownian force. The instantaneous velocity changes direction very rapidly and its absolute value is several orders of magnitude larger than the average velocity. Consequently, the drag force on the motor is stochastic and cannot be treated as a conservative force. The effect of a conservative load force on the motor is different from that of a viscous drag force that simply opposes the motor motion in any direction and whose magnitude increases with the velocity.4) Therefore, for protein motors, it is necessary to distinguish the case where the motor is loaded with a conservative force and the case where the motor is loaded with a viscous drag. When a protein motor works against a viscous load, the thermodynamic efficiency defined above does not apply. A commonly employed measure of efficiency in this case is the Stokes efficiency, defined by replacing f in Eq. (5) with the average viscous 4) A conservative load force tends to drive the motor backwards (opposite to the positive motor direction) and its magnitude is independent of the velocity. A viscous drag simply damps the motor velocity, while the Brownian force and/or the chemical reaction excite the veloc-

ity. If the chemical reaction is halted, the viscous drag will relax the motor to thermal equilibrium with the surrounding fluid, while the conservative force will drive the motor in the opposite direction.

13

14 Force Generation in Molecular Motors

drag force, f D = ζ v. Here ζ = kB T/D is the drag coefficient and D is the diffusion coefficient, which can be computed or measured independently [12]: η Stokes =

ζ vδ , −G

or

η Stokes =

ζ v2 −G·r

(6)

Although ζ v2 has the dimension of energy per unit time, ζ v2 is not the rate of the work done by the motor motion on the fluid medium5) . Indeed, it is not clear what energy per unit time ζ v2 measures. However, since ζ v2 increases with v, the quantity ζ v2 does measure some aspect of the motor’s mechanical performance. So the Stokes efficiency is the ratio of this ‘mechanical performance’ index to the energy supply. Of course, for Eq. (6) to be a valid measure of efficiency, it has to satisfy ηStokes ≤ 100 %. This is true, but the proof is not trivial. One can show that Eq. (6) measures how close the motor comes to delivering a constant force [1, 12, 13]. Given the free energy supply — G, the maximum average velocity is achieved if – G is utilized to generate a roughly constant driving force (independent of motor position). When the driving force is constant, the Stokes efficiency is 100%. When the Stokes efficiency is close to 100%, the driving force is close to a constant force [13]. A high Stokes efficiency for the F1 motor was one of the factors that implicated the progressive binding of ATP as the force-generating process, for only in this fashion could a constant torque be generated [1]. To summarize, if the thermodynamic efficiency, ηTD , is close to 1, this means that the motor motion is tightly coupled to the chemical reaction. However, when the Stokes efficiency, ηStokes , is close to 1, this means that the motor is delivering a nearly constant force. Since the Stokes and thermodynamic efficiencies measure different properties of the motor, it is worthwhile to measure both efficiencies experimentally.

7 Discussion

The basic principle underlying force generation in molecular motors seems at first glance pretty simple: proteins use short range attractive intermolecular forces to bias small thermal fluctuations or to rectify larger diffusions. The former we call ‘power strokes’, the latter ‘Brownian ratchets’. But the details are all important, and they are devilishly diverse. At some level of abstraction the operation of a molecular motor can be viewed as the motion of a point moving down a multidimensional free energy surface [1, 20, 21]. Indeed, many models start from this viewpoint and try and deduce what the laws of physics can say about the general properties of 5) ζ v2 should not be confused with ζ v2 . By the equipartition theorem of statistical meachanics, ζ v2 equals ζ kB T m−1 at equilibrium,

where m is the motor mass. ζ v2 is several orders of magnitude larger than ζ v2 .

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes 15

the surface. While useful from the viewpoint of theory, models at this level of abstraction are likely to be unsatisfying to biologists who seek a more mechanistic understanding, akin to how an engineer would understand the design and operation of an automobile motor. This involves dealing with the details of protein structure. The study of molecular motors, more than most other areas in biology, draws on a diversity of fields. Biochemistry elucidates the kinetics of the energy supplying reactions and thermodynamics establishes constraints on the energetic transactions. Mutation studies isolate the key functional amino acids and mechanical measurements provide criteria that circumscribe possible mechanisms. Finally, microscopy and X-ray crystallography provides the sine qua non structures, for it is nearly impossible to deduce how a machine works without knowing what it looks like. However, all of these studies combined cannot produce a mechanistic theory of how motor proteins work; this requires the unifying power of mathematical modeling. For until the operation of a motor can be formulated as equations and solved, our knowledge is, at best, qualitative and uncertain, hardly better than a plausible cartoon that may, or may not, obey the laws of chemistry and physics and in any event cannot be compared quantitatively with experiments.

Appendix

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes

Here we discuss four model examples to illustrate the role of Brownian fluctuations and the classification of power strokes and Brownian ratchets. For simplicity, we consider the one-dimensional motion of an object. A1.1 Example 1: A power stroke without Brownian fluctuations

Suppose the object is deterministically moved forward a fixed unit distance, x, per unit time, t (e.g. 0.01 nm in 1 µs). The trajectory of the object is shown in Figure A1a. The object moves forward uniformly with a velocity of (x)/(t). This model example is a pure power stroke without Brownian fluctuations. It is mostly relevant for macroscopic motors. Because of the large inertia, macroscopic motors tend to move at a roughly constant velocity and the effect of Brownian fluctuations is negligible. For protein motors, because of the small size, at room temperature the motion is dominated by Brownian fluctuations. One may argue that this is a hypothetical example of a power stroke protein motor at zero temperature.

16 Force Generation in Molecular Motors

Fig. A1 Trajectory of the object for model 1. The object moves forward uniformly with an average velocity of v = x/t = x/1. (b) Trajectory of the object for model 2. The object is driven by a constant force and is subject to Brownian fluctuations. The average velocity of the object is v = x/t. (c) Trajectory of the object for model 3. The object is moved by Brownian fluctuations, and the net drift comes from biasing fluctuations. The average velocity of the object is v = 0.92·x/t. The free energy consumption per unit length is the same as

that in Example 2. Examples 2 and 3 have similar statistical behaviors, and so it is experimentally difficult to distinguish between them. (d) Trajectory of the object for model 4. The object is moved by Brownian fluctuations, and the net drift comes from rectifying large fluctuations. The free energy consumption per unit length is the same as that in Examples 2 and 3 but the average velocity of the object is v = 0.1 x/t, significantly lower than that of Examples 2 and 3.

Since protein motors can only function in a certain temperature range and in a certain fluid medium, this model is not really relevant for protein motors, and so we must modify it to incorporate Brownian fluctuations. A1.2 Example 2: A power stroke with Brownian fluctuations

Suppose, in addition to the deterministic unit forward displacement per unit time, in each unit time the object makes n = 10 independent random moves. Each random move is one unit displacement either forward or backward with equal probability, p (e. g. flipping a fair coin: p = 0.5). This random motion is added to simulate

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes 17

Brownian fluctuations. The trajectory of the object is shown in Fig. A1b. The average velocity of the object is v = x/t, but the motion is stochastic. If we view the displacement in each unit time as the ‘instantaneous velocity’ for that unit time, then the instantaneous velocity is much larger than the average velocity. The inset shows the details of the trajectory; it is evident that the deterministic drift is buried in the random motions and can be detected only by looking at long time correlations. The number of random moves per unit time is proportional to the diffusion coefficient, D, of the object: 2Dt = n(x)2 . This allows us to calculate the diffusion coefficient from which the drag coefficient, ζ , is computed from the Einstein relation: ζ = kB T/D. In this example, the constant force driving the deterministic forward motion is f = ζ v and the free energy consumption per unit length is δvx. For n = 10, the free energy consumption per unit length is 0.2 kB T. A model such as this is appropriate for describing a charged object, such as a DNA strand, driven through a fluid medium by an electrical potential gradient. A1.3 Example 3: A Brownian ratchet that biases fluctuations

In the two examples above, the net drift is caused directly by a driving force (e. g. an electric potential gradient), and the Brownian fluctuations do not affect the net drift. Next, we consider situations where the net drift is actually caused by biasing or rectifying Brownian fluctuations. In the absence of other influences, the forward and backward fluctuations have the same probability. However, if internal barriers are established to block, partially or completely, the backward fluctuations, then the object is more likely to fluctuate forward. Suppose in each unit time, t, the object makes 10 independent random moves (fluctuations). Each random move is 1 unit displacement, x, either forward or backward with equal probability (p = 0.5). Suppose that each time the object passes a multiple of 5 × x, a barrier is established at that location. At a barrier, the object can fluctuate forward in two ways: a forward Brownian fluctuation and a backward Brownian fluctuation that is reflected by the barrier. The object can move back past the barrier only if a backward Brownian movement ‘breaks’ the barrier. The probability of breaking a barrier depends on the strength of the barrier. Let pf be the probability of forward fluctuation at the barrier and pb be that of backward fluctuation. G/kB T = log(pf /pb ) gives the free energy (in units of kB T) required to break the barrier. If we use pf = 0.73 and pb = 0.27, the corresponding free energy drop at the barrier is 1 kB T Since the barriers are separated by 5 × x, the free energy consumption per unit length is 1/5 = 0.2 kB T, the same as that used in Example 2. The trajectory of the object is shown in Fig. 1Ac. The average velocity of the object is v = 0.92 × x/t, similar to Example 2. Because of the small free energy drop (1 kB T) associated with each barrier, it only partially blocks backward fluctuations. The inset shows the details of the trajectory. This model is an example of a Brownian ratchet that biases small fluctuations. The motion of the object is stochastic and indistinguishable

18 Force Generation in Molecular Motors

from that of Example 2 where the object is driven by a constant force and subject to Brownian fluctuations. In this example, the object is directly moved by Brownian fluctuations. The net drift results from biasing fluctuations. For this reason, this model can be classified as a Brownian ratchet. However, it has the same phenomenological behavior as the power stroke motor in Example 2, and so it is very difficult — and unnecessary — to experimentally distinguish it from a power stroke motor. A1.4 Example 4: A Brownian ratchet that rectifies fluctuations

Finally, consider the model Example 3, but now suppose that in each unit time, t, the object makes 10 independent random moves (fluctuations). Each random move is one unit displacement, x, either forward or backward with equal probability (p = 0.5). Now suppose that each time the object passes a multiple of 100 × x, a barrier of 20 kB T is established at that location. The barrier is very high: the probability of a forward fluctuation is pf = 1–3.8 × 10−11 ≈1 and the probability of a backward fluctuation that surmounts the barrier is pb = 3.8 × 10−11 ≈ 0. Since the barriers are 100 × x apart, the free energy consumption per unit length is 20/100 = 0.2 kB T, the same as that in Examples 2 and 3. The trajectory of the object is shown in Fig. A1d. The average velocity of the object is v = 0.1x/t, significantly lower than that of Examples 2 and 3. Because of the large free energy drop associated with each barrier, it almost completely blocks backward fluctuations. Thus, this model is a Brownian ratchet that rectifies large fluctuations. The stochastic motion of the object is different from that of Examples 2 and 3. The object advances slowly in large ratchet steps: once it passes a multiple of 100x, it almost never goes back. One can add a ‘load’ to the above examples by decreasing the probability of a forward fluctuation and increasing that of a backward fluctuation at every location. One can also combine the power stroke and ratchet by adding both a deterministic motion and larger barriers, or alternatively by adding both small barriers and larger barriers. But this would complicate the model and obscure the simplicity of the distinctions we are trying to illustrate. However, we should point out that there is a way to use the time series data itself to estimate how much of the motor driving force can be ascribed to ratchet and power stroke contributions. This involves constructing an Effective Driving Potential from the time series data of the motor; this is discussed in Wang and Oster [12].

A2 A Closer Look at Binding Free Energy

The simple description given in the text of ATP binding to the catalytic site of F1 or of ions binding to the rotor of F0 , conceals a great deal of complexity because it neglects the role of solvent effects. Charges in aqueous solution are always hydrated, surrounded by a shifting cohort of hydrogen bonded waters. Before ATP can bind

A2 A Closer Look at Binding Free Energy

Fig. A2 Enthalpic and entropic changes during desolvation and binding can be followed by plotting—H, vs. TS. In these coordinates, the free energy change, G, is plotted as linear contour lines decreasing up and to the right. The formation of one hydrogen bond between the enzyme and nucleotide can be represented schematically as a reversible path showing a single desolvation and binding process. In state 1, the enzyme and nucleotide sites are hydrated (solid circles). Removing a water molecule from one site entails an enthalpic increase, H12 . This is followed by an entropic increase, TS23 , as the water escapes into solution. Finally, the removed water hydrogen bonds with other waters resulting in an enthalpic decrease, H34 . Similar changes accompany the release of a water molecule

from the other site during the transition from state 4 to state 7. Now the two empty sites must be brought close together, entailing an entropic decrease, TS78 , and an enthalpic decrease, H89 as the sites bind. Thus the overall free energy change, G19 , has enthalpic and entropic components (black bars) that depend on many factors, especially whether the water binds more strongly to the sites than to other waters. Note that the sequence 1 → 9 is only meant to show the enthalpic and entropic changes. It does not represent the actual sequence of what occurs during the desolvation and binding process. In particular, 2 → 3 (the removed water diffusing into solution) and 3 → 4 (the removed water bonding with other waters) occur simultaneously and cannot be separated.

to the catalytic site to initiate the hydrolysis cycle (or sodium binding to the rotor charge in the F0 motor cycle), both must shed their water coats. The shedding of the water coats is progressive as the hydrogen bonds form between ATP and the catalytic site. Before each hydrogen bond can form, the hydration water molecules must be shed from the donor and receptor just before they form the bond (otherwise they will be re-hydrated quickly). Consider the overall process of the formation of one hydrogen bond between ATP and the catalytic site. This entails a number of energetic and entropic transactions. We can plot the process schematically as the path shown in Fig. A2. (This path does not represent the actual non-equilibrium process, but rather a ‘reversible work’ path to illustrate the separate entropic and enthalpic transactions). We see that, even in the simplest case, a single association event entails four enthalpic and four entropic changes when water molecules break their hydrogen bonds to charged sites, escape into solution, re-bond with other

19

20 Force Generation in Molecular Motors

waters, and finally two sites associate. Clearly, solvent effects can tip the free energy balance, but it is seldom easy to compute how. The binding transition that generates the power stroke as embodied in Eq. (2) and Fig. 3 is also an oversimplified description. The hydrogen bonds are not arrayed in a linear sequence (as suggested by the term ‘binding zipper’), nor are they either ‘on’ (zipped) or ‘off’ (unzipped). The bonding surfaces are complex, hydrogen bonds have a finite range and angular dependence, and thermal motions create a stochastic pattern of graded bonding interactions. Nevertheless, molecular dynamics studies demonstrate that the free energy changes gradually and nearly linearly as the nucleotide unbinds or binds to the catalytic site.

References 1 OSTER, G. and H. WANG. 2000a. Reverse engineering a protein: The mechanochemistry of ATP synthase. Biochim. Biophys. Acta 1458: 482–510. 2 WANG, H. and G. OSTER. 1998. Energy transduction in the F1 motor of ATP synthase. Nature 396: 279–282. 3 BOCKMANN, R. 2002. Nanoseconds molecular dynamics simulation of primary mechanical energy transfer steps in F1 -ATP synthase. Nat. Struct. Biol. 9: 198–202. 4 SUN, S., et al. 2003. Elastic energy storage in beta sheets with application to F1 -ATPase. Eur. Biophys. J. 32: 676–683. 5 OSTER, G. and H. WANG. 2000b. Why is the efficiency of the F1 ATPase so high? J. Bioenerg. Biomembr. 332: 459–469. 6 ANTES, I., et al. 2003. The unbinding of ATP from F1 -atpase. Biophys. J.. 85: 695–706. 7 DIMROTH, P., et al. 1999. Energy transduction in the sodium F-ATPase of Propionigenium modestum. Proc. Natl. Acad. Sci. USA 96: 4924–4929. 8 OSTER, G., et al. 2000. How F0 -ATPase generates rotary torque. Proc. Roy. Soc. 355: 523–528. 9 MAHADEVAN, L. and P. MATSUDAIRA. 2000. Motility powered by supramolecular springs and ratchets. Science 288: 95–99. 10 MOGILNER, A. and G. OSTER. 1996a. Cell motility driven by actin polymerization. Biophys.J. 71: 3030–3045. 11 MOGILNER, A. and G. OSTER. 1996b. The physics of lamellipodial protrusion. Euro. Biophs. J. 25: 47–53.

12 WANG, H. and G. OSTER. 2002a. Ratchets, power strokes, and molecular motors. Appl. Phys. A 75: 315–323. 13 WANG, H. and G. OSTER. 2002b. The Stokes efficiency for molecular motors and its applications. Europhys. Lett. 57: 134–140. 14 MENZ, R., et al. 2001. Structure of bovine mitochondrial F1 -ATPase with nucleotide bound to all three catalytic sites: implications for the mechanism of rotary catalysis. Cell 106: 331–341. 15 STOCK, D., et al. 2000. The rotary mechanism of ATP synthase. Curr. Opin. Struct. Biol. 10: 672–679. 16 JUNGE, W., et al. 2001. Inter-subunit rotation and elastic power transmission in F0 F1 -ATPase. FEBS Lett. 251: 1–9. 17 UEMURA, S., et al. 2002. Kinesin–microtubule binding depends on both nucleotide state and loading direction. Proc. Natl Acad. Sci. USA 99: 5977–5981. 18 HANCOCK, W. and J. HOWARD. 1999. Kinesin’s processivity results from mechanical and chemical coordination between the ATP hydrolysis cycles of the two motor domains. Proc. Natl Acad. Sci. USA 96: 13147–13152. 19 PESKIN, C. S. and G. OSTER. 1995. Coordinated hydrolysis explains the mechanical behavior ofkinesin. Biophys.J. 68: 202s–210s. 20 BUSTAMANTE, C., et al. 2001. The physics of molecular motors. Acc. Chem. Res. 34: 412–420.

References 21 WANG, H., et al. 1998. Force generation in RNA polymerase. Biophys.J. 74: 1186–1202. 22 ISRAELACHVILI, J. and B. NINHAM. 1977. Intermolecular forces the long and short of it. J. Colloid Interface Sci. 58: 14–25.

23 PEDERSEN, P., et al. 2000. ATP Synthases in the year 2000: evolving views about the structures of these remarkable enzyme complexes. J. Bioenerget. Biomembr. 32: 325–332.

21

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

1

Enzyme Catalysis Andreas Sebastian Bommarius Georgia Institute of Technology, Atlanta, USA

Bettina R. Riebel Emory University School of Medicine, Atlanta, USA

Orginally published in: Biocatalysis. Andreas Sebastian Bommarius and Bettina R. Riebel. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30344-1

1 Introduction

Enzymes are a class of macromolecules with the ability both to bind small molecules and to effect reaction. Stabilizing forces such as hydrophobic effects only slightly dominate destabilizing forces such as Coulombic forces of equal polarity; thus the Gibbs free enthalpy of formation of proteins, Gformation , is only weakly negative. In an enzyme reaction, initially free enzyme E and free substrate S in their respective ground states initially combine reversibly to an enzyme–substrate (ES) complex. The ES complex passes through a transition state, Gtr,cat , on its way to the enzyme–product (EP) complex and then on to the ground state of free enzyme E and free product P. From the formulation of the reaction sequence, a rate law, properly containing only observables in terms of concentrations, can be derived. In enzyme catalysis, the first rate law was written in 1913 by Michaelis and Menten; therefore, the corresponding kinetics is named the Michaelis–Menten mechanism. The rate law according to Michaelis–Menten features saturation kinetics with respect to substrate (zero order at high, first order at low substrate concentration) and is first order with respect to enzyme. Important milestones in the rationalization of enzyme catalysis were the lockand-key concept [1], Pauling’s postulate (1944) and induced fit [2]. Pauling’s postulate claims that enzymes derive their catalytic power from transition-state stabilization; the postulate can be derived from transition state theory and the idea of a thermodynamic cycle. The Kurz equation, kcat /kuncat ≈ KS /KT , is regarded as the mathematical form of Pauling’s postulate and states that transition states in the case of successful catalysis must bind much more tightly to the enzyme than Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

2 Enzyme Catalysis

ground states. Consequences of the Kurz equation include the concepts of effective concentration for intramolecular reactions, cooperativity of numerous interactions between enzyme side chains and substrate molecules, and diffusional control as the upper bound for an enzymatic rate. Enzymes demonstrate rate accelerations (kcat /kuncat ratios) of ≤ 1.4 · 1017 and proficiencies [kcat /(kuncat K M ) (= 1/K T )], of ≤ 10 M−1 . Surprisingly, for a range of enzymatic reactions kcat is within two orders of magnitude whereas kuncat varies by more than six orders of magnitude; the highest rate accelerations are observed for reactions with low kuncat . Every (bio)catalyst can be characterized by the three basic dimensions of merit – activity, selectivity and stability – as characterized by turnover frequency (tof) (= 1/kcat ), enantiomeric ratio (E value) or purity (e.e.), and melting point (T m ) or deactivation rate constant (kd ). The dimensions of merit important for determining, evaluating, or optimizing a process are (i) product yield, (ii) (bio)catalyst productivity, (iii) (bio)catalyst stability, and (iv) reactor productivity. The pertinent quantities are turnover number (TON) (= [S]/[E]) for (ii), total turnover number (TTN) (= mole product/mole catalyst) for (iii) and space–time yield [kg (L · d)−1 ] for iv). Threshold values for good biocatalyst performance are kcat > 1 s−1 , E > 100 or e.e. > 99%, TTN > 104 –105 , and s.t.y. > 0.1 kg (L · d)−1 .

2 Characterization of Enzyme Catalysis 2.1 Basis of the Activity of Enzymes: What is Enzyme Catalysis?

Enzymes are a class of multifunctional, multivalent macromolecules with the ability to bind small molecules and, much more importantly, subsequently to effect reaction. Stabilizing forces such as hydrophobic effects [3] only slightly dominate destabilizing forces such as Coulombic forces of equal polarity; thus the Gibbs free enthalpy of formation, Gformation , is weakly negative. Without exception, enzymes are proteins and consist of one or more linear chains of, with rare exceptions, the common 20 proteinogenic amino acids. The fairly frequent disulfide bonds between cysteines are formed through crosslinking of side-chain interactions but do not constitute branching of the backbone chain. Extremely rarely, an additional genetically coded 21st amino acid is found, such as selenocysteine in formate dehydrogenase or glutathione peroxidase [4, 5], or pyrrolysine, the “22nd” amino acid, in methanogens belonging to the domain of archaea [6, 7], or p-fluorophenylalanine in a redesigned E. coli cell [8]. All three examples used a stop codon to code for the unusual amino acid. Non-proteinogenic amino acids are found much more commonly in non-enzyme proteins. One prominent example is collagen, from gelatin, which mainly consists of triplets of hydroxyproline–glycine–proline, in which the hydroxyproline is generated via posttranslational hydroxylation. Other proteins apart from enzymes

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

2 Characterization of Enzyme Catalysis 3

exist that can bind substrates and catalyze reactions, such as catalytic antibodies [9, 10] or ribozymes [11, 12]. On the other hand, there are many proteins that do not catalyze any reaction, and thus are not enzymes, but function as oxygen storage and shuttle units (hemoglobin and myoglobin), receptors (signal transduction proteins such as the EGF-receptor) or as structural proteins (myelin). 2.2 Enzyme Reaction in a Reaction Coordinate Diagram

The idea of an enzyme reaction consisting of both a binding step and a subsequent reaction step is embodied in a free enthalpy-reaction coordinate (G-ξ ) diagram depicting the change in Gibbs free enthalpy G over the extent of reaction ξ (zeta) (Figure 1). In an uncatalyzed reaction, one or more substrates or reactants initially are in their respective ground states; during the reaction, the reactants pass the point of maximum free enthalpy Gtr,uncat , termed the “transition state”, and continue to the ground state of the product(s). In an enzyme reaction, initially free enzyme E and free substrate S likewise are in their respective ground states. From the ground state, enzyme and substrate combine reversibly to an enzyme–substrate (ES) complex. If [S] > K M , the ES complex forms an “intermediate”, i.e., a local minimum of free enthalpy. (If [S] < K M , the free enthalpy of the ES complex is higher than the enzyme’s ground state; at [S] = K M , both free enthalpy levels are the same.) The ES complex passes through another transition state, Gtr,cat , on its way to the enzyme–product (EP) complex and then on to the ground state of free enzyme E and free product P. The maximum of the Gibbs free enthalpy between the ground states of substrate and product forms the Gibbs free enthalpy of activation with the energy difference G= , which determines the rate constant of the reaction. Like every catalyst, an enzyme decreases the value of G= and thus accelerates the reaction. (An agent increasing the value of G= is termed an “anti-catalyst”.)

Fig. 1 Free enthalpy reaction coordinate diagram for an enzyme reaction.

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Enzyme Catalysis

2.3 Development of Enzyme Kinetics from Binding and Catalysis

From the idea of enzyme kinetics as a binding and a reaction step with the corresponding course of the energy curve in the Gibbs free enthalpy–reaction coordinate (G – ξ ) diagram, the reaction scheme represented by Eq. (1) can be drawn. E+S⇔ES→EP→(⇔)E+P

(1)

Among the several ways of verifying or disproving such a reaction scheme, the derivation of a rate law linking a product formation rate or substrate consumption rate with pertinent concentrations of reactants, products, and auxiliary agents such as catalysts probably has the greatest utility, as conversion to product can be predicted. A proper rate law contains only observables, and no intermediates or other unobservable parameters. In enzyme catalysis, the first rate law was written in [13] by Michaelis and Menten (the corresponding kinetics is therefore aptly named the Michaelis–Menten (MM) mechanism). The kinetic schemes for the description of enzyme reactions are similar to those for homogeneous and heterogeneous catalysis and were developed roughly contemporaneously. In heterogeneous catalysis, a mechanism postulating a reaction of a mobile substrate molecule on a solid surface is modeled by a rate law named after Langmuir and Hinshelwood, and later also after Hougen and Watson [14] (LHHW). LHHW and MM both assume a limiting number of active sites which can be saturated with reacting molecules from a continuous phase. The kinetic scheme according to Michaelis–Menten for a one-substrate reaction [13] assumes three possible elementary reaction steps: (i) formation of an enzyme–substrate complex (ES complex), (ii) dissociation of the ES complex into E and S, and (iii) irreversible reaction to product P. In this scheme, the product formation step from ES to E + P is assumed to be rate-limiting, so the ES complex is modeled to react directly to the free enzyme and the product molecule, which is assumed to dissociate from the enzyme without the formation of an enzyme–product (EP) complex [Eq. (2)]. E+S⇔ES→E+P

(2)

In the derivation according to Michaelis and Menten, association and dissociation between free enzyme E, free substrate S, and the enzyme–substrate complex ES are assumed to be at equilibrium, K S = [ES]/([E] · [S]). [The Briggs–Haldane derivation (1925), based on the assumption of a steady state, is more general.] With this assumption and a mass balance over all enzyme components ([E]total = [E]free + [ES]), the rate law in Eq. (3) can be derived. v=vmax [S]/(K M +[S])=kcat [E]total [S]/(K M +[S])

(3)

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

3 Sources and Reasons for the Activity of Enzymes as Catalysts

In contrast to [E]free , [E]total is observable. Eq. (3) is written with K M , the Michaelis constant, instead of the equilibrium binding constant K S ; unless the enzyme reaction is very fast (Section 4.3.); i.e., in almost all cases, K M ≈ K S . In Eq. (3), the reaction rate is traditionally denoted by v [concentration/time] and kcat is the reaction rate constant [time−1 ]. The equation describes a two-parameter kinetics, with a monotonically rising reaction rate with respect to substrate concentration and saturation at high substrate concentration. The maximum reaction rate at saturation is denoted by vmax , with vmax = kcat [E]. The K M value corresponds to the substrate concentration at half saturation (vmax /2) and is a measure of binding affinity of the substrate to the enzyme: a high K M value corresponds to loose binding, and a low value to tight binding between enzyme and substrate. At low substrate concentration, more precisely if [S] K M , Eq. (3) simplifies to v = vmax [S]/K M and consequently is first order with respect to the substrate concentration [S]. In contrast, at high substrate concentration, more precisely if [S] K M , Eq. (3) simplifies to v = vmax and consequently is zeroth order with respect to [S]. In all situations, v is proportional (first-order with respect) to [E]. 3 Sources and Reasons for the Activity of Enzymes as Catalysts 3.1 Chronology of the Most Important Theories of Enzyme Activity

Just one year after Ostwald’s hypothesis about the existence of catalysts in 1893, when nobody yet had a clear idea of the structure and composition of enzymes, Emil Fischer voiced the idea for the first time that a substrate molecule fits into the pocket of an enzyme, the “lock-and-key hypothesis” [1]. Both the lock (enzyme) as well as the key (substrate) were regarded as rigid. This hypothesis was modified later in many ways. According to Haldane [15], catalysis of a reaction occurs only if a catalyst in the active center is complementary to the transition state of the substrate during the reaction. Therefore, the transition state between substrate and products fits best into a pocket close to the enzyme. The substrate molecule is subject to strain upon binding to the active center and changes its conformation to fit into the active center; the key (the substrate) does not fit completely into the lock but is strained and bent. In a further modification of this concept, a reaction is accelerated if a catalyst stabilizes the transition state; in contrast, stabilization of the ground state leads to a deceleration of the reaction. This concept of transition-state stabilization, formulated first by Haldane, was expanded later by Linus Pauling [16, 17]. The notion of lowering of G= attributed to the stabilization of the transition state by the catalyst or to the destabilization of the ground state in comparison to the transition state is generally accepted nowadays. Instead of assuming a “solid” enzyme, in which active center the substrate molecule is “bent” (the concept of substrate strain), the idea was developed [2, 18]

5

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

6 Enzyme Catalysis

that enzymes can embrace the substrate molecule flexibly in the active center and effect reaction by the formation of specific interactions, the so-called “induced fit”. This picture is especially appropriate with allosterically activated enzymes or in situations in which part of the enzyme molecule has to turn or move over longer distances to effect catalysis (hinge movement), as for instance with most NAD(P)(H)dependent enzymes [19]. An equation for the acceleration of the reaction rate by a deliberate catalyst and the relative strength of the binding to the transition state and ground state was formulated in 1963 by Kurz [20], who combined the ideas of the thermodynamic cycle and transition-state theory and whose result, unrecognized by Kurz, can be regarded as a quantification of the Pauling postulate [20]. Just a few years later Jencks originated the idea of transition-state analogs as inhibitors and cited references in the literature [21]. Finally, it was observed [22] that the best inhibitors for serine proteases such as subtilisin, chymotrypsin, trypsin or elastase all led to geometrically similar transition states for the hydrolysis reaction and all bound in the complementary oxyanion hole. All four proteases belonged to the small number of enzymes whose three-dimensional structure was known by then. 3.2 Origin of Enzymatic Activity: Derivation of the Kurz Equation

The key equation resulting from the application of two known theories, the Born–Haber cycle and the transition-state theory, was formulated for any catalyst and without reference to enzymes [20]; a good derivation can be found in the article by Kraut [23]. Transition-state theory is based on two assumptions, the existence of both a dynamic bottleneck and a preceding equilibrium between a transition-state complex and reactants. Eq. (4) results, where k denotes the observed reaction rate constant, κ the transmission coefficient, and ν the mean frequency of crossing the barrier. k=κ·ν·K =

(4)

All correction factors such as tunneling, back-crossing of the barrier, and solvent frictional effects are captured by κ, which is between 0.1 and 1 for the reactions in solution in question here. The equilibrium constant K = is expressed by partition functions, where the mode normal to the reaction coordinate ν is approximated by the term kB T/hν. In the resulting Eq. (5), ν cancels out.

k=κ·(k B T/ h)·K =

(5)

Note that K = is not the thermodynamic equilibrium constant and kB T/h is not a universal frequency for the decomposition of the transition-state complexes into products. The thermodynamic cycle relevant for further discussion is shown in Figure 2.

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

3 Sources and Reasons for the Activity of Enzymes as Catalysts E +

S

E + S≠

KS

KT ES

E + P

[ES] ≠

Fig. 2 Thermodynamic (Born–Haber)-cycle.

If one compares the first-order rate constant for the enzyme-catalyzed singlesubstrate elementary reaction ke with the one for the uncatalyzed reaction ku , Eq. (6) is obtained with Eq. (4). ke /ku =(κe ·νe ·K e= )/(κu ·νu ·K u= )

(6)

In the case of a simple elementary enzyme reaction, ke is identical to kcat . By utilizing the thermodynamic cycle the quotient of the constants of formation of the transition state K e = /Ku = can be equated with the quotient of the dissociation constants for substrate KS and transition state K T [Eq. (7)]. kcat /ku =(κe ·νe ·K S )/(κu ·νu ·K T )

(7)

Although only a few data are available for the comparison of κ e ·ν e , and κ u ·ν u , it is assumed that the two terms do not differ much in magnitude, so that Eq. (8) holds approximately. kcat /ku ≈K S /K T

(8)

This is the equation derived by Kurz, expressed here for the case of an enzyme reaction. The ratio of enzyme-catalyzed to uncatalyzed reaction kcat /ku or kcat /kuncat , i.e., the rate acceleration, often reaches orders of magnitude of 1010 to 1012 [24], and in the best case so far 1.4 · 1017 (orotidine monophosphate decarboxylase, OMPD), which translates into a proficiency kcat /(kuncat · K M ) of 1023 M−1 [25]. This underscores the quality of enzymes as catalysts. In comparison, the best value for the rate acceleration of catalytic antibodies is 2.3 · 108 [26], and typical values are 103 to 105 [27]. Radzicka [24] found that kcat for the reactions in question is often within one or two orders of magnitude; interestingly, the highest efficiencies are observed with reactions of very low kuncat . Thus, Eq. (8) means that transition states in the case of successful catalysis must bind much more tightly to the enzyme than ground states; in this way, Eq. (8) is the mathematical form of the Pauling postulate in 1944 [17]. It is emphasized again that the action of enzymes through the stabilization of the transition state is not an independent concept but is derived from transition-state theory and the idea of a thermodynamic cycle. 3.3 Consequences of the Kurz Equation

The Kurz equation [Eq. (8)] has simplified and channeled many discussions in enzymology Many other explanations of enzyme action and many phenomena

7

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

8 Enzyme Catalysis

observed with enzyme reactions can be reduced to the validity of Eq. (8), as shown in the following paragraphs. Ĺ The principle of transition state stabilization can explain why many enzymes show higher activity against larger, sterically more demanding substrate molecules than against smaller ones. Paradoxically, the cause of the enhanced activity was an increased kcat , not a decreased K M value, i.e., an enhanced maximum rate and not a tighter substrate binding. This phenomenon led to the postulation of “induced fit”. But if the enzyme template binds the transition state more tightly than the ground state, it can be expected that peripheral parts of the substrate molecule have an important role for the binding in the transition state but not necessarily for the binding in the ground state. Conformational changes of the enzyme template do not need to be postulated. Frequently, however, conformational changes during binding and catalysis are observed which are then also called “induced fit” by several authors. Ĺ The exact loci of binding and catalysis cannot be distinguished exactly. In subtilisin, the amino acids forming the catalytic triad, Ser221–His64– were replaced in all combinations by Ala. Even the triple mutant without any amino acid from the original catalytic center displayed a 1000-fold higher reaction rate than the uncatalyzed reaction, and the remainder of the enzyme molecule bound the substrate better in the transition state than in the ground state [28]. Ĺ The numerous interactions between enzyme side chains and substrate molecules act synergistically, so the fit is quite exact. This cooperativity enhances substrate specificity, for a small change of substrate can effect a large change in the binding of the transition state. Triose phosphate isomerase is only 1/1000 as effective if the residue Glu165 is moved away from the substrate by even 1 Å, as was demonstrated with the exchange of Glu165 for Asp165 [29]. The change of only one amino acid of the triad in subtilisin causes an activity decrease of 10−6 , whereas the exchange of all three causes a decrease of only 7 × 10−7 and not of 10−18 [28]. The results seem to be explained by an all-or-nothing situation rather than additivity Ĺ Intramolecular reactions proceed much more rapidly than intermolecular ones owing to entropic stabilization. Page and Jencks have calculated the entropies of translation, rotation, and vibration [30]. The entropy loss of the reactants already realized in the enzyme–substrate (ES) complex allows for a much faster reaction than in the situation without complexation, binding, or geometric orientation. Introduction of an effective concentration [S]eff alleviates the trouble of comparing results of intermolecular catalysis with rate constants of dimension [concentration · time]−1 (second-order reaction) with those of intramolecular catalysis with constants of dimension [time]−1 . The high values often achieved for [S]eff (often > 10 M) reflect the efficiency of intramolecular catalysis. In this context, enzymes are often referred to as “entropy traps”, because many contributions to entropy are “frozen in” after binding of the substrate to the enzyme molecule. Ĺ Many enzymatic reaction mechanisms are accelerated or even only made possible by the absence of water or other solvents. The enzyme supports desolvation

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

3 Sources and Reasons for the Activity of Enzymes as Catalysts Table 1 Wholly or partially diffusion-controlled enzyme reactions adapted from [52]

Substrate

Enzyme

k cat [s−1 ]

K M [M]

kcat /KM [s−1 · M−1 ]

CO2 ⇔ HCO3 − H2 O2 fumarate benzylpenicillin

Carbonic anhydrase Catalase Fumarase β-Lactamase

1 × 106 4 × 107 800 2 × 103

0.012 1.1 5 × 10 −6 2 × 10−5

8.3 × 107 4 × 107 1.6 × 108 1 × 108

by redirecting part of the possible binding energy liberated from the binding between enzyme and substrate. Ĺ Upper boundary of rate constant of an enzyme reaction: diffusion control The upper boundary of the reaction rate is reached when every collision between substrate and enzyme molecules leads to reaction and thus to product. In this case, the Boltzmann factor, exp(−E a /RT), is equal to 1 in the transition-state theory equations and the reaction is diffusion-limited or diffusion-controlled (owing to the difference in mass, the reaction is controlled only by the rate of diffusion of the substrate molecule). The reaction rate under diffusion control is limited by the number of collisions, the frequency Z of which can be calculated according to the Smoluchowski equation [Smoluchowski, 1915; Eq. (9)]. −dc/dt=−8π RD·c 2

(9)

R is the normalized radius of both participating particles (1/r 1 + 1/r 2 ), D the normalized diffusion coefficient (1/D1 + 1/D2 ). The collision frequency Z is then calculated from Eq. (10). Z=2RT/(3000η·[(r 1 +r 2 )2 /r 1 r 2 ])

(10)

As the enzyme molecule is much larger than the substrate or product molecule (r E r S ) and thus diffuses much more slowly (DE DS ), Eqs. (9) and (10) can be simplified to Eqs. (11) and (12). −dc/dt=−8π RDE ·c 2

(11)

Z=2RT/(3000η·rE )

(12)

In the case of a highly viscous solution the influence of viscosity η can dominate [31]. For simple, uncharged particles in water at 25 ◦ C the second-order rate constant is 3.2 × 10 (M · s)−1 [32, 33]. Some cases of wholly or partially diffusion-controlled enzyme reactions are listed in Table 1. Rearrangement of Eq. (7) results in Eq. (13). kcat /K S =(ku ·κe ·ve )/(κu ·vu ·K T )

(13)

9

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

10 Enzyme Catalysis

Eq. (13) is the manifestation of the second-order rate constant for a reaction between free enzyme and free substrate to give free enzyme and free product; thus, the rate constant cannot exceed the maximum value for kdiff , i.e., 3.2 × 109 M−1 · s−1 . A perfectly evolved enzyme therefore features a decreased binding constant K s , which means a stronger binding in the transition state, possibly up to the thermodynamic limit. Limitation of a reaction by translational diffusion in solution is a rather rare case. Much more frequently the limitation of the observed overall reaction rate is by external mass transfer (through a laminar film around a solid macroscopic carrier) or internal mass transfer (diffusion of substrate or product through the pores of a solid carrier or a gel network to an enzyme molecule in the interior of the carrier). 3.4 Efficiency of Enzyme Catalysis: Beyond Pauling’s Postulate

Let us return one more time to the Kurz equation [Eq. (8)], which is regarded as the quantification of Pauling’s postulate that transition-state stabilization, i.e., the tighter binding between enzyme and transition state, expressed by K T , as compared to binding between enzyme and ground state, expressed by K S , is the source of catalytic rate enhancement. kcat /kuncat ≈K S /K T

(8)

Identifying K S with K M , K T approximately equals the ratio (kuncat · K M )/kcat , which is the inverse of termed the “proficiency”, which has been determined experimentally for a number of systems (see Section 4.4 below). If the Kurz equation captured the whole essence of enzyme catalysis, K T should be proportional to (kuncat · K M )/kcat . However, Figure 3 reveals that the correlation of K T (≡ “K TS ”) for some enzyme reactions is much better with kuncat (≡ “knon ”) than with kcat [34]. This seeming contradiction to Pauling’s postulate and the notion of the transitionstate energy and conformation as the sole source of enzymatic catalytic power cast the geometry of the substrate ready to interact with the enzyme, i.e., substrate preorganization, into a new and more important role. While the discussion on this subject is far from settled, a novel concept based on molecular mechanics calculations of the stability of different substrate conformations, termed substrate conformers, has emerged which emphasized substrate preorganization. The substrate conformers which allow reactions to occur most easily clearly resemble the transition state and are called “near-attack conformers” (NACs) [34, 35]. The relative rate constants of anhydride formation krel from mono-p-bromophenyl esters were found to correlate with the mole fractions (P) of ground-state conformers that could be classified as NACs according to Eq. (14) [8]. log krel =0.94log P+7.48 (r 2 =0.915) The results are depicted in Figure 4.

(14)

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Performance Criteria for Catalysts, Processes, and Process Routes

Fig. 3 Comparison of the relationship of K TS with the enzymatic reaction rate constant, kcat , and with the uncatalyzed solution or reference reaction rate constant knon [34].

4 Performance Criteria for Catalysts, Processes, and Process Routes 4.1 Basic Performance Criteria for a Catalyst: Activity, Selectivity and Stability of Enzymes

Every catalyst, and thus also every biocatalyst, can be characterized by the three basic dimensions of merit, namely activity, selectivity, and stability. Additional, but not frequently employed, performance criteria beyond the basic dimensions are discussed in Section 4.3. 1. Biocatalysts often feature much better selectivity than non-biological catalysts, so they are developed for use because of their selectivity, be it enantioselectivity, chemo- or regioselectivity. 2. As activity is straightforward to measure and necessary to know for even the basic experimental protocol, enzyme activity is often well studied. In contrast to other catalysts, most enzymes are only active and stable in a very limited temperature and pH range (mostly between 15 and 50 ◦ C or pH 5 and 10). 3. Unlike activity, stability of enzymes is often interpreted simplistically as thermal stability, i.e., a temperature beyond which the enzyme loses stability. Although this quantity is important, first every statement of stability at a certain temperature depends on exposure time and thus is often ambiguous; and second, for biocatalytic process applications, a more important quantity is the process or operational stability, which is the long-term stability under specified conditions.

11

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

12 Enzyme Catalysis

Fig. 4 Log of the relative rate constants of anhydride formation krel from mono-p-bromophenyl esters vs. the log of the probability (P) of NAC formation of each monophenyl ester of the various dicarboxylic acids [34].

4.1.1 Activity The value of the (overall) enzyme activity is usually provided in “International Units” or “Units”:

1 Unit≡1 IU≡1 U≡1 µmol min−1 In most cases a value for the overall activity of enzyme is not very interesting. Much more important are both the specific activity, scaled to the mass of catalyst, and the volumetric activity, based on the activity per unit volume: 1 U (mg protein)−1 =1µmol (min mg protein)−1 ≡specific activity and 1 U mL−1 =1µmol (min mL)−1 ≡volumetric activity All activity data are meaningless without a specification of the conditions of measurement. Connonen, the activity of enzymes is provided at nearly physiological

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Performance Criteria for Catalysts, Processes, and Process Routes

conditions at 30 ◦ C and pH 7.5, unless specified otherwise. Any enzyme sold commercially comes with information about the detailed assay conditions used to arrive at the specific (solid preparation) or volumetric (liquid preparation) activity. To judge the quality of a biocatalyst, its specific activity can serve as a guideline: the threshold for a useful biocatalyst is 1 U (mg pure protein)−1 ; the threshold for a biocatalyst developed for plant-scale level, however, lies much higher, rather closer to 100 U (mg pure protein)−1 . The volumetric activity of a (bio)catalyst can be enhanced by simple addition of more catalyst to a system. Specific activity, however, must be improved through optimization of the reaction conditions, or through variation of the structure of the carrier or even of the enzyme. A mass-independent quantity of activity, superior to specific activity, is the turnover frequency (tof), defined as: turnover frequency (tof)=

number of catalytic events time×number of active sites

The turnover frequency allows performance comparison between different catalyst systems, biological and/or non-biological. Its threshold is at 1 event per second per active site. According to the definition, a turnover frequency can be determined only if the number of active sites is known. For an enzyme reaction obeying Michaelis–Menten kinetics, Eq. (15) holds. tof=1/kcat

(15)

4.1.2 Selectivity The notion of selectivity needs to be specified further: point selectivity is the incremental selectivity, usually towards product, at a specific degree of conversion, whereas integral selectivity is the overall average selectivity at the same specific degree of conversion. Owing to the importance of enantiomeric purity of target product molecules in life science applications and the pre-eminent position biocatalysts enjoy with respect to the achievement of that goal, enantioselectivity is the most important kind of selectivity in the context of biocatalysis. The enantiomeric ratio or E value [36] serves as a measure of enantio-selectivity at a certain degree of conversion [Eq. (16)].

E =(kcat /K M )A /(kcat /K M )B =ln([A]/[A]0 )/([B]/[B]0 )

(16)

Although the E value is measured at a specific degree of conversion, nevertheless it is an integral selectivity measure. This is even more obvious in the case of the other measure commonly employed, the enantiomeric excess, e.e. [Eq. (17)], which reflects overall selectivity up to the point of isolation of the product. e.e.=([A]−[B])/([A]+[B])×100%

(17)

13

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

14 Enzyme Catalysis

The threshold for an enantioselective (bio)catalyst is either at E = 100 or at 99% e.e., depending on whether one is evaluating either the reaction itself or a reaction product. [These values are linked to the requirement by the Food and Drug Administration of the United States (FDA), the most important body in the world for setting guidelines for both procedures for novel drug approval and drug performance evaluation, that a separate toxicological study be conducted for any impurity, including an enantiomer of the active drug, exceeding a concentration level of 0.5%.] Besides enantioselectivity, regioselectivity and to a lesser extent, chemoselectivity are also important issues of enzymatic reaction selectivity. 4.1.3 Stability Stability of an enzyme is usually understood to mean temperature stability, although inhibitors, oxygen, an unsuitable pH value, or other factors such as mechanical stress or shear can decisively influence stability. The thermal stability of a protein, often employed in protein biochemistry, is characterized by the melting temperature T m , the temperature at which a protein in equilibrium between native (N) and unfolded (U) species, N ⇔ U, is half unfolded. The melting temperature of a protein is influenced on one hand by its amino acid sequence and the number of disulfide bridges and salt pairs, and on the other hand by solvent, added salt type, and added salt concentration. Protein structural stability was found to correlate also with the Hofmeister series [37–39] [Eq. (18)].

Tm =T0 +K [S]

(18)

In Eq. (18), T 0 is the melting temperature in the absence of salt, [S] the salt concentration, and K the corresponding coefficient. Storage stability over time under fixed conditions of temperature, pH value and concentration of additives often can be expressed by a first-order decay law (analogous to radioactive decay) [Eq. (19)]. [E ]t =[E]0 exp(−kd ·t)

(19)

The validity of a first-order decay law over time for the activity of enzymes according to Eq. (19), with [E]t and [E]0 as the active enzyme concentration at time t or 0, respectively, and kd as the deactivation rate constant, is based on the suitability of thinking of the deactivation process of enzymes in terms of Boltzmann statistics. These statistics cause a certain number of active protein molecules to deactivate momentarily with a rate constant proportional to the amount of active protein [for evidence for such a “catastrophic” decomposition, see Craig [40]]. The relevant stability criteria for process applications (the process stability or operating stability) is discussed in the next section.

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Performance Criteria for Catalysts, Processes, and Process Routes

4.2 Performance Criteria for the Process

In evaluating a biocatalyst for a given processing task, there are performance criteria to be met not only for the biocatalyst but also for the process. The dimensions of merit important when determining, evaluating, or optimizing a route for a process are (i) product yield, (ii) (bio)catalyst productivity, (iii) (bio)catalyst stability, and (iv) reactor productivity. 4.2.1 Product Yield Chemical yield to product is most important for the economics of the process. Product yield is inversely proportional to the amount of reactants required per unit of product output. As the key substrate and other raw materials in most mature processes make up more than 50% of the variable cost, a high product yield is indispensable for an economic process. The yield of product y is linked to selectivity σ and the degree of conversion x by Eq. (20).

y=x·σ

(20)

so that the product concentration [P] can be calculated from Eq. (21). [P]=[S]0 ·y=[S]0 ·x·σ

(21)

Yields much less than 100% are no longer acceptable either economically or ecologically. In the separation of a racemate, where the yield per run is limited to 50% per pass, this means that internal or external racemization is necessary unless either the substrate is inexpensive enough to lose up to 50% or the co-product can be marketed in similar amounts. Both of the latter situations are highly unlikely. A threshold for a sufficiently high yield is hard to define, as the threshold value seems to correlate inversely with the unit value of the product. While for basic, large-volume chemicals yields of 98 or 99% are absolutely essential, the situation in fine chemicals calls for 90–95% yield, and in the initial stage of production of extreme performance chemicals, such as pharmaceuticals, yields of > 80% are very acceptable; sometimes values down to 50% have to be encountered. Acceptable yields depend on the number of process steps, including product isolation. If all the steps are assumed to fetch 90% yield, the overall yield depends on the number of steps n as in Eq. (22). yoverall =(ystep )n =(0.9)n

(22)

so that for a one-step process yoverall,1 = 90%, for three-step process yoverall,3 = 73%, and for a ten-step process yoverall,10 = 35%!

15

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

16 Enzyme Catalysis

4.2.2 (Bio)catalyst Productivity If a large amount of (bio)catalyst is added to a substrate solution better results are achieved than if just a small amount of a highly productive catalyst is used. To even call an agent a catalyst, one condition is that the agent be added, as a minimum, in substoichiometric amounts. The smaller the amount of catalyst that has to be added for the same result, the better its performance. The relevant criterion is the dimensionless turnover number, TON [Eq. (23)].

TON=

substrate amount of product = =[S]/[C] amount of catalyst catalyst ratio

(23)

The turnover number is not used frequently in biocatalysis, possibly as the molar mass of the biocatalyst has to be known and taken into account to obtain a dimensionless number, but it is the decisive criterion, besides turnover frequency and selectivity, for evaluation of a catalyst in homogeneous (chemical) catalysis and is thus quoted in almost every pertinent article. Another reason for the low popularity of the turnover number in biocatalysis, apart from the challenge of dimensionality, is the focus on reusability of biocatalysts and the corresponding greater emphasis on performance over the catalyst lifetime instead of in one batch reaction as is common in homogeneous catalysis [41]. For biocatalyst lifetime evaluation, see Section 4.2.3. The turnover number of a catalyst does not refer to the timescale of activity of that catalyst in a process. If the timescale of activity is taken into consideration, the productivity of the catalyst is recovered, expressed as a productivity number, PN, defined in Eq. (24). PN=

mass of product unit catalyst×time

(24)

4.2.3 (Bio)catalyst Stability As in all catalytic processes, catalyst stability is a key process criterion. In contrast to mere temperature or storage stability, which refer to the catalyst independently of a process, the operating stability or process stability is the relevant and decisive dimension of merit. It is determined by comparing the amount of product generated with the amount of catalyst spent. The relevant quantity, also sometimes found in homogeneous catalysis, is the total turnover number (TTN) [Eq. (25)].

TTN=

moles of product produced mole of catalyst spent

(25)

Quite common in applied biocatalysis, where the purity of biocatalyst often is not known, is the expression of biocatalyst stability as an the enzyme consumption number (e.c.n.) [Eq. (26)]. e.c.n.=

g of enzyme amount of enzyme preparation spent = amount of product generated kg (or lb) of product

(26)

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Performance Criteria for Catalysts, Processes, and Process Routes

The e.c.n. value depends on process parameters such as temperature, pH value, and concentrations of substrate(s) and product(s). With the molar masses of enzyme and product, e.c.n. and TTN can be interconverted [Eq. (27)]. TTN=(1000/e.c.n.)×(MWenzyme /MWproduct )

(27)

It should be emphasized that the TTN is not a completely suitable quantity for the evaluation of operating stability because the number of moles of biocatalyst is not a suitable reference for the complexity and cost of its manufacture. However, the values for both numerator and denominator in Eq. (27) are usually known and can be expressed in monetary terms. For checking the application of such a biocatalyst, the contribution of the biocatalyst to the overall cost can be assessed readily. The relevant parameter for studies of operating stability of enzymes is the product of active enzyme concentration [E]active and residence time τ , [E]active · τ . In a continuously stirred tank reactor (CSTR) the quantities [E]active and τ are linked by Eq. (28), where [S0 ] denotes the initial substrate concentration, x the degree of conversion and r(x) the conversion-dependent reaction rate [42, 43]. ([E]active ·τ )/[S0 ]=x/r (x)

(28)

For a comparison and discussion of the concepts of stability at rest [Eq. (19)] and operating stability [Eq. (28)]. The threshold value for sufficient biocatalyst stability depends on the application, as does the value for product yield. For any application in synthesis the TTN should exceed 10 000, and for large-scale processing a value of > 1 000 000 is preferred. 4.2.4 Reactor Productivity For the assessment of reactor productivity that is independent of catalyst, the spacetime yield (s.t.y) [Eq. (29)]. is considered, as in any chemical process.

s.t.y.=mass of product generated/(reactor volume×time)[kg(L·d)−1 ]

(29)

An increase in s.t.y. in the same reactor is equivalent to an increase in reaction rate. When considering the Michaelis–Menten equation [Eq. (3): v = vmax [S]/(K M + [S]) = kcat [E][S]/(K M + [S])], there are three possible ways to achieve a maximum reaction rate: 1. High substrate concentration (increase of space-yield). Enzyme reactions can be accelerated by increasing the substrate concentration up to the limit of saturation (≈ 10K M ). If [S] K M , the enzyme is saturated and Eq. (3) is reduced to Eq. (30).

v=vmax =kcat ·[E]

(30)

17

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

18 Enzyme Catalysis

At substrate saturation, the reaction is zeroth-order with respect to substrate. With a loosely binding substrate, i.e., a high K M value, the enzyme is often saturated only above the solubility limit of the substrate. To improve the reaction rate or the s.t.y. further, according to Eq. (29) the yield per unit time must be increased, which means enhancing either the biocatalyst concentration and/or the time constant kcat . 2. High enzyme concentration. The reaction rate and s.t.y. can be enhanced by increasing the catalyst concentration [E]; in practice, however, in contrast to the formalism of Eq. (3), owing to an either excessive viscosity increase or excess of deactivated protein in the reactor, a maximum limit of enzyme concentration is reached. 3. Increase in the time constant. According to Eq. (3), kcat signifies the time constant of the enzyme reaction [time−1 ], and the corresponding reaction is performed over a time scale 1/kcat [time]. An increase can be effected through the temperature (Arrhenius behavior) but also for instance through changing a protecting group in peptide synthesis [44]. An increase in kcat also increases the acceleration ratio kcat /kuncat [24] of the enzyme catalyst, in contrast to case 2) (see Section 4.3).

A minimum threshold value for reactor productivity can be set at a space-timeyield of about 100 g (L · d)−1 , a value which tends to be compromised more by lack of substrate solubility than by biocatalyst reactivity. Well-developed biocatalytic process often feature space–time yields of > 500 g (L · d)−1 or even > 1 kg (L · d)−1 [44–46]. 4.3 Links between Enzyme Reaction Performance Parameters 4.3.1 Rate Acceleration While users of biocatalysts are often concerned first and foremost about a high maximum rate vmax , a high kcat value, and possibly also a low K M value, a good (bio)catalyst is one that enhances the chemical background rate by as high a factor as possible, at least 105 , possibly 1010 or even more. The dimensionless ratio of the enzyme catalytic rate constant over the uncatalyzed (i.e., the chemical background) rate constant, kcat /kuncat , is called the rate acceleration. An alternative nomenclature used in the literature, the efficiency or proficiency, should be reserved for the term kcat /(kuncat · K M ) quantifying the total energetic cost of enzymatic catalysis [47]. Surprisingly, when analyzing the source of good rate accelerations, i.e., high kcat /kuncat values, or the source of high proficiencies, i.e., high kcat /(kuncat · K M ) values, kcat was found to be within two orders of magnitude for a variety of enzymes whereas kuncat varied by several orders of magnitude, irrespective of the value of kcat (Figures 5 and 6; [24, 48]). The data provides clear guidance that the biggest improvement in enzyme catalysts can be achieved for reactions with very low chemical background rate constants and not by optimizing rate constants or specificities which are already fairly high.

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Performance Criteria for Catalysts, Processes, and Process Routes

10 5 rate constant (e–1)

enzyme rate constants (kcat)

catalyzed

1 t 1/2 1 minute

10 –5 t1/2 1 week uncatalyzed

t 1/2 100 years 10

–10

t 1/2 1 million years 10–15 t 1/2 4 billion years

nonenzymatic rate constants (knon)

Fig. 5 Logarithmic scale comparison of kcat and kuncat (= knon ) for some representative reactions at 25 ◦ C. The length of each vertical bar represents the rate enhancement. [48]. ADC: arginine decarboxylase; ODC: orotidine 5 -phosphate decarboxylase; STN:

staphylococcal nuclease; GLU: sweet potato $-amylase; FUM: fumarase; MAN: mandelate racemase; PEP: carboxypeptodase B; CDA: E. coli cytidine deaminase; KSI: ketosteroid isomerase; CMU: chorismate mutase; CAN: carbonic anhydrase.

4.3.2 Ratio between Catalytic Constant kcat and Deactivation Rate Constant kd The rate equation for a deactivating enzyme in a batch reactor [Eq. (31)] reads:

x·[S]0 −K M ln(1−x)=[E]0 ·(kcat /kd )·{1−exp(−kd ·t)}

(31)

The influence of deactivation depends linearly on the dimensionless ratio kcat /kd , which might serve as a ratio to assess quickly the potential of a deactivating enzyme for synthesis. 4.3.3 Relationship between Deactivation Rate Constant kd and Total Turnover Number TTN Specific enzyme consumption [U per kg product], a hands-on parameter, can be obtained from Eq. (32) [49], where avol,0 [U L−1 ] is the initial volumetric activity).

specific enzyme consumption=(avol,0 ·kd )/(s.t.y.)[U(kg product)−1 ]

(32)

A high value for s.t.y. and low values for [E] (a modified enzyme concentration [g enzyme L−1 ]), avol,0 , or kd all lead to a favorable, i.e., low, specific enzyme consumption. To convert the specific enzyme consumption into the total turnover number TTN, the specific enzyme consumption, sp.e.c. [U (kg product)−1 ] [Eq. (32)] first has to be

19

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

20 Enzyme Catalysis 1010 k cat / K m

enzyme efficiencies (k cat /K m)

(M –1 s–1)

catalyzed

10 5

1 t 1/2 1 minute

10 –5 t1/2 1 week uncatalyzed

t 1/2 100 years 10 –10 rate constant (s–1)

k non (s–1)

t 1/2 1 million years 10 –15 t 1/2 4 billion years

nonenzymatic reaction constants (k non)

Fig. 6 Logarithmic scale comparison of kcat /K M and kuncat (≡ knon ) for some representative reactions at 25◦ C. The length of each vertical bar represents the transition-state affinity or catalytic proficiency (its reciprocal). For abbreviations, see Figure 5 [48].

converted into the enzyme consumption number e.c.n. [g enzyme (kg product)−1 ] [Eq. (26)].

e.c.n.=(sp.e.c.·[E] )/avol,0 [g enzyme(kg product)−1 ]

(33)

If Eq. (32) is inserted into Eq.(33) and the resulting equation into Eq. (27),the result is Eq.(34), which with [E] /MWenzyme = [E]0 [mol L−1 ] becomes Eq. (35). TTN[mol product(mol catalyst)−1 ] =(MWenzyme /MWproduct )(100×s.t.y.)/([E] ·kd )

(34)

TTN=(1000×s.t.y.)/MWproduct ·[E]0 ·kd

(35)

A favorable, i.e., high, TTN is achieved by driving up the s.t.y. of a pertinent experiment while keeping both [E]0 and kd low. (The conversion factor of 1000 does not appear if s.t.y. is expressed in [g (L · d)−1 ] and not [kg (L · d)−1 ].)

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

4 Performance Criteria for Catalysts, Processes, and Process Routes

4.4 Performance Criteria for Process Schemes, Atom Economy, and Environmental Quotient

Roger Sheldon, of TU Delft (Delft, The Netherlands), has compiled a list of trends for chemical and biochemical process technology [50]; they are towards Ĺ Ĺ Ĺ Ĺ Ĺ

catalytic reactions instead of stoichiometric ones, reactions with 100% selectivity at 100% conversion, high substrate concentrations, no detrimental solvents, and no change of solvent during the process, enhanced use of solid or volatile acids and bases as well as pH-stat techniques.

We recognize the importance of high conversion, high selectivity, and thus high product yields; and of high substrate concentration, often leading to high s.t.y. and TTN. However, there are several additional guidelines which help to optimize a complete process. Some of those criteria have been discussed above in the context of the catalyst or the process, such as enantioselectivity or diastereoselectivity instead of simple selectivity to product, and catalyst performance data such as turnover frequency (tof), turnover number (TON), total turnover number (TTN), and space–time-yield (s.ty). Others have to deal with inputs and outputs of the process apart from reactants, catalysts, and products, such as: 1. the atom economy, i.e., the fraction of carbon (or other elements) of the substrate that is utilized or recovered in the product 2. the specific consumption (consumption per kg product) of solvents, salts, and other auxiliaries such as work-up materials (active carbon, filter aids), and 3. the degree of recovery of all materials, ranging from main process components to solvents, salts, and traces as well as contents of trace streams, often termed ancillary losses.

ad 1) Atom economy The degree of utilization of inputs in the final product is termed the atom economy [51]. An atom-economic process uses as many atoms as possible in the reaction stoichiometry Examples of simple, but atom-economic, chemical processes are the manufacture of maleic anhydride by air-oxidation of butane/butene instead of benzene, or of propylene oxide by air-oxidation of propylene (propene) instead of by ring-closing substitution of epichlorohydrin (with loss of HCl). An example of a process with low atom economy is any synthesis with protection/deprotection steps, such as those that occur during peptide synthesis (while possibly still featuring very high product yields). ad 2) Specific consumption of solvents, salts, and auxiliaries This can be captured by evaluating the environmental quotient, EQ [50], consisting of a relative mass of by-product versus target product [kg kg−1 ] (or for our American readers, lb lb−1 )

21

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

22 Enzyme Catalysis Table 2 Comparison of conditions for laboratory- and industrial-scale processes

Medium Substrate pH control Time Temperature, pH gradients

Laboratory-scale processes

Industrial-scale processes

dilute solutions often natural with buffer sufficient for reaction none

(highly) concentrated media often unnatural titration with acid/base; solid acids/bases fast reactions desired heat and mass transfer influence reaction

multiplied by a measure of the environmental unfriendliness. Water or even NaCl might carry a factor of 1 or close to 1, whereas mercury, halogenated solvents, or other toxic by-products might carry a factor of 10 or even 100. ad 3) Recovery of all materials Eco-balances have become increasingly popular with regulators to gauge the impact of discharges. Increasingly, such instruments replace simple levying of costs on discharges or setting inflexible upper limits for concentrations of a range of ecologically unfriendly agents. When aiming for scale-up or when actually already running a process on a pilot or a full industrial scale instead of trying to gain knowledge about a problem under controlled laboratory conditions, one should ensure that the different goals are reflected in different measures taken to obtain data or to develop a catalyst or process for an ultimate large scale. Table 2 contrasts typical laboratory practices with conditions prevailing on production scale. It is prudent, however, for processes to be practiced on a large scale to be developed accordingly, even during the test phase on lab scale. Suggested Further Reading Alan FERSHT, Structure and Mechanism in Protein Science, Freeman, New York, 1999.

William P. JENCKS, Mechanisms in Chemistry and Enzymology, Dover, New York, 1975.

References 1 E. FISCHER, Einfluss der Configuration auf die Wirkung der Enzyme, Ber. dtsch. chem. Ges. 1894, 27, 2985–2993. 2 D. E. KOSHLAND Jr., Application of a theory of enzyme specificity to protein synthesis, Proc. Natl. Acad. Sci. U.S.A., 1958, 44, 98–104. 3 C. TANFORD, Physical Chemistry of Macro-molecules, Wiley, New York, 1961. 4 I. CHAMBERS, J. FRAMPTON, P. GOLDFARB, N. AFFARA, W. MCBAIN, and P. R. HARRISON, The structure of the mouse

glutathione peroxidase gene: the selenocysteine in the active site is encoded by the ‘termination’ codon, TGA, EMBO J. 1986, 5, 1221–1227. 5 F. ZINONI, A. BIRKMANN, T. C. STADTMAN, and A. BOECK, Nucleotide sequence and expression of the selenocysteinecontaining polypeptide of formate dehydrogenase (formate-hydrogenlyase-linked) from Escherichia coli, Proc. Natl. Acad. Sci. USA 1986, 83, 4650–4654.

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

References 6 B. HAO, W. GONG, T. K. FERGUSON, C. M. JAMES, J. A. KRZYCKI, and M. K. CHAN, A new UAG-encoded residue in the structure of a methanogen methyltrans-ferase, Science 2002, 296, 1462–1466. 7 G. SRINIVASAN, C. M. JAMES, and J. A. KRZYCKI, Pyrrolysine encoded by UAG in Archaea: charging of a UAG-decoding specialized tRNA, Science 2002, 296, 1459–1462. 8 R. A. MEHL, J. C. ANDERSON, S. W. SANTORO, L. WANG, A. B. MARTIN, D. S. KING, D. M. HORN, and P. G. SCHULTZ, Generation of a bacterium with a 21 amino acid genetic code, J. Am. Chem. Soc. 2003, 125, 935–939. 9 S. J. POLLACK, J. W. JACOBS, and P. G. SCHULTZ, Selective chemical catalysis by an antibody, Science 1986, 234, 1570–1573. 10 A. TRAMONTANO, K. D. JANDA, and R. A. LERNER, Chemical reactivity at an antibody binding site elicited by mechanistic design of a synthetic antigen, Proc. Natl. Acad. Sci. USA 1986, 83, 6736–6740. 11 T. R. CECH, RNA as an enzyme, Sci. Am. 1986, 255(5), 64–75. 12 T. R. CECH, RNA. Fishing for fresh catalysts, Nature, 1993, 365, 204–205. 13 L. MICHAELIS and M. L. MENTEN, Die Kinetik der Invertinwirkung, Biochem. Z. 49, 1913, 333–369. 14 O. A. HOUGEN and K. M. WATSON, Chemical Process Principles, Part III, Wiley, New York. 1947. 15 J. B. S. HALDANE, Enzymes, M.I.T. Press, Cambridge/MA, USA, 1965. 16 L. PAULING, Molecular architecture and biological reactions, Chem. Eng. News 1946, 24, 1375–1377. 17 L. PAULING, Chemical achievement and hope for the future, Am. Sci. 1948, 36, 51–58. 18 D. E. KOSHLAND Jr., G. NEME´ THY, and D. FILMER, Comparison of experimental binding data and theoretical models in proteins containing subunits, Biochemistry 1966, 5, 365–385. 19 T. J. STILLMAN, A. M. B. MIGUEIS, X.-G. WANG, P. J. BAKER, L. BRITTON, P. C. ENGEL, and D. W. RICE, Insights into the mechanism of domain closure and

20

21

22

23 24

25

26

27

28

29

30

31

substrate sof glutamate dehydrogenase from Clostridium symbosium, J. Mol. Biol. 1999, 285, 875–885. J. L. KURZ, Transition state characterization for catalyzed reactions, J. Am. Chem. Soc. 1963, 85, 987–991. W. P. JENCKS, in: Catalysis in Chemistry and Enzymology, McGraw-Hill, New York, 1969, pp. 288–291. J. KRAUT, Serine proteases: structure and mechanism of catalysis, Ann. Rev. Biochem. 1977, 46, 331–358. J. KRAUT, How do enzymes work?, Science 1988, 242, 533–540. A. RADZICKA and R. WOLFENDEN, A proficient enzyme, Science 1995, 267, 90–93. J. K. LEE and K. N. HOUK, A proficient enzyme revisited: the predicted mechanism for orotidine monophosphate decarboxy-lase, Science 1997, 276, 942–945. G. ZHONG, R. A. LERNER, and C. F. BARBAS III, Enhancement of the repertoire of catalytic antibodies with aldolase activity by combination of reactive immunization and transition state theory, Angew. Chem. Int. Ed. 1999, 111, 3957–3960. J. HASSERODT, Organic synthesis supported by antibody catalysis, Synlett 1999, 12, 2007–2022. P. CARTER and J. A. WELLS, Dissecting the catalytic triad of a serine protease, Nature, 1988, 332, 564–568. D. JOSEPH-MCCARTHY, L. E. ROST, E. A. KOMIVES, and G. A. PETSKO, Crystal structure of the mutant yeast triosephos-phate isomerase in which the catalytic base glutamic acid 165 is changed to aspartic acid, Biochemistry 1994, 33, 2824–2829. M. I. PAGE and W. P. JENCKS, Entropic contributions to rate accelerations in enzymic and intramolecular reactions and the chelate effect, Proc. Natl. Acad. Sci. USA 1971, 68, 1678–1683. U. EICHHORN, A. S. BOMMARIUS, K. DRAUZ, and H.-D. JAKUBKE, Synthesis of dipeptides by suspension-to-suspension conversion via thermolysin catalysis – from analytical to preparative scale, J. Peptide Sci. 1997, 3, 245–251.

23

c02 (JWBK148-Bommarius)

January 17, 2008

18:57

Char Count=

24 Enzyme Catalysis

32 H. GERISCHER, J. F. HOLZWARTH, D. SEIFERT, and L. STROHMAIER, Flow method with integrating observation for very fast irreversible reactions, Ber. Bunsenges. Physik. Chem. 1969, 73, 952–955. 33 C. CREUTZ and N. SUTIN, Vestiges of the inverted region for highly exergonic electron-transfer reactions, J. Am.Chem. Soc. 1977, 99, 241–243. 34 T. C. BRUICE and S. J. BENKOVIC, Chemical basis for enzyme catalysis, Biochemistry 2000, 39, 6267–6274. 35 T. C. BRUICE, A View at the Millenium: the efficiency of enzymatic catalysis, Acc. Chem. Res. 2002, 35, 139–148. 36 C. S. CHEN, Y. FUJIMOTO, G. GIRDAUKAS, and C. SIH, Quantitative analyses of biochemical kinetic resolutions of enantiomers, J. Am. Chem. Soc. 1982, 104, 7294–7299. 37 F. HOFMEISTER, Zur Lehre von der Wirkung der Salze. Zweite Mitteilung, Arch. Exp. Pathol. Pharmakol. 1888, 24, 247–260. 38 P. H. VON HIPPEL and K.-Y. WONG, Neutral salts: the generality of their effects on the stability of macromolecular conformation, Science 1964, 145, 577–580. 39 J. K. KAUSHIK and R. BHAT, A mechanistic analysis of the increase in the thermal stability of proteins in aqueous carboxylic acid salt solutions, Protein Sci. 1999, 8, 222–233. 40 D. B. CRAIG, E. A. ARRIAGA, J. C. Y. WONG, H. LU, and N. J. DOVICHI, Studies on single alkaline phosphatase molecules: reaction rate and activation energy of a reaction catalyzed by a single molecule and the effect of thermal denaturation – the death of an enzyme, J. Am.Chem. Soc. 1996, 118, 5245–5253. 41 H. U. BLASER, F. SPINDLER, and M. STUDER, Enantioselective catalysis in fine chemicals production, Appl. Catal. A: General 2001, 221, 119–143. 42 C. WANDREY, Reaction engineering studies on enzyme catalysts for the development of continuous processes, 913: Habilitation Thesis, TH Hannover, Germany, 1977. 43 A. S. BOMMARIUS, K. DRAUZ, U. GROEGER, and C. WANDREY, 1992, Membrane

44

45

46

47

48

49

50

51

52

53

54

bio-reactors for the production of enantiomer ically pure α-Amino acids, in: Chirality in Industry, A. N. COLLINS, G. N. SHELDRAKE, and J. CROSBY (eds.), Wiley & Sons Ltd., London, 930: Chapter 20, pp. 371–397. A. FISCHER, A. S. BOMMARIUS, K. DRAUZ, and C. WANDREY, A novel approach to enzymatic peptide synthesis using highly solubilizing Nα -protecting groups of amino acids, Biocatalysis 1994, 8, 289–307. J. D. ROZZELL, Biocatalysis at commercial scale: myths and realities, Chimica Oggi 1999, 951: (6/7), 42–47. A. S. BOMMARIUS, M. SCHWARM, and K. DRAUZ, Comparison of different chemo-enzymatic process routes to enantiomeri-cally pure amino acids, Chimia 2001, 55, 50–59. E. A. TAYLOR, D. R. J. PALMER, and J. A. GERLT, The lesser ‘burden borne’ by o-succinyl-benzoate synthase: an ‘easy’ reaction involving a carboxylate carbon acid, J. Am. Chem. Soc. 2001, 123, 5824–5825. R. WOLFENDEN and M. J. SNYDER, The depth of chemical time and the power of enzymes as catalysts, Acc. Chem. Res. 2001, 34, 938–945. U. KRAGL, D. VASIC-RACKI, and C. WANDREY, Continuous production of L-tert-leucine in series of two enzyme membrane reactors, Bioproc. Eng. 1996, 14, 291–297. R. A. SHELDON, Consider the environmental quotient, CHEMTECH 1994, 3, 38–47. B. M. TROST, On inventing reactions for atom economy, Acc. Chem. Res. 2002, 35, 695–705. A. FERSHT, Enzyme Structure and Mechanism, 2nd edition, Freeman, New York, 1985. P. T. ANASTAS and T. C. WILLIAMSON, Frontiers in green chemistry, Green Chemistry 1998, 1–26. M. VON SMOLUCHOWSKI, Versuch einer mathematischen Theorie der Koagulations-kinetik kolloidaler L¨osungen, Z. physik. Chem. 1917, 92, 129–168.

1

Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes Ariel B. Lindner, Zelig Eshhar, and Dan S. Tawfik Weizmann Institute of Science, Rehovot, Israel

Originally published in: Catalytic Antibodies. Edited by Ehud Keinan. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30688-6

Abbreviations

TS – transition state; TSA – TS analog; OAS – oxyanion stabilization; OAH – oxyanion hole; AChE – acetylcholinesterase; D-Abs – D-Antibodies (esterolytic antibodies D2.3, D2.4, and D2.5).

1 Introduction

During the past 16 years, knowledge gained from studies of natural hydrolases and the mechanism of hydrolytic reactions led to a variety of strategies for generating antibodies that catalyze hydrolytic reactions. In particular, ester hydrolysis became the most intensely studied antibody-catalyzed reaction. Esterolytic antibodies provided the first ‘proof of principle’ of antibody-based catalysis, spearheading the wide spectrum of antibody catalysts detailed in the chapters of this book. Although esterolytic antibodies do not, in general, exhibit enzyme-like rates, knowledge may be extracted from the many mechanistic and structural studies of hydrolytic antibodies concerning the mechanism of action of hydrolytic enzymes. Comparing hydrolytic antibodies to hydrolytic enzymes is the underlying theme of this chapter. Specific-base-(hydroxyl)-catalyzed and general-base-catalyzed is the mechanism by which most natural esterases act (Fig. 1). The reaction proceeds via relatively low activation barriers (the addition TS being higher than the elimination TS [1]), and a well-defined singular tetrahedral, oxyanionic intermediate (I1 , Fig. 1a). Inherent in hydroxyl-catalyzed ester hydrolysis is the linear of the hydrolytic rates. In implementing these basic mechanisms, which are also seen in solution, enzymes apply complex mechanisms, combining a variety of catalytic forces, as exemplified Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Fig. 1 Mechanisms of ester hydrolysis. (a) Hydroxide-catalyzed ester hydrolysis. The rate-limiting step consists of an exogenous hydroxide attack on the ester’s carbonyl, forming, via the addition TS1, a tetrahedral oxyanionic intermediate (I1 ). The latter collapses, via the elimination TS2, to form the carboxylate and alcohol products. Top formula: a representative phosphonate TSA, where, for the D-antibodies, R = glutaryl glycine and R = p-nitrobenzyl [24]. The numbers correspond to bond lengths de-

rived from ab initio calculations [1]. (b) The endogenous nucleophile mechanism of Serine hydrolases. The nucleophilic attack is carried out by the active site’s seryl alkoxide, activated by an active-site general-base residue (B:) to form a tetrahedral oxyanionic intermediate (I2 ). This is followed by a rate-limiting deacylation step mediated by a hydroxide ion generated in situ by the basecatalyzed proton elimination of an activesite water molecule, releasing the products via a second intermediate (I3 ) [2].

by three major families of hydrolytic enzymes: seryl/cysteyl hydrolases (Fig. 1b), aspartyl hydrolases (Fig. 1c), and zinc hydrolases (Fig. 1d). In essence, two major catalytic forces are utilized by these enzymes [2]: i) A nucleophilic attack is mediated by a precisely positioned nucleophile, either an endogenous general base (seryl alkoxide (Fig. 1b, step I)) or an explicitly bound specific base (a hydroxyl generated via general-base-catalyzed

1 Introduction

Fig. 1 (c) The general base – general acid (“push-pull”) mechanism of Aspartic hydrolases. An active-site bound water molecule is activated by a general-base carboxylate residue, whereas the oxyanion tetrahedral intermediate (I4 ) is stabilized by a proton donation from a second aspartyl residue. Residues maintaining the carboxylates’ protonation level [135] are not shown for clarity. An alternative “symmetric” model can be found in [136]. (d) The metal-mediated

mechanism of Zinc hydrolases. The nucleophilic attack is carried out by a zinc-bound water molecule, activated by a general-base Glu residue. The formed oxyanionic intermediate (I5 ) is stabilized by its coordination to the metal ion. Though a similar fivecoordinated zinc intermediate has recently been observed [137], a four-coordinated intermediate was previously suggested [2, 138].

deprotonation of an active-site water molecule; Fig. 1a, step II, 1c and 1d), to yield a tetrahedral, oxyanionic intermediate. ii) Oxyanionic stabilization (OAS) of the tetrahedral, oxyanionic intermediate and the TSs leading to its formation and breakdown. The negative charge developed on the carbonyl oxygen is stabilized by hydrogen bonding (Fig. 1b, c) or proton transfer (general-acid catalysis), or by a positive charge (Fig. 1d). iii) Protonation of the alkoxide leaving group by general-acid catalysis.

The relative positioning of various active-site residues involved in the above mechanisms provides the recognition and stabilization of the tetrahedral transition

3

4 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

state (TS). It is therefore expected that, although described as separate forces or contributions, their effect on catalysis is coupled. This coupling is indeed encoded in the enzymes’ active-site structure – for example, the seryl residue (Fig. 3), the aspartyl residues (positioned by a shared a hydrogen bond; Fig. 1b) and the zinc ion (Fig. 1c) all provide both the endogenous nucleophile as well as a hydrogen bond to the TS oxyanion. In addition to mechanism above, enzymes take advantage of a combination of other forces, including medium (solvation/desolvation effects), substrate (groundstate) destabilization, remote recognition, as well as co-ordinated conformational changes [2, 3]. Thus, the remarkable catalytic efficiency of natural enzymes, at times limited only by the rate of diffusion, is the outcome of many forces acting in concert. The separation of these factors and the assessment of their independent contributions to catalysis is far from trivial, as exemplified by various mutagenesis studies (e.g., see [4, 5]). Catalytic antibodies, raised to mimic single facets of the complex enzymatic regimes, provide a unique opportunity to examine and quantify enzymatic forces and mechanisms of action individually. 2 Chapter Overview

The first and most widely applicable strategy for generating esterolytic antibodies is by immunizing with analogs of the tetrahedral, oxyanionic intermediates of ester hydrolysis (Fig. 1a), which are generally referred to as transition state analogs (TSAs). Analysis of the structure, mechanism and catalytic efficiency of these antibodies provides the opportunity to compare and assess oxyanion stabilization (OAS) as the primary source of catalysis in both esterolytic antibodies and natural, hydrolytic enzymes (Section 2.1). Surprisingly, several antibodies directed towards OAS (e.g., 43C9 [6]) have incorporated, haphazardly, an endogenous nucleophile as part of their mechanism of action. These antibodies, together with studies attempting to directly obtain antibodies with an endogenous nucleophile (e.g., reactive immunization [7] and chemical modification strategies [8, 9]), form the basis for our discussion on nucleophilic reactivity (Section 2.2). This is accompanied by an analysis of “chemical rescue” experiments, where strong exogenous nucleophiles supplement the initial OAS activity of various antibodies [10–13] (Section 2.3). Several studies, attempting at the generation of esterolytic antibodies with general-acid/base mechanism either by a “bait and switch” strategy (Section 2.4) or metal incorporation (Section 2.5), are then qualitatively assessed. Finally, the roleof conformational changes, suggested in a number of catalytic antibodies [14–20], is described vis-a-vis known contributions of conformational changes to natural enzymes (Section 2). 2.1 Catalysis by Oxyanion Stabilization (OAS)

The most commonly used TSA haptens for the generation of esterolytic antibodies are phosphonates, although other phosphate derivatives such as phosphinates,

2 Chapter Overview

phos-phonoamidates, and phosphothioates have also been used. This indirect approach for the generation of biocatalysts was first formulated by Jencks [21]. Immunization with these analogs generates antibodies that catalyze ester hydrolysis via preferential binding of the reaction’s tetrahedral, oxyanionic TS compared to the planar, uncharged ester substrate (Fig. 1a). This approach relies on optimal representation of the reaction’s TSs by the TSA. It has been shown, for example, that substrate analogs completely fail to generate esterolytic antibodies [22]. Thus, as discussed below, TSA design is a primary issue. 2.1.1 Fidelity of TSA design Phosphate derivatives that are potent inhibitors of hydrolases were shown to exhibit TSA characteristics, albeit with some important discrepancies ([23] and references therein). Ab initio calculations suggest that these TSAs better present the elimination TS2 , vis-a-vis the addition TS1 , the higher and thus the rate-limiting barrier of the two TSs [1] (Fig. 1a). The TSA P-O bonds overestimate the corresponding lengths of the TS, apart from their being significantly shorter (1.48 Å) than the 2.2 Å distance in the addition TS between the incoming hydroxyl nucleophile and the ester’s carbonyl carbon (Fig. 1a). The symmetrical charge partition between the phosphonates’ oxygens overestimates the charge formed in the actual TS and of the incoming hydroxyl nucleophile. Both of the phosphonates’ oxygens are expected to bait antibodies with H-donor residues. This is crucial to stabilizing the developing oxyanionic charge, yet “baiting” a basic residue would be preferable in the position representing the nucleophilic attack. The latter may act as a general base to activate the incoming nucleophile or as an endogenous nucleophile. Nonetheless, despite the above-described infidelities, phosphonate TSAs (and related derivatives) are fairly close mimics of the TSs of ester hydrolysis and hence elicit antibodies in which OAS is the primary, or often the sole, source of catalysis. 2.1.2 Esterolytic Antibodies Based Solely on Oxyanion Stabilization Our comparison of OAS in esterolytic antibodies with natural enzymes is based on a number of antibodies in which OAS was shown to be the only source of the catalytic activity (group I antibodies). This group includes antibodies D2.3, D2.4 and D2.5 [10, 24–26], 48G7 [27, 28], 29G11 [29, 30], and 6D9 [31–33] (Group I; Fig. 2), which show most if not all of the following characteristics: i) The observed rate acceleration correlates well with the differential affinity toward the TSA vs the substrate: K TSA /K S ≥ kcat /kuncat (Table 1) ii) A linear phase is observed in the pH-rate profile iii) Crystal structure indicating active-site residues that comprise the oxyanion hole iv) Mutation of OAS residues eliminates catalysis (Table 2).

Other antibodies (Group II; Fig. 2) for which structural data is available, CNJ206 [14, 34], 17E8 [29, 30, 35], and 7C8 [11] were all raised against similar or identical TSAs to those of Group I, and exhibit K TSA /K S ≤ kcat /kuncat . This suggests that forces additional to or other than OAS take part (Sections 2.2 and 2.4). Nonetheless, their catalytic efficiencies may be taken as an upper limit to the contribution of OAS to

5

6 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Fig. 2 The “oxyanion hole” of esterolytic antibodies. Oxyanion-stabilizing residues are depicted from the TSA complexes of antibodies CNJ206 (1KNO in PDB), 17E8 (1EAP), D2.3 (1YEC), 48G7 (1AJ7), 6D9

(1HYX), and 7C8 (1CT8), viewing from the oxyanion representing the incoming hydroxyl nucleophile. CPK color codes: gray: carbon, red: oxygen, blue: nitrogen, orange: phosphate, and green: chlorine.

their activity. Antibody 43C9, where an endogenous nucleophile was shown to be involved [6], is not considered, as the two mechanisms that affect its catalysis (OAS and nucleophilic catalysis) are difficult to separate (Section 2.2). Detailed comparisons of the hydrolytic antibodies’ crystal structures in the presence of the respective TSAs are available in several reviews [36–37] . A common motif (“canonical binding array”) of interactions with the TSA oxyanionic core was

2 Chapter Overview Table 1 Oxyanion stabilization by hydrolytic antibodies

kcat /kuncat G‡ cat-uncat G◦ TSA-S Catalytic Antibody × 103 Kcal/mol Kcal/mol OAH residuesa)

Secondary OAH residuesb)

Group I D2.3 D2.4 D2.5 48G7 29G11 6D9

110 36 1.9 16 2.2 0.9

7.0 6.3 4.5 5.8 4.6 4.1

7.0 6.3 4.3 6.3 6.0 4.1

Tyr100dH, Asn34L Tyr100eH, Asn34L Tyr100eH, Ser34L-H2 O His35H, (Tyr96H) Lys93H, (Tyr96H) His27d

Trp95H, Tyr96L-H2 O Trp95H, Tyr96L-H2 O Trp95H, Tyr96L-H2 O Arg96L, Tyr33H His35H, Tyr96L –

a) Residues in hydrogen-bond distance from the phosphonate’s oxyanion representing the ester substrate carbonyl (Fig. 1a), assigned by comparison of the antibodies’ phosphonate- and substrate analog-bound structures (D2.3, D2.4 and D2.5) [128], mutagenesis analysis (see Table 2), pH-dependency analysis (D-antibodies [10], CNJ206 [7, 129]C8 [11]), and docking (48G7, 29G11, and 17E8 [39]). b) Residues in hydrogen-bond distance from the phosphonate’s oxyanion representing the incoming hydroxide nucleophile (Fig. 1a) (see text for assignment).

revealed, identifying the residues contributing to the binding of the two oxyanions (Table 1, Fig. 2). Convergent evolution is evident by the sequence similarities as well as structural homology. Antibodies CNJ206 and 17E8 exhibit only 41–52% differences in the primary sequence of the heavy- and light-variable regions, yet share 8 out of 10 contact residues with the hapten. Antibody 48G7 shares the same light chain with CNJ206 and has 7 of 10 of the heavy chain’s CDR residues in common with CNJ206 (for the detailed analysis see [36–38]). Another example of convergence is seen in the structures of D2.3 and D2.4, sharing the same germline genes with 16 positions difference in their variable regions. In particular, the CDR3 H loop of D2.4 differs in 4 positions and includes an amino acid insertion compared to D2.3, as a result of different D-J segments’ junction. Consequently, their CDR3 H loops adopt Table 2 Oxyanion hole mutants of hydrolytic antibodies

Antibody

OAH residue mutant

k cat /kuncat wt = 1.0

D2.3/4b)

AsnL34 → Gly TyrH100d → Phe TyrH100d → Gly TyrH100d → Lys TyrH100d → Ser His35H → Glu His35H → Gln His27d → Ala

1.1 n.d. n.d. n.d. 0.08 0.04 0.6 n.d.

48G7a) 6D9 a) b)

Mutants expressed as recombinant Fab fragments [28]. Mutants expressed as scFv chimera of D2.4 VL /D2.3 VH [10]. n.d. – catalytic activity not detected.

7

8 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

distinctly different structures yet place the critical OAH residue Tyr100d (D2.3) and Tyr100e (D2.4) at the exact same position with respect to the TSA’s oxyanion [26]. The TSAs of Group I antibodies share two epitopes: an aryl or benzyl leaving group, and a phosphonate monoester (Fig. 2). While aromatic moieties are well known as immunogenic epitopes (hence their common use in TSA haptens), Tantillo and Houk suggested that the determining epitope may be the bidentate “transition state epitope” oxyanionic moiety. This is supported by the identification of similar motif in crystal structures of arsonates, sulfonates and DNA binding antibodies. Interestingly, the origin of many of the residues in contact with the TSA phosphonates can be tracked down to the germline genes (e.g. TyrH33 of 48G7, HisH35 of CNJ206 and 48G7, His27d of 6D9). Exceptions do occur though, as in the structures of 6D9 and 7C8 raised against a secondary p-nitrophenyl phosphonate. In contrast to other Group I TSAs, in antibodies 6D9 and 7C8, the linker via which the TSA hapten is linked to a carrier protein, is located on the leaving group and not on the phosphonate (Fig. 2). The active site of 6D9 and 7C8 is shallower, the phosphonate is less buried than in other Group I antibodies, and the active-site residues in contact with the TSA are fewer and different [11]. Despite the evident convergence between Group I’s active sites, the immune system provided some variations within the interactions forming the oxyanion hole (Table 1). The distinction between the position of the oxyanion hole and that of the entering hydroxy nucleophile was determined by the respective structure in the presence of substrate analogs (as in the D-antibodies), by pH profiles pointing to key hydrogen-bond donor residues (D-Abs, CNJ206), and by mutagenesis (D-Abs, 48G7, 17E8), and was complemented by docking in silico (Table 1, Fig. 2) [39]. 2.1.3 Antibody Oxyanion Holes The D-antibodies (D2.3 and D2.4), the most efficient antibodies of Group I, are taken here as a model for examining the contribution of active-site residues acting OAH. Two such residues (Tyr100dH and Asn34L) were suggested to form the crystal structures of the D-antibodies in complex with the TSA and a substrate analog (Fig. 2). In the crystal structure of D2.3 with an amide substrate analog (1YEF in PDB), 3.8 Å is the distance between the aspargine’s δ-amide proton donor and the scissile bond carbonyl-oxygen (compared to the 2.6 Å distance from the oxygen of TyrH100d), suggesting that Asn34L interaction may be specific to the transition state. In addition, D2.5, lacking Asn34L, has 20–50 fold lower catalytic efficiency than D2.3 and D2.4 (Table 1). However, mutating Asn34L to Glycine did not affect the rate of catalysis (Table 2), leaving Tyr100d the primary functional OAH residue. This was further supported by several Tyr100d mutants (Table 2) and pH profiles of binding and catalysis [10]. The pH-binding profiles revealed a 1.6–1.9 units difference in pK a between the substrate-antibody and the TSA-antibody complexes. Thus, the negative charge marking the difference between the substrate and the TSA (and hence the actual TSs of the reaction) dramatically strengthens the tyrosyloxyanion H-bond, thus rendering it the key residue in the D-Abs oxyanion hole [10]. The specific free energies, 6.3 and 7 Kcal/mol for the complexation of D2.4 and D2.3

2 Chapter Overview

with the TSA G◦ TSA –G◦ s ) are identical to the reduction in the activation energy barrier derived for the D-Abs from their rate acceleration (GC‡ cat –G‡ uncat ) (Table 1). This tight correlation suggests that the D-Abs active sites convert binding energy into catalysis with close to 100% efficiency, corresponding to the contribution of the tyrosyl OAH. Judging from Tables 1 and 2, tyrosine is the most efficient single residue for OAS. Further, a hierarchy may be derived where Tyr (contributing up to 7.0 Kcal/mol) > His (4.1 Kcal/mol, assumed from 6D9 single His OAH and48G7 His35HGlu mutant) ≥ Lys (∼3 Kcal/mol, 29G11 Lys-main chain amide OAH combination is weaker than 48G7) > backbone amide (2.2 Kcal/mol, judged by comparison of CNJ206 to wt48G7; 48G7 HisH35Glu mutant’s activity, which is derived from Tyr97H backbone amide, is approximately half that of CNJ206 with two backbone amides). Though highly speculative, this order of activities may be justified by the oxyanion pK a s found for serine hydrolase substrates, ranging between 7 and 10 [40, 41]. It is expected that hydrogen bonds between donors and acceptors with matching pK a would form the strongest hydrogen bonds [42]. Tyrosine’s hydroxyl (pK a ≈10) may afford the closest match to ester’s TS pK a . It is expected that this pK a would be considerably lower than the pK a of the TSs, leading, for example, to amide hydrolysis. The former is often stabilized in nature by OAH comprised of amide bonds that have a pK a much higher than the hydroxyl of tyrosine (see below) [5, 43]. Matching pK a values may also explain the significantly stronger interaction of the D-Abs’ tyrosine (∼7.0 Kcal/mol) compared to the average binding energy of a charged group via a single hydrogen bond (3.5–4.5 Kcal/mol) [44]. 2.1.4 Oxyanion holes–Antibodies vs. Enzymes The versatility of OAS solutions adopted by the immune system is comparable to, or perhaps even higher than, that observed in natural hydrolytic enzymes. Excluding metal hydrolases, which are not easily mimicked by antibodies (Section 2.5), the overall majority of enzymes use a combination of two (e.g., chymotrypsin, trypsin) or three (e.g., AChE, Fig. 3) backbone amides to form their OAHs. Their contribution to the catalytic activity cannot be addressed directly by mutation analysis or separated according to the individual contribution of each backbone NH. OAS could only be assessed by attributing the residual activity of the triad mutants to OAS [e.g., trypsin [45], acetylcholinesterase (AChE) [46]; Table 3]. A minority of enzymes were found to utilize a combination of a backbone NH groups as H-donor together with a side-chain NH group [e.g., Asn in subtilisin (Fig. 3), Gln, or Ser (Table 3)]. Interestingly, the recently characterized prolyl oligopeptidase-like family [43, 47, 48] and the structurally similar cocE esterase (Fig. 3) [49] utilize a tyrosine side chain (Table 3) together with the backbone NH of the seryl nucleophile. Indeed, the efficient OAS by catalytic antibodies such as the D-antibodies may indicate that tyrosine could be as efficient as or even more efficient than NH groups. The contribution of the OAH side chain to the overall catalytic rates of these enzymes was assessed by mutation of the putative side chain (Table 3). Such studies are complicated by mutation-induced structural changes of the active site as clearly

9

10 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Fig. 3 The ‘oxyanion hole’ of hydrolytic enzymes. Oxyanion-stabilizing residues are depicted from the following inhibitor-enzyme crystal structures: acetylcholinesterase (AChE) with m(N,N,N-trimethylammonio)-

2,2,2-trifluoroacetophenone (TMTFA) (1AMN in PDF), cocaine esterase I (cocE) with phenyl boronic acid (1JU3), and subtilisin with D-p-chlorophenyl-1-acetamidoboronic acid (1AVT).

seen in the cocE Tyr44Phe mutant’s structure. As a result of a loss of H-bonding to the mutated tyrosine, three Trp residues change their position, and the Phe ring is tilted by 20◦ compared to the wild-type tyrosyl ring [5]. In addition, given the average bacterial translation error rate of 5 × 10−4 per amino acid [50], wild-type revertants may significantly affect these single-mutants’ analyses. Histidine-containing OAH (as in antibody 6D9; Fig. 2) have not yet been found in nature. The lower efficiency of His in OAS is seen in hydrolytic antibodies (see above). The latter may be due to the relatively low pK a of the His side chain or the more demanding desolvation of the charged His residue compared to the more hydrophobic tyrosine, aspargine, or serine. Interestingly, several structural studies (NMR of trypsin-leupeptin complex [51]; X-ray of proteaseA-chymostatin [52]) identified the oxygen carbonyl of the aldehyde inhibitors pointing toward the active-site’s His, suggesting that the latter has comparable affinity to the oxyanion with respect to the original OAH two backbone NH groups. This observation is in agreement with the OAS efficiency scale postulated from the catalytic antibodies’ data. The estimated values for the individual residue’s contribution to OAS are generally lower for enzymes (Table 3) than for the D-antibodies and most other esterolytic antibodies (Tables 1 and 2). It may be that the enzymes’ mutagenic studies underestimate the OAH side chain’s contribution, as the remaining H-bond by the backbone NH at the OAH is strengthened in the absence of its side-chain partner, hence compensating for its absence. Such compensation may explain the mild effect of 48G7 His35HGlu mutation (40-fold reduction in catalytic rate), compared to 6D9 His27dAla mutation. The latter lacks a backbone NH, which is present in 48G7’s OAH (Table 2). The only natural OAH suggested to have a similar contribution to catalysis as that of the antibodies described above is of AChE, which has three hydrogen donors (Fig. 3) (compared to 1–2 in most antibodies). Did enzyme evolution miss out on an efficient OAS strategy that is easily implemented by antibodies? Few plausible answers come to mind. The OAH may not have evolved independently in nature, but paralleled (or followed) the appearance of a catalytic triad that takes care of nucleophilic and proton-transfer catalysis (Fig. 1b). Once the latter was established,

2 Chapter Overview Table 3 Contribution of oxyanion stabilization to enzymes, mutagenesis studies

Enzyme

OAH

Subtilisin Asn155, Thr220, (Ser221)

CarboxypeptidaseA Trypsin

Gln19

(Ser42) Prolyloligopeptidase Oligo-peptidaseB CocE

AChE

Arg127 (Gly193) (Ser195) (Ser195) Gln19Ala (Cys25) Ser42 Ser42Ala (Gln121) Tyr473 (Asn555) Tyr452 (?)e) Tyr44 (Tyr114) (Gly118) (Gly119) (Ala201)

OASa) ‡ G cat−uncat Kcal/mol

Ref.

Asn155Gly Asn155Ala Thr220Ala Asn155Ala Thr220Ala Ser221Alab) Arg127Alac) Ser195Ala

3 4.2 1.8 6.0

[4] [130] [130] [130]

4.8 6.0 4.2

[4] [131] [45]

3

[132]

3.6

[133]

Tyr473Phe

3.7

[43, 47]

Tyr452Phe Tyr44Phe

3.6 > 4.4d)

[48] [5]

-f)

5–7

[46]

Mutation

a) Contribution to oxyanion stabilization derived from OAH-residue mutation’s effect on catalytic rates or: b) Contribution to oxyanion stabilization derived from residual activity after mutation of the enzyme’s endogenous nucleophile. c) Though Thr220 is only within 4.0 Å of the oxyanion, dynamic simulations suggest this residue’s participation in subtilisin’s OAS. d) Limited by detection of mutant’s activity. Significant structural perturbation of mutant’s active site e) As crystal structure is unavailable, the identity of an assumed second OAH residue (analogous to the homologous Asn555 of prolyloligopeptidase (see separate entry) is unknown. f) Estimated by the authors based on AChE-TSA crystal structure and accumulative data.

together with other mechanisms such as substrate destabilization [46] [52], the hydrolytic activity may have been sufficiently high (diffusion-limited at times [46]) to make further improvements in OAS unnecessary. Alternatively, it was recently suggested that the charged imidazole of the subtilisin triad’s histidine has a major role of lowering the pK a of the TS oxyanion, thereby diminishing the OAH binding energy [40]. In such a scenario, the major role of the OAH is substrate alignment rather than TS stabilization as was initially suggested [53]. It remains to be seen whether these results would extend to other natural enzymes.

11

12 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

In conclusion, hydrolytic antibodies validate the fundamental role of OAS in enzyme catalysis. They do so with similar or perhaps better efficiency than enzymes. OAS measured for antibodies could be thus used as an upper limit to the contribution of OAS in natural hydrolytic enzymes, providing a reference value for the contribution of their other mechanistic features (see below).

2.1.5 Antibody Affinity and Rate Acceleration Limitation Despite the limitations of TSA design, our analysis suggests that many antibodies do convert TSA binding energies to catalysis with nearly 100% efficiency (Table 1). This may suggest a facile route to increased catalytic efficiency via antibodies with higher affinity. However, it appears that antibodies with affinities higher than 1 nM may not be easily obtained. Data collected on the affinity of a large number of antibodies [54] suggest an antibody of 0.1–1 nM [55]. This is derived from the upper range for the association rate (< 5 × 10 M−1 s−1 ) and a dissociation rate (koff ) that is sufficient to trigger B-cell activation (10−3 –10−4 s−1 ). These studies suggest (see [54, 55] and references therein) that 0.1–1nM affinities are sufficient for mounting an efficient immunological response, arguing that the immune system is unlikely to produce higher affinity antibodies. By inference, with a low-average K M (≈KS ) in the mM range [56–58] a “catalytic ceiling” of K TS /KS = kcat /kuncat = 106 − 107 may be derived for catalytic antibodies raised against TSA haptens even if these happen to perfectly mimic the reaction TSs. Thus, the turnover number (kcat ) of antibody-mediated hydrolysis of esters commonly targeted by catalytic antibodies (kuncat 10−6 –10−5 s−1 ) is expected to be well under 10s−1 and the specificity constant (kcat /K M ) ≤ 104 M−1 s−1 . Under the immunological constraints limiting K TSA , a lower K M (or higher substrate affinity) would result in a lower rate acceleration, without lowering the specificity constant. Indeed, a survey of the published catalytic antibodies conforms with the above theoretical limitation (Fig. 4). Moreover, in some instances, not all of the TSA binding energy is directed towards TS stabilization, but rather to non-contributing epitopes of the TSA hapten. For example, antibody 48G7 was shown to bind its TSA with 30 000-fold higher affinity compared to its germline counterpart, resulting in only ∼80-fold improvement of kcat /K M [16]. Although independent values of kcat and K M were not reported, it is likely that a significant part of the increase in binding energy upon maturation was directed to both the substrate and the TSA, and therefore led to no increase in rate acceleration. Similarly, attempts to improve the rate of esterolytic antibodies by directed evolution aimed toward higher TSA affinities have shown that major increases in affinity may result in minor rate improvements [59, 60]. Improving the rates of esterolytic antibodies is not a purely academic challenge. The limitations in rate hinder at least two of the most promising biomedical applications of hydrolytic antibodies – cocaine detoxification [61] and pro-drug therapy [62]. Catalytic forces other than OAS may help in closing the gap of ∼100-fold reactivity [5, 62] required for their successful implementation. These are discussed below.

2 Chapter Overview

Fig. 4 Rate enhancements exhibited by catalytic antibodies: a compilation of rate accelerations exhibited by all catalytic antibodies [139]. The theoretical threshold is set by the affinity ceiling of antibodies as described

in the text. The only antibody to surpass the theoretical threshold dictated by the immune system is 38C2 [140], achieved by reactive immunization (see text for details).

2.1.6 Nucleophilic Catalysis The vast majority of esterolytic antibodies make use of a solution (exogenous) hydroxide ion to generate the tetrahedral, oxyanionic intermediate that leads to ester hydrolysis. This is not particularly effective, as at or close to neutral pH, hydroxide ion concentrations are very low. The enzymatic solutions to this problem are basically two: (i) nucleophilic catalysis by an active-site residue (an endogenous nucleophile) via an acyl-enzyme intermediate; (ii) a general-base residue and a bound water molecule leading to the in situ generation of an appropriately positioned hydroxide ion. Inspired by natural hydrolytic enzymes that utilize a covalent catalytic mechanism (Fig. 1b), and with the goal of catalyzing the more demanding reaction of amide hydrolysis, several attempts were made to introduce an endogenous nucleophile into antibodies’ combining sites. But, in fact, the more effective cases of endogenous nucleophilic catalysis seem to be in antibodies in which the endogenous nucleophile was incorporated haphazardly. Antibodies in which endogenous nucleophilic catalysis plays a role are discussed in Section 2.2 below. In the same vein, attempts were made to generate antibodies in which the nucleophilic hydroxide is generated in situ by general-base catalysis (discussed in Section 2.4 below). Alternatively, the efficiency of the exogenous nucleophilic attack can be enhanced by applying nucleophiles more potent than hydroxide (“chemical rescue”; Section 2.3). 2.2 Endogenous Nucleophiles in Hydrolytic Antibodies 2.2.1 Reactive immunization (RI) Several attempts were made to introduce endogenous nucleophiles into antibody combining sites, either by chemical modification [9, 63] or by immunization. The

13

14 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Scheme 1

latter was aimed at “reactive immunization”, whereby the labile phosphonate diester 1 (Scheme 1), a known covalent inhibitor of hydrolases, was used as a hapten [7, 64]. During the immunization process, an RI hapten should covalently trap antibodies with a nucleophilic residue, thereby forming a highly stable complex that is selectable by the immune system. There is, however, a major caveat to this methodology. Hydrolysis of the activated p-methylsulfonate ester substrate via nucleophilic catalysis will result in an that is significantly more stable than the original substrate. For effective catalysis to occur, an would be necessary to activate the hydrolysis of the acyl-enzyme intermediate (Fig. 1b). Such a residue is counter-selected in this system, as it would also hydrolyze the covalent phosphonyl hapten-antibody bond, thus eliminating the selectable advantage of the reactive immunization. Moreover, hapten 1 is labile (estimated half-life mice’s serum is 24 h), resulting in a mixture of haptens, 1 and 2, (the bi- and mono-phosphonate esters, respectively). Unexpectedly, the affinity to the “baiting” hapten 1 is rather low (in the micromolar range) compared to the high nanomolar affinity of antibodies raised against hapten 2. The kinetic parameters of anti-1 antibodies, suggest that OAS (represented by hapten 2) does not account for the RI-derived antibodies’ catalytic efficiencies (e.g., for antibody 15G12, K TSA(2) /K M = 2; kcat /kuncat = 6.6 × 105 ). However, there is no other evidence to support nucleophilic catalysis in the hydrolytic antibodies raised against RI hapten 1 (e.g., burst kinetics, trapping of the covalent intermediate, non-competitive inhibition by hapten 1). Some of the RI antibodies were reportedto catalyze the dephosphonylation of their cognate hapten 1, albeit with a low turnover (< 3), because of product inhibition [7] (such single turnover rate enhancement by

2 Chapter Overview

ester-hydrolytic antibodies, raised against the ground state hapten, was reported as early as 1980 [65]). Schowen has recently suggested that TS flexibility, where the classical bipyramidal dephosphonylation TS may adopt a more tetrahedral-like conformation, renders it available for stabilization by the anti-phosphonate antibodies [66]. It should be mentioned that, although RI achieved a limited success with ester hydrolysis, it has proven highly effective for reactions such as aldol condensation, retro-aldol and Michael addition reactions that proceed via a covalent Schiff-base enzyme-substrate intermediate [67, 68]. 2.2.2 Serendipity and 43C9 Antibody It has been suggested that several hydrolytic antibodies raised against phosphonate TSAs include an endogenous nucleophile as part of their mechanism of action. Unfortunately, in most cases, only partial evidence is available. In general, proving a nucleophilic mechanism is not a trivial matter, and several independent pieces of evidence need to provided, including the direct identification of an acyl-enzyme intermediate, before a definite statement can be made [2]. A “ping pong” mechanism by antibody PCP21H3 (kcat /kuncat not determined) in a transesterification reaction of a vinyl ester has been suggested. The antibody exhibits burst kinetics for the hydrolysis of a related p-nitrophenyl ester [17]. Product inhibition, common to many catalytic antibodies, was detected as well, yet this does not outweigh the evidence for the covalent mechanism. A similar burst in product formation interpolated to a 1:1 molar ratio of product to antibody was reported for antibody 6F11 (kcat /kuncat = 3.4 × 103 ) [69]. Chemical modification oftyrosyl residues resulted incomplete activity loss. Antibody 20G9 (kcat /kuncat = 500) exhibits “burst kinetics”, and this led to the suggestion of nucleophilic catalysis [70–72]. However, the kinetic analysis is complicated by strong product inhibition, and no evidence apart from the “burst” is available. Incubation of the antibody with N-cbz-glycine -O-phenyl ester resulted in multiple acylation of the antibody and a significant decrease in its activity, yet acylation of an active-site residue such as lysine that is not directly involved in catalysis may account for this result [73]. In addition, the reported acid-limb pH profile fitting a tyrosyl pK a (9.6), as well as the loss of activity when chemically modifying tyrosyl residues with tetranitromethane (TNM), could well be explained by this residue’s participation in OAS [10, 73, 74]. Antibody 7C8 (kcat /kuncat = 3.4 × 103 ) also exhibits a pH profile with an acidlimb with a pK a ≈ 9.0. The rate is also insensitive to the addition of a strong nucle-ophile (hydroxylamine; see also Section 2.3 below). These led to the suggestion of nucleophilic catalysis by an active-site tyrosyl hydroxyl [11]. However, D2.3 and D2.4 antibodies also manifest the above characteristics [10], yet their active site tyrosine (H100d) was shown to serve as an OAH rather than a nucleophile. Similarly, 7C8’s crystal structure reveals a sole residue that may serve as an OAH, Tyr95H. Another clear demonstration of the difficulties associated with proving a nucleophilic mechanism is antibody 17E8. Its crystal structure and hydroxylamine partitioning experiments suggested the existence of a Ser-His dyad, reminiscent of natural serine hydrolases [13, 35]. However, direct evidence of an acyl-enzyme

15

16 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

intermediate was not provided, and the proposed mechanism involved the disruption of a hydrogen bond between His35H and TrpH47 that is conserved in all known antibody structures [36]. Indeed, nucleophilic catalysis was ruled out when a variant of 17E8 in which the Ser, assumed to act as nucleophile, was replaced by Gly exhibited a similar rate enhancement [30]. Most significantly, analysis of the antibodies described above suggests that even if an is indeed part of their mechanism, the resulting rate enhancements are 2–3 orders of magnitude lower than is achieved with antibodies raised against the same haptens and that act by OAS only (see above). Antibody 43C9, raised against the phosphonoamidate 3 (Scheme 1) [75, 76] stands out from all the above examples, not only because of its high reactivity (it catalyzes the hydrolysis of both a p-nitrophenyl ester and a p-nitroanilide and, although its rate enhancement is within the range of other known esterolytic antibodies, its specificity constant is so far unrivalled (kcat /KM = 2.8 × 107 M−1 s−1 )), but also because of the unambiguous assignment of a nucleophilic mechanism. In the case of antibody 43C9, there is ample evidence to prove a nucleophilic mechanism. The pH-rate profile suggests a change in the rate-limiting step: while at pH > 9 product release is limiting, at more acidic pH an acylation step determines the rate [77]. This was demonstrated by pre-steady-state kinetics, where a burst of antibody-equivalent product release kinetics was observed [77]. Finally, the effect of p-substituents of the phenolic leaving group on the rate of catalysis [78] and the identification by electrospray mass spectroscopy of an acyl-enzyme intermediate (with HisL91 as the acylated residue) unambiguously support the proposed mechanism [6]. A detailed catalytic mechanism based on 43C9 crystal structure was put forward where a nucleophilic attack, carried out by HisL91, is accompanied by OAS via the side chains of HisH35 and ArgL96 (Fig. 5) [79]. The serendipitous selection of a His side chain as the nucleophilic catalyst of 43C9 is quite interesting. Not only is the His’s imidazole side chain a better nucleophile toward activated esters than the seryl alcohol moiety, but also its pK a is optimal for catalysis and theresulting acyl-enzyme intermediate is much more labile than an ester intermediate. Thus, His makes an ideal nucleophilic catalyst. Nevertheless, His was never recorded as a nucleophile in natural hydrolytic enzymes. One possible explanation is that with non-activated ester substrates, imidazole prefers acting as a general base rather than a nucleophile, hinting at nature’s preference for other nucleophiles as alcohols and thiols [80]. The OAH of 43C9 is equivalent to other known antibodies (Table 1, Fig. 2), suggesting a comparable contribution to its rate enhancement. Thus, the nucleophilic mechanism does not seem to contribute much to the rate acceleration of ester hydrolysis. This can be explained by the absence of a general base residue (Fig. 1b) that would catalyze the rate-limiting de-acylation step, thus replacing an activated p-nitrophenyl ester with a less active acyl-imidazolium moiety. However, nucleophilic catalysis may contribute significantly to anilide hydrolysis, where formation of the intermediate, rather than its breakdown, is rate-determining. The nucleophilic mechanism may also release 43C9 from having a kcat /K M limited by low affinity to the ground state substrate, as prescribed for efficient catalytic antibodies utilizing solely the energy gained

2 Chapter Overview

Fig. 5 The mechanism of catalysis by antibody 43C9 [79]. (i) A nucleophilic attack by an endogenous nucleophile – HisL91 side chain. The nucleophilic imidazole is maintained in the correct tautomer by a network of H-bonds with TyrL36 and TyrH95 side chains. (ii) The resulting oxyanionic intermediate is stabilized by H-bonding to His35H and ArgL96. Bond rearrangement, coupled with a proton transfer cascade, re-

sults in the acyl intermediate and the release of the aniline leaving group. The aniline release is mediated by protonation by a water molecule (not shown). (iii) An exogenous nucleophilic attack by a hydroxide ion on the acyl-enzyme intermediate. (iv) Bond rearrangement results in the release of the carboxylate product and the free antibody (see hapten 3 structure for R and R ).

from preferential TS vis-a-vis substrate binding, yielding ca. 2 orders of magnitude higher specificity constant, in parity with natural enzymes’ values. Quantifying the individual contribution of the nucleophilic attack is further complicated by the fact that a HisL91Gln mutation of 43C9 results in total loss of hydrolytic activity, yet retains wild-type levels of affinity to its hapten and substrates [81]. Thus, in agreement with natural enzymes, even in a somewhat simplified model, different catalytic mechanisms seem to be coupled. Importantly, the nucleophilic mechanism endows 43C9 with the ability to hydrolyze the more stable p-nitroanilide substrate, unattainable by the antibodies using OAS as their only source of reactivity. Unlike natural amidases, 43C9 does not catalyze the hydrolysis of non-activated amides. The rate-limiting step in their hydrolysis involves the protonation of the amine-leaving group via a general acid residue (as in natural amidases, Fig. 1b), not found in 43C9 active site.

17

18 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Fig. 6 The tetrahedral intermediate formed by various exogenous nucle-ophiles.

2.3 Exogenous Nucleophiles and Chemical Rescue in Hydrolytic Antibodies

The nucleophilic attack of a solution’s (exogenous) hydroxide anion seems to constitute the rate-limiting step of all hydrolytic antibodies. In most cases, the hydroxide reacts directly with the ester substrate and, in rarer cases, with an acyl-antibody intermediate. This step therefore merits careful examination. One of the most useful analytical tools is the replacement of hydroxide by alternative nucleophiles. Negatively charged, tetrahedral intermediates, homologous to the hydroxyl-mediated catalytic mechanism (Fig. 1a) can be formed by the attack on the ester’s carbonyl by other nucleophiles (Fig. 6). Thus, it may be expected that small nucleophiles, stronger than the hydroxyl anion, may be active in the antibodies’ active sites, resulting in higher catalytic rates. A similar strategy was previously used to “chemically rescue” hydrolytic enzymes where the nucleophilic residue was mutated. The mutants’ catalytic activity can often be restored to rates only 100- to 300-fold lower than the intact enzyme [82–84]. One example of effective “chemical rescue” of hydrolytic antibodies is with the peroxide anion, a ∼200-fold stronger nucleophile than hydroxide, that was found to enhance the turnover rates of D2.3 and D2.4 antibodies to a similar extent (Fig. 7a, Table 4) [10]. The concentration-dependent rate acceleration exhibits a rate ceiling at a kcat that is 500- to 2000-fold higher than that of the antibodies’ hydroxidemediated catalysis. The rate ceiling is due to a change in the rate-limiting step observed at high peroxide concentrations – a conformational change becomes rate limiting instead of the nucleophilic attack. The observed rate enhancement of the D-antibodies in the presence of peroxide compares favorably with the estimated contribution of the deacylation rate-limiting step of various natural hydrolases (e.g., ca. 200-fold for cocaine esterase cocE [10, 49]. The D-Abs’ maximal rate of 6 s−1 is also comparable to many natural esterolytic enzymes, such as cocE [5], thioesterases [85], and carboxylesterases [86], and is within the lower range of typical rates of (1–200 s−1 ). This suggests that a potent exogenous nucleophile can match, in the framework of efficient OAS antibodies, the role of the enzyme’s general-base deacylation mechanism (e.g., the His-Asp dyad in Fig. 1b). The secondorder rate for peroxide anion reactivity with the D-antibodies was calculated to be 6 × 105 M−1 s−1 . This is significantly lower than the directly measured second-order rate for water-mediated deacylation of chymotrypsin (< 4M−1 s−1 ) and compares favorably with its deacylation by an exogenous amine nucleophile (tyrosinamide: 1.7 × 104 M−1 s−1 ), a direct result of the enzyme’s specificity for the leaving group

2 Chapter Overview

Fig. 7 The effect of exogenous nucleophiles on the rate of antibody-mediated ester hydrolysis. Rate acceleration factors are the ratios of net initial rates observed in the presence of the nucleophile at a given con-

centration and in its absence. (a) Effect of peroxide anion on the hydrolysis of pnitrophenyl ester by D2.3 and D2.4 [10]. (b) Effect of hydroxylamine on chloramphenicol ester hydroysis (data derived from [11]).

Table 4 Nucleophile effect on D-Abs’ catalysis

Nucleophilea) (× 106 M−1 )b) − OH −O

2H

NH2 NH2 NH2 OH N3 −

pK a) (× 106 M−1 )b)

Buffer (× 106 M−1 )b)

Antibody D2.3

Antibody D2.4

15.4 11.6 8.1 6.0 4.0

13 2800 0.24 0.45 1 × 10−3

12 1270 0 0 n.d.

14 3200 0.008 0.006 0.01 × 10−3

a) A linear correlation links the nucleophilicity of most nucleophiles to their pK a s. However, the reactivity of the α-nucleophiles cited above (i.e. excluding OH− ) is far higher than predicted from their pK a (e.g. 103 -fold for hydroxylamine). b) Results are given in “molar reactivity” – the rate acceleration factor at 1 M nucleophile as depicted in [10].

19

20 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Scheme 2

[87]. Importantly though, the D-Abs’ rate accelerations (kcat /kuncat ) are relatively unchanged when peroxide replaces hydroxide, suggesting that the D-Abs are equally complementary to the respective TSs of both hydroxide and peroxide. The D-Abs exhibit absolute preference toward small, negatively charged nucleophiles (i.e., hydroxylate and peroxylate) for the displacement of the ester’s alcohol moiety, whilst other nucleophiles (e.g., hydroxylamine) do not affect the rate (Table 4). We believe that this specificity is a result of (i) the selection, by the negatively charged phosphonate anion, of antibodies with a cluster of positively charged residues, creating a channel to the antibody active site, and (ii) the snug fit of the antibody to its hapten and therefore to the TS, preventing the entry of bulky nucleophiles such as azide [10]. These results suggest that it may be possible to optimize the electrostatic properties of the channel through which the nucleophile approaches the active site, perhaps by engineering positively charged residues in and around this channel without interfering with the properties of the active-site core [88], thereby increasing the rate of catalysis. The rate of hydrolysis by antibody 6D9 (raised against hapten 4, Scheme 2) is also increased by the presence of the α-nucleophile hydroxylamine. Unlike the D-abs, both the rate (kcat , or kcat /K M ) and the rate acceleration (kcat /kuncat ) increase, albeit to a minor extent (up to ∼2 fold at 100 mM hydroxylamine) (Fig. 7b) [11]. The source of recognition of hydroxylamine by 6D9 was not addressed, yet the Lys32L residue [11] is located in a position suitable for general-base activation of the nucleophile. Alternatively, it may act as an H-bond donor to orient the nucleophile toward attacking the chloramphenicol ester’s carbonyl. It would thus be interesting to study the effect of Lys32L mutants on the hydroxylamine-mediated reaction, as well as attempting the hydrolysis of the more demanding amide hydrolysis in this system. Another reported case of “chemical rescue” concerns antibody 14–10. Using the heterologous immunization approach, where mice were challenged with both haptens 5 and 6 (Scheme 2), Masamune and coworkers attempted the generation of a p-nitroanilide hydrolyzing antibody that makes use of OAS, general-base catalysis

2 Chapter Overview

Scheme 3

(by “baiting” with a positively-charged amine hapten) and an exogenous phenol nucleophile. The best antibody achieved, 14–10, catalyzed both the lactonization of substrate 7 (Scheme 3) via an intramolecular phenol nucleophilic attack (kcat = 1.5 × 10−2 min−1 , K M = 1.27 mM [89]) and the intermolecular attack of added phenol nucleophile to substrate 8 to produce phenyl propionate, 9 (kcat = 1.3 × 10−4 min−1 , K M(2) = 0.37 mM, K M(phenol) = 0.14 mM [12]). In both cases the phenol does not seem to participate in the background buffercatalyzed reactions. Thus, the antibody catalyzes kinetically “disfavored” reactions, resulting in kcat /kuncat = 2 × 104 and 630 for the intramolecular and intermolecular reactions, respectively (the uncatalyzed rates serving as an upper limit of the background rates). Interestingly, 14–10 does not hydrolyze the ester product 9. This could be only partly explained by the lack of a negatively charged TSA to elicit an OAH, as similar, uncharged haptens (e.g., phosphono-amidates [62], and β-amino alcohols [90]) were shown to elicit efficient hydrolytic antibodies. Nonetheless, as the buffer-mediated hydrolysis rate of 9 proceeds twice as fast as the antibodymediated acyl transfer, the net reaction can be formally considered as the activated amide’s hydrolysis. Future studies may reveal whether the phenol nucleophilicity is indeed enhanced by a general-base residue, as implied by 14–10’s pH optimum (8.0 compared with the phenol’s solution pK a of 10). 2.4 General Acid/Base Mechanisms in Hydrolytic Antibodies

The most notable example of hydrolytic enzymes that make use of general acid and base catalysis is aspartic peptidases. This family of hydrolases utilizes the concerted action of two carboxylate residues (Fig. 1c). The first carboxylate serves as a general base to align and activate the water nucleophile, and subsequently as a general acid to protonate the leaving group. The second serves as the OAH and may be regarded as a general acid, donating a proton to the formed carboxylate. In order to mimic such mechanisms with catalytic antibodies, several “bait and switch”

21

22 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes Table 5 Kinetic parameters of “bait and switch”-raised antibodies

Antibody

Hapten

kcat a) (min−1 ) pH = 7.8

27A6 30C8 2D5

10 11 12

0.004 0.005 3.5

K M (mM)

kcat /KM M−1 s−1 )

kcat /kuncat b)

Ref.

0.25 1.1 0.5

0.7 0.07 120

1.4 × 105 1.2 × 105 1.3 × 105

[134] [90] [69]

a)

Values for antibodies 30C8 and 27A6 were derived from pH-rate profiles. Although the same substrate and reaction are involved, a ∼1000-fold difference exists between the uncatalyzed rates reported of one group [134] and those of the other [69]. b)

haptens were devised with the aim of “baiting” oppositely charged residues in the antibody’s active site that may serve in a general acid/base mechanism. An example of this strategy (haptens 10–12) and the kinetic rates of the resulting antibodies is summarized in Table 5. Haptens 10–12 (Scheme 4) were designed to bait different types of residues in the elicited antibodies’ binding sites. The positively-charged hapten 11 was designed to bait a general-base residue (e.g., a negatively-charged carboxylate), whereas hapten 10 aims at baiting a general-acid residue that would facilitate the rate-determining protonation of the amine-leaving group of substrate 14. Nonetheless, the catalytic activity of the best antibodies raised against haptens 10 and 11 is remarkably similar (Table 5), suggesting that even if “baiting” did occur, it does not effect to a significant extent the antibodies’ catalytic efficiency. Moreover, whilst the antibodies raised against haptens 10–12 all catalyze the hydrolysis of the ester substrate, 13, with similar kinetic parameters, the amide substrate 14 is hydrolyzed by none

Scheme 4

2 Chapter Overview

of these antibodies, not even by those antibodies raised against hapten 12. The effectiveness of the “bait and switch” strategy in raising hydrolytic antibodies is further questioned by the fact that hapten 12 was designed to bait a general-base catalyst while maintaining the phosphonate tetrahedral negatively charged TSA core for the generation of OAH residues. On the other hand, in haptens 10 and 11, a planar, uncharged carbon represents the tetrahedral TS. The catalytic rate of the best anti-12 antibody, 2D5, is also in agreement with rates of the above-described OAS antibodies, suggesting that even if an active-site general base has been generated, its contribution to catalysis is not very significant. By comparing the catalytic rates of antibodies 30C6 and 27A6 to 2D5, one could conclude that the tetrahedral phosphonate is a better mimic of the TSs of ester hydrolysis than the β-alcohol moiety, or, in other words, the phosphonate TSA “baits” a ∼500-fold more efficient OAH. Other attempts to utilise the “bait and switch” strategy yielded similar rate enhancements to the examples described above, with kcat /kuncat values in the range 102 − 104 (see, for example [91–93]). Future studies of antibodies raised by the “bait and switch” strategy may shed more light on their mechanism and may allow an improvement in their rates. Notably, as is the case with reactive immunization, the “bait and switch” approach was applied to non-hydrolytic reactions with greater success. This is evident, for example, in the Glu residue acting as a general base in antibody 4B2’s mediated catalysis of an allylic rearrangement reaction (PDB code 1F3D [94]). In contrast, the crystal structure of antibody 1D4, raised against a charged β-amino phosphate, does not show the expected positioning of a catalyzing general-base residue in its active site [95]. Finally, it is encouraging to note that a structural model of the chemiluminescent anti-phosphonate 7F11 antibody (kcat /kuncat = 1.2 × 103 , kcat /KM = 5.9 × 105 M−1 s−1 [96]) indicates the presence of two aspartic acid residues in positions resembling the natural aspartyl protease. Based on this model, the authors suggested that the 7F11 mechanism is similar to the natural counterpart, but that its much inferior catalytic efficiency may be due to the imperfect positioning of the apartyl residues. Antibody 7F11 also suffers from severe product inhibition, enabling an estimated turnover of ca. 5 cycles [97]. Nevertheless, if its active-site structure does prove similar to its natural counterpart, it may serve as a starting point for the generation of more potent hydrolases. 2.5 Metal-Activated Catalytic Antibodies

Several studies have attempted to raise catalytic antibodies with metal ion-mediated hydrolysis, following the zinc-protease mechanism (Fig. 1d). Many enzyme models based on zinc have been created that exhibit high rate enhancements [98], suggesting that the generation of metallo-antibodies with enzymatic activities is a promising avenue. However, this approach has thus far been explored with limited success. Lerner and coworkers reckoned that the “bait and switch” hapten 11 may elicit a carboxylate residue in the antibodies’ active site that would serve as a metal-binding ligand. Furthermore, the pyridinium’s methyl group will induce a

23

24 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Scheme 5

cavity for the metal ion. Indeed, in the presence of Zn+ , antibody 84A3 catalyzes with high specificity the hydrolysis of a pyridinium ester (kcat /kuncat = 1200 [99]), the structure of which is related to the immunizing hapten. No catalysis was observed in the absence of the metal ion or with other, structurally similar ester substrates. The antibody suffers from severe substrate inhibition, probably due to non-productive binding of the substrate, probably in a mode not involving a zinc ion, as judged by the affinity difference to the substrate in the presence or absence of Zn+ (3.5 µM and 840 µM, respectively). In an attempt to overcome this hurdle, the mercury(II)-phosphorodithioate unsaturated complex 15 (Scheme 5) was used to elicit Hg-dependent hydrolysis of ester 16 [100]. The best anti-15 antibody, 38G2, exhibited Michaelis-Menten kinetics (kcat /kuncat = 300, K M(Hg) = 87 µM and K M(16) = 345 µM), multiple turnovers and absolute dependency on Hg(II) for catalysis. While far from enzyme-like reactivity, these studies widen the spectra of possible mechanisms that could be tailored by antibody mimics. 2.6 Substrate Destabilization

Remote interactions, away from the ester or amide carbonyl reaction core, play an important role in many natural enzymes. Such interactions may contribute to catalysis via substrate destabilization (for a recent review see [101] and references therein). For example, Sussman and coworkers postulated that as much as 5 × 105 (corresponding to TS stabilization of 8 kcal/mol) of AchE’s spectacular rate enhancement (1013 ) is derived from orienting the substrate to a highly unfavorable conformation, perfectly aligning the substrate for the nucleophilic attack to take place. This is achieved by strong binding to the choline ammonium charged moiety [46]. Several examples of the identification and possible role of substrate destabilization by hydrolytic antibodies are given below. The docking of the activated anilide substrate in the crystal structure of 43C9 suggests that the substrate’s amide bond is twisted by 140◦ out of planarity, thus destabilizing the bond’s resonance and facilitating its hydrolysis [79]. Similarly, 6D9 antibody’s structure suggests that the antibody locks its ester substrate in the sin unfavorable conformation, resulting in substrate destabilization. In both the above examples, a quantitative assessment of the contribution of substrate destabilization to the rate acceleration is not available. A 5-fold increase in the catalytic rate accompanied by a smaller effect of higher K M (2-fold) was seen between the catalysis of the glutaryl-glycine nitrophenylester

3 The Role of Conformational Changes in Catalytic Antibodies

substrate and a caproyl-elongated substrate by antibody D2.3, suggesting a remote effect of the substrate’s linker moiety. Crystal structure analysis revealed different interactions with the two linkers that may account for the kinetic differences [102]. When the linker terminal glycine was replaced by either the D- or L-Ala enantiomers, a ∼20-fold difference in D2.3’s specificity constant (kcat /K M ) was observed in favor of the l-Ala enantiomer [103] (for the enantioselectivity potential of the esterolytic active sites of various antibodies see [39]). Another example is set by antibody 17E8 catalyzing the hydrolysis of n-formylnorleucine (nLeu) phenyl ester. The specificity factors (kcat /K M ) observed with a series of substrates where the nLeu side chain was replaced by different linear and branched alkyl side chains extend over 2 orders of magnitude. But rate enhancements higher than those with the original substrate (namely with a Leu side chain) were not obtained. These results suggested that 17E8 interactions with the side chain are used to stabilise both the TS and the ground state. However, the replacement of nLeu side chain with a more hydrophilic ethyl-cysteyl side chain resulted in a 20-fold increase in kcat (43 vs 2.1 s−1 ) accompanied by a similar decrease in ground state binding (K M = 4 vs 0.18 mM with nLeu). This rendered the new substrate as efficient as the original substrate (kcat /K M ≈ 1.2 × 104 M−1 s−1 ) and suggested that substrate destabilization may play a key role in the mechanism of 17E8 with the ethyl-cysteyl substrate [104].

3 The Role of Conformational Changes in Catalytic Antibodies

Structural dynamics are part of the nature of proteins, though their identification on the molecular level and their mechanistic roles are still a subject of much research. In enzymes, the identification of conformational changes is generally elusive, as they are typically much faster than the chemical step and product release [2]. Nonetheless, structural fluctuations were observed in serine proteases when comparing their free and boronic acid inhibitor-complexed structures. Such plasticity may be an integral part of the catalytic cycle, mediating, amongst other steps, substrate binding, TS formation, stabilization, and product release (for a recent example see [105, 106]). Several hydrolytic enzymes were also shown to utilize an induced-fit mechanism to tune their specificity, either to a broad (e.g., alpha lytic protease) or extremely narrow (e.g., factor D) substrate spectrum (see [101] and reference therein). In addition, a recent kinetic study suggests that conformational changes control the aperture of AChE’s active site and are coupled to its catalytic activity [107, 108]. There had been several speculations regarding the role and effect that conformational changes may have on catalysis by antibodies. Padlan, for example, speculated that the of antibodies may hinder their catalytic efficiency [109], whilst Benkovic and Stewart suggested that antibodies elicited against TSA might be limited by a “lock-and-key” rigidity and will lack the conformational isomerism that characterizes enzymatic catalytic cycles [20]. In a more general context, the issue of

25

26 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Fig. 8. Schematic representation of the two mechanisms of conformational changes linked to antibody-ligand complex formation. Ab and Ab’ represent two antibody isomers exhibiting low and high affinity, respectively.

conformational isomerism in antibodies has been a subject of much research and debate. In the past, kinetic (e.g., [110, 111]) and structural (e.g., [112] and references therein) studies provided strong evidence of the dominance of structural changes in antibodies. Conformational changes can be generally described by the inducedfit model (the ligand binding a low-affinity form of the antibody and inducing a subsequent conformational change that leads to a high affinity complex) or by preequilibrium between two different conformational species of the same antibody, only one of which binds the ligand with high affinity. The two models can be combined in one scheme (Fig. 8). While kinetic evidence supports the existence of both mechanisms, structural evidence has been put forward thus far for induced fit only. However, a recent study combining pre-steady-state kinetics in solution and X-ray crystallography provides conclusive evidence for pre-equilibrium in an antibody [113]. The identification of conformational changes in catalytic antibodies and their possible role in catalysis are discussed in this section.

4 Conformational changes in catalytic antibodies

The advent of catalytic antibodies has brought a renewed interest in antibody structure and evolution. In particular, structures of catalytic antibodies underlined the understanding of their mechanism at the molecular level (over 50 published structures of catalytic antibodies in PDB as of end of 2002). As is the case with many ordinary antibodies, conformational changes are observed with catalytic antibodies as well – in several catalytic antibodies, the structure of the TSA-bound antibody differs from the uncomplexed structure. But, in the absence of kinetic studies in solution, the interpretation of the crystallographic data regarding the mechanism of these con-formational changes is ambiguous. Further complications from the comparison of bound vs unbound antibody crystal structures arise from the effect of packing forces (e.g., the antibodies’ arrangement in the crystal in antibody 1D4 [95], CDR1 packed inside the active site of antibody 43C9, blocking the diffusion of the hapten [79]), and the buffer used for crystallization (e.g., antibody D2.3, see below and [15]). A recurrent structurally observed conformational change is a relative movement between the heavy- and light-chains’ variable regions [114], as exemplified by antibody CNJ206 where a 7◦ rotation and a ∼1 Å translation change were found between the TSA unbound and bound structures. In addition, a distinct isomerization was noted in the active site, where a dramatic change of the entire H3 CDR loop – from an open state to a closed state – is observed.

4 Conformational changes in catalytic antibodies

This changes the binding site completely – from a shallow form in uncomplexed antibody to a deep and narrow cavity in the TSA-bound state. This is assumed to be a TSA-induced conformational change, but in fact the TSA-bound conformation may be pre-existing, as neither the oxyanion hole nor the part of the active site that binds the phenolic leaving group is formed in the free form [14]. Induced-fit movement of CDR-H3 residues (H99-H102) was also suggested to enhance surface complementarity to the TSA vs the substrate and product of the catalyzed retroDiels-Alder reaction, thus implying that better rate enhancement is gained by the conformational changes (antibody 10F11, [19]). Crystal structures of antibody 1D4, catalyzing a syn elimination reaction, indicated conformational changes limited to CDR-H2. The conformational change increases the contacts with the TSA (notably residue Arg58), and may contribute to catalysis by constraining the substrate to an elimination-favorable orientation [95]. The free and TSA-complexed structures of several catalytic antibodies were solved, alongside the structures the putative germline antibodies from which these catatylic antibodies diverged [27, 115–117]. A general trend has been suggested, whereby the germline antibodies exhibit a different free and bound structure, whilst the mature antibodies exhibit the same structure. The observed isomerizations between the free and bound antibodies seem to occur mainly at CDR3H residues. In antibody 48G7’s precursor, tyrosines H98 and H99 shift by up to 5 Å upon TSA binding together, with a 4.5◦ 4 change in the V-regions’ orientation, resulting in better packing of the TSA’s p-nitrophenyl ring by TyrH99 [27, 117]. The hapten of antibody AZ-28, catalyzing an oxy-Cope reaction, induces a rotation of CDR-H3 away from the binding site to yield better π–π stacking with the hapten’s phenyl ring as well as enhanced orbital overlap to the reaction’s TS [115]. In the precursor of the oxidating, periodate-dependent antibody 28B4, better packing of the TSA phenyl ring is also achieved by movement of residues H95–H99 [116]. As judged by crystallography, the matured antibodies related to these germline antibodies exhibit the same structure when free and bound to the TSA. Interestingly, in antibodies 48G7 and 28B4, the TSA’s p-nitrophenyl ring adopts a different conformation in the mature vs the germline-complexed structures, locking them in a different binding pocket. This is surprising, as, in the case of the mature 48G7, the germline’s pocket remains unchanged. The above studies [118] promoted the hypothesis that the germline antibodies exhibit high conformational diversity, and that, during affinity maturation, the hapten binding conformation becomes dominant [110]. However, structural differences between the bound and the free state are commonly observed in mature antibodies ([112] and references therein), and at least one study has shown similar structures of the mature Diels-Alderase antibody 39-A11 and its putative germline precursor in both bound and unbound states [119]. The crystallographic studies were not accompanied by kinetics in solution that indicate the existence of a pre-equilibrium between different unbound conformations, in either the mature or the germline antibodies. Indeed, a kinetic study of mature antibodies that showed no structural differences between their free and TSA-bound states clearly indicated a pre-equilibrium between two antibody conformers, one exhibiting low affinity

27

28 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

toward the TSA and another high affinity and catalytic activity. The slow, minutescale isomerization between the two conformers resulted in hysteretic, “memorylike” behavior [15]. Moreover, it has also been shown that the crystallization buffer shifts the equilibrium toward the active conformer, resulting in both the free and TSA-bound antibodies yielding the same structure [15]. Also of interest are the mutations that lead to affinity maturation and their effect on TSA binding and rate of catalysis. In antibodies 48G7 and AZ-28 the mutations occur away from the active site. The importance of remote residues on catalytic efficiency was also noted in mutants of antibody 17E8 selected by phage display for better TSA binding. A double mutant was achieved, with ten-fold improvement of kcat /K M [120]. In antibody 28B4, however, three of the somatic mutations (nine in total), are indirect contact with the hapten [116]. In all three antibodies, somatic mutations increased the TSA-binding affinity (up to 40 000-fold in 48G7 [117]). Interestingly, the catalytic activity of the mature 48G7 was increased compared to its germline counterpart (see Section 2.1), whereas that of antibody AZ-28 was diminished, hinting at the role of structural flexibility in this antibody’s catalytic activity (the catalytic activity of 28B4 germline was not reported). Structural flexibility may also explain the increase in the catalytic activity of the ArgH100aGln mutant of antibody 43C9 (a CDR-H3 mutation remote from the active site), which reduces product inhibition [81]. 4.1 The contribution of structural isomerization to catalysis

The structural studies described above do not indicate a direct role for conformational changes in antibody catalysis, nor do they provide a measure of their contribution to TS stabilization and rate of catalysis. In the first quantitative study of conformational changes in catalytic antibodies, two independent kinetic measurements (one related to TSA binding and another directly to catalysis) performed with the D-Abs (D2.3, D2.4, D2.5) were used to reveal and quantify the contribution of conformational changes to catalysis [10, 15]. Pre-steady-state TSA-binding kinetics of D2.3, D2.4 and D2.5 antibodies, based on the change of intrinsic tryptophan fluorescence upon hapten binding, indicated the formation of a low micromolar affinity primary complex followed by a conformational change toward the final nanomolar affinity complex (Fig. 9a,b and Table 6 [15]). The rate of isomerization (6 s−1 ) revealed by fluorescence quenching measurements with both the 1b and 1p TSAs is significantly higher than the rate of hydrolysis of the D-Abs (0.05 – 0.22 s−1 ) and is therefore invisible in the catalytic, productrelease assay. However by increasing the turnover rate by means of a strongly reactive nucleophile (i.e., peroxide anion, see Section 2.3 above), a “rate ceiling” equal to the isomerization rate was observed (Fig. 9c) [10]. The isomerization provides 25 to 140fold higher affinity to the TSA (Table 6) and a concomitant increase to the D-Abs catalytic rates. This is yet another manifestation of the true representation of the actual TS by the TS analog. The “induced-fit” isomerization contributes not only to rate enhancement but also to specificity. A second,

5 Conclusions

Fig. 9 Induced fit exhibited by the Dantibodies. (a) Two-step kinetics reflected by the stopped-flow trace of fluorescence quenching of antibody D2.4 in the presence of TSA. The fast phase (shaded) is enlarged in the inset. Each phase was fitted to a single exponential rate equation. (b) The effect

of TSA concentration on the slow bimolecular binding step and on the fast isomerization step [15]. (c) D2.4 (full circles) and D2.3 (empty circles) catalytic rates dependency on the peroxide anion exogenous nucleophile’s concentration [10].

minute-scale, isomerization step is unique to antibody D2.3 [15]. This very slow conformational change increases the affinity to the TSA 1b antigen ∼7 fold yet decreases its affinity to TSA 1p by ∼2 fold (Table 6). These results may suggest that the immune system may select high-affinity and high-specificity antibodies that still exhibit a considerable degree of conformational change, and that affinity maturation does not necessarily involve a rigid “lock-and-key” binding mechanism [27, 37].

5 Conclusions

The examples given in this review emphasize the ability of catalytic antibodies to mimic multiple aspects of enzyme catalysis. In addition, several other enzyme characteristics are faithfully mimicked by hydrolytic antibodies but are beyond the scope of this review. Hydrolytic antibodies can exhibit both a fine substrate specificity that rivals natural enzymes (e.g., 2H6 and 21H3 [121], the D-Abs [25]

29

0.24 ± 0.02 0.20 ± 0.02 0.15 ± 0.01 0.3 ± 0.03 0.20 ± 0.03 0.20 ± 0.02 0.6 0.7 0.1 0.6 0.4 0.5

5±1 6±1 1 ± 0.1 6±1 7±2 4±1

0.2 ± 0.1 0.10 ± 0.05 0.01 ± 0.003 0.04 ± 0.01 0.2 ± 0.1 0.08 ± 0.03

Kfast+ s−1b) kslow+ s−1 kslow0.04 0.02 0.01 0.007 0.03 0.02

0.013 ± 0.002 0.001 ± 3 × 104 – – – –

Kslow s−1 kslowest+ s−1

0.002 ± 3 × 104 0.0023 ± 3 × 104 – – – –

kslowest-

0.15 2.3 – – – –

3±1 25 ± 3 1 ± 0.4 4±1 10 ± 2 10 ± 2

Kslowest nM KD c)

b)

Because of high error in its determination kfast+ was back-calculated from the overall dissociation constant (K d ) after measuring all the other kinetic parameters. Calculated from K d . c) Measured by fluorescence equilibrium titration.

a)

4 × 105 3 × 105 2 × 106 5 × 105 5 × 105 4 × 105

TSA µM−1 s−1 kfast+ s−1a) kfast- µM

D2.3 1b 1p D2.4 1b 1p D2.5 1b 1p

Ab

Table 6 Kinetic parameters of D-Abs’ binding to p-nitro-phenyl (1p)- and benzyl (1b)-phosphonate TSAs

30 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

References

and 12F12 [93] antibodies), as well as very broad specificity that may be induced by bulky TSA linker moieties (e.g., [122, 123]). In addition, high enantioselectivity was reported for several ester hydrolytic antibodies (e.g., [124, 125]). Metal (Pd)-mediated allosterity was also described [126]. Rivaling the rates of enzyme catalysis, by catalytic antibodies or any other enzyme mimic, is a daunting challenge. Antibodies, like all other enzyme mimics, are still much inferior to natural enzymes in their catalytic efficiency. Yet, despite the limited rates achievable by hydrolytic antibodies, their generation and the associated research effort have provided unique insights into enzyme catalysis and antibody diversity, evolution and structure. The clear message from 16 years of research into hydrolytic antibodies is that it is possible to mimic individual forces and contributions to enzymic catalysis with enzyme-like efficiencies. However, it is the combination of the different catalytic forces and contributions described here that may yield antibodies with efficiencies rivaling natural enzymes. Moreover, a significant enhancement in rate may require the incorporation of several catalytic elements and a parallel improvement of the antibody’s dynamics. These complex structural and mechanistic elements are not likely to be induced by designing better haptens or engineering existing antibodies. Thus, the most general and effective way of improving the catalytic potency of antibodies would be to select esterolytic antibodies directly for higher rates and turnover ([127] and references therein). Such selections can affect all the factors concerned, including those linked directly with the active site (e.g., inducing endogenous active-site nucleophiles) and those peripheral to it (e.g., improving the approach of exogenous nucleophiles or the rate of conformational isomerism). The existing hydrolytic antibodies described above that exhibit low to medium rate accelerations and effective turnover may serve as the ideal starting point for such an evolutionary process. References 1 TANTILLO, D. J., HOUK, K. N., J. Org. Chem. 64 (1999), p. 3066–3076 2 FERSHT, A. R., Enzyme structure and mechanism, W.H. Freeman, New-York 1985 3 HAMMES, G. G., Biochemistry 41 (2002), p. 8221–8228 4 CARTER, P., WELLS, J. A. Proteins 7 (1990), p. 335–342 5 TURNER, J. M., LARSEN, N. A., BASRAN, A., BARBAS, C. F. 3rd, BRUCE, N. C., WILSON, I. A., LERNER, R. A., Biochemistry 41 (2002), p. 12297–12307 6 KREBS, J. F. SIUZDAK, G. DYSON, H. J. STEWART, J. D., BENKOVIC, S. J., Biochemistry 34 (1995), p. 720–723 7 WIRSCHING, P., ASHLEY, J. A., LO, C. H., JANDA, K. D., LERNER, R. A., Science 270 (1995), p. 1775–1782

8 BALDWIN, E., SCHULTZ, P. G., Science 245 (1989), p. 1104–1107 9 POLLACK, S. J., NAKAYAMA, G. R., SCHULTZ, P. G., Science 242 (1988), p. 1038–1040 10 LINDNER, A. B., KIM, S. H., SCHINDLER, D. G., ESHHAR, Z., TAWFIK, D. S., J. Mol. Biol. 320 (2002), p. 559–572 11 GIGANT, B., TSUMURAYA, T., FUJII, I., KNOSSOW, M., Struct. Fold Des. 7 (1999), p. 1385–1393 12 ERSOY, O. FLECK, R. SINSKEY, A. MASAMUNE, S., J. Am. Chem. Soc. 120 (1998), p. 817–818 13 GUO, J. HUANG, W. SCANLAN, T. S., J. Am. Chem. Soc. 116 (1994), p. 6062–6069 14 CHARBONNIER, J. B., CARPENTER, E., GIGANT, B., GOLINELLI-PIMPANEAU, B., ESHHAR, Z., GREEN, B. S., KNOSSOW, M.,

31

32 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

15 16

17

18

19

20 21

22

23 24

25

26

27

28

29

30

Proc. Natl. Acad. Sci. U S A 92 (1995), p. 11721–11725 LINDNER, A. B., ESHHAR, Z., TAWFIK, D. S., J. Mol. Biol. 285 (1999), p. 421–430 WEDEMAYER, G. J., PATTEN, P. A., WANG, L. H., SCHULTZ, P. G., STEVENS, R. C., Science 276 (1997), p. 1665–1669 WIRSCHING, P., ASHLEY, J. A., BENKOVIC, S. J., JANDA, K. D., LERNER, R. A., Science 252 (1991), p. 680–685 MUNDORFF, E. C., HANSON, M. A., VARVAK, A., ULRICH, H., SCHULTZ, P. G., STEVENS, R. C., Biochemistry 39 (2000), p. 627–632 HUGOT, M., BENSEL, N., VOGEL, M., REYMOND, M. T., STADLER, B., REYMOND, J. L., BAUMANN, U., Proc. Natl. Acad. Sci. U S A 99 (2002), p. 9674–9678 STEWART, J. D., BENKOVIC, S. J., Nature 375 (1995), p. 388–391 JENCKS, W. P., Catalysis in chemistry and enzymology, McGraw-Hill Book Company, New-York 1969 TAWFIK, D. S., CHAP, R., GREEN, B. S., SELA, M., ESHHAR, Z., Proc. Natl. Acad. Sci. U S A 92 (1995), p. 2145–2149 MADER, M. M., BARTLETT, P. A., Chem. Rev. 97 (1997), p. 1281–1302 TAWFIK, D. S., GREEN, B. S., CHAP, R., SELA, M., ESHHAR, Z., Proc. Natl. Acad. Sci. U S A 90 (1993), p. 373–377 TAWFIK, D. S., LINDNER, A. B., CHAP, R., ESHHAR, Z., GREEN, B. S., Eur. J., Biochem. 244 (1997), p. 619–626 CHARBONNIER, J. B., GOLINELLI, P. B., GIGANT, B., TAWFIK, D. S., CHAP, R., SCHINDLER, D. G. KIM, S. H., GREEN, B. S., ESHHAR, Z., KNOSSOW, M., Science 275 (1997), p. 1140–1142 WEDEMAYER, G. J., WANG, L. H., PATTEN, P. A., SCHULTZ, P. G., STEVENS, R. C., J. Mol. Biol. 268 (1997), p. 390–400 PATTEN, P. A., GRAY, N. S., YANG, P. L., MARKS, C. B., WEDEMAYER, G. J., BONIFACE, J. J. STEVENS, R. C. SCHULTZ, P. G., Science 271 (1996), p. 1086–1091 BUCHBINDER, J. L., STEPHENSON, R. C., SCANLAN, T. S., FLETTERICK, R. J., J. Mol. Biol. 282 (1998), p. 1033–1041 GUO, J., HUANG, W., ZHOU, G. W., FLETTERICK, R. J., SCANLAN, T. S., Proc. Natl. Acad. Sci. U S A 92 (1995), p. 1694–1698

31 MIYASHITA, H., KARAKI, Y., KIKUCHI, M., FUJII, I., Proc. Natl. Acad. Sci. U S A 90 (1993), p. 5337–5340 32 MIYASHITA, H., HARA, T., TANIMURA, R., FUKUYAMA, S., CAGNON, C., KOHARA, A., FUJII, I., J. Mol. Biol. 267 (1997), p. 1247–1257 33 O. KRISTENSEN, VASSYLYEV, D. G., TANAKA, F., MORIKAWA, K., FUJII, I., J. Mol. Biol. 281 (1998), p. 501–511 34 R. ZEMEL, SCHINDLER, D. G., TAWFIK, D. S., ESHHAR, Z., GREEN, B. S., Mol. Immunol. 31 (1994), p. 127–137 35 ZHOU, G. W., GUO, J., HUANG, W., FLETTERICK, R. J., SCANLAN, T. S., Science 265 (1994), p. 1059–1064 36 MACBEATH, G., HILVERT, D., Chem. Biol. 3 (1996), p. 433–445 37 TANTILLO, D. J., HOUK, K. N., Chem. Biol. 8 (2001), p. 535–545 38 GOLINELLI-PIMPANEAU, B., J. Immunol. Methods 269 (2002), p. 157–171 39 TANTILLO, D. J., HOUK, K. N., J. Comput. Chem. 23 (2002), p. 84–95 40 O’CONNELL, T., DAY, P. R. M., TORCHILIN, E. V., BACHOVCHIN, W. W., MALTHOUSE, J. G., BioChem. J. 326 (1997), p. 861–866 41 O’SULLIVAN, D. B., O’CONNELL, T. P., MAHON, M. M., KOENIG, A., MILNE, J. J., FITZPATRICK, T. B., MALTHOUSE, J. P., Biochemistry, (1999) 38 p. 6187–6194 42 GERLT, J. A., GASSMAN, P. G., Biochemistry 32 (1993), p. 11943–11952 43 SZELTNER, Z., RENNER, V., POLGAR, L., Protein Sci. 9 (2000), p. 353–360 44 FERSHT, A. R., SHI, J. P., KNILL-JONES, J., LOWE, D. M., WILKINSON, A. J., BLOW, D. M. BRICK, P., CARTER, P., WAYE, M. M., WINTER, G., Nature 314 (1985), p. 235–238 45 COREY, D. R., CRAIK, C. S., J. Am. Chem. Soc. 114 (1992), p. 1784–1790 46 HAREL, M., QUINN, D. M., NAIR, H. K., SILMAN, I., SUSSMAN, J. L., J. Am. Chem. Soc. 118 (1996), p. 2340–2346 47 SZELTNER, Z., REA, D., RENNER, V., FULOP, V., POLGAR, L., J. Biol. Chem. 277 (2002), p. 42613–42622 48 JUHASZ, T., SZELTNER, Z., RENNER, V., POLGAR, L., Biochemistry 41 (2002), p. 4096–4106 49 LARSEN, N. A., TURNER, J. M., STEVENS, J., ROSSER, S. J., BASRAN, A., LERNER, R. A.,

References

50 51

52 53 54

55 56 57 58 59

60

61

62

63 64

65

66 67 68

69

BRUCE, N. C., WILSON, I. A., Nat. Struct. Biol. 9 (2002), p. 17–21 PARKER, J., MicroBiol. Rev. 53 (1989), p. 273–298 ORTIZ, C., TELLIER, C., WILLIAMS, H., STOLOWICH, N. J., SCOTT, A. I., Biochemistry 30 (1991), p. 10026–10034 DELBAERE, L. T., BRAYER, G. D., J. Mol. Biol. 183 (1985), p. 89–103 HENDERSON, R., J. Mol. Biol. 54 (1970), p. 341–354 CHAPPEY, O., DEBRAY, M., NIEL, E., SCHERRMANN, J. M., J. Immunol. Methods 172 (1994), p. 219–225 FOOTE, J., EISEN, H. N., Proc. Natl. Acad. Sci. USA 92 (1995), p. 1254–1256 THOMAS, N. R., Appl. BioChem. Biotechnol. 47 (1994), p. 345–372 THOMAS, N. R., Nat. Prod. Rep. 13 (1996), p. 479–511 STEVENSON, J. D., THOMAS, N. R., Nat. Prod. Rep. 17 (2000), p. 535–577 BACA, M., SCANLAN, T. S., STEPHENSON, R. C., WELLS, J. A., Proc. Natl. Acad. Sci. U S A 94 (1997), p. 10063–10068 TAKAHASHI, N., KAKINUMA, H., LIU, L., NISHI, Y., FUJII, I., Nat. Biotechnol. 19 (2001), p. 563–567 DENG, S. X., de PRADA, P., LANDRY, D. W., J. Immunol Methods 269 (2002), p. 299–310 WENTWORTH, P., DATTA, A., BLAKEY, D., BOYLE, T., PARTRIDGE, L. J., BLACKBURN, G., Proc. Natl. Acad. Sci. USA 93 (1996), p. 799–803 POLLACK, S. J., JACOBS, J. W., SCHULTZ, P. G., Science 234 (1986), p. 1570–1573 DATTA, A., WENTWORTH Jr., P., SHAW, J. P., SIMEONOV, A., JANDA, K. D., J. Am. Chem. Soc 121 (1999), p. 10461–10467 KOHEN, F., KIM, J. B., LINDNER, H. R., ESHHAR, Z., GREEN, B., FEBS Lett 111 (1980), p. 427–431 SCHOWEN, R. L., J. Immunol. Methods 269 (2002), p. 59–65 WAGNER, J., LERNER, R. A., BARBAS III, C. F., Science 270 (1995), p. 1797–1800 ZHONG, G. F., LERNER, R. A., BARBAS, C. F., Angew. Chem. Int. Ed. 38 (1999), p. 3738–3741 TSUMURAYA, T., TAKAZAWA, N., TSUNAKAWA, A., FLECK, R., MASAMUNE, S., Chem. Eur. J. 7 (2001), p. 3748–3755

70 ANGELES, T. S., SMITH, R. G., DARSLEY, M. J., SUGASAWARA, R., SANCHEZ, R. I., KENTEN, J. SCHULTZ, P. G., MARTIN, M. T., Biochemistry 32 (1993), p. 12128–12135 71 ANGELES, T. S., MARTIN, M. T., BioChem. Biophys. Res. Commun. 197 (1993), p. 696–701 72 MARTIN, M. T., NAPPER, A. D., SCHULTZ, P. G., REES, A. R., Biochemistry, (1991), 30, p. 9757–9761 73 GIGANT, B., CHARBONNIER, J. B., GOLINELLI-PIMPANEAU, B., ZEMEL, R. R., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., Eur J., BioChem. 246 (1997), p. 471–476 74 TAWFIK, D. S., CHAP, R., ESHHAR, Z., GREEN, B. S., Protein Eng. 7 (1994), p. 431–434 75 STEWART, J. D., KREBS, J. F., SIUZDAK, G., BERDIS, A. J., SMITHRUD, D. B., BENKOVIC, S. J., Proc. Natl. Acad. Sci. USA 91 (1994), p. 7404–7409 76 JANDA, K. D., SCHLOEDER, D., BENKOVIC, S. J., LERNER, R. A., Science 241 (1988), p. 1188–1191 77 BENKOVIC, S. J., ADAMS, J. A., BORDERS, Jr., C. L., JANDA, K. D., LERNER, R. A., Science 250 (1990), p. 1135–1139 78 GIBBS, R. A., BENKOVIC, P. A., JANDA, K. D., LERNER, R. A., BENKOVIC, S. J., J. Am. Chem. Soc 114 (1992) 79 THAYER, M. M., OLENDER, E. H., ARVAI, A. S., KOIKE, C. K., CANESTRELLI, I. L., STEWART, J. D., BENKOVIC, S. J., GETZOFF, E. D., ROBERTS, V. A., J. Mol. Biol. 291 (1999), p. 329–345 80 JENCKS, W. P., Catalysis in chemistry and enzymology, McGraw-Hill Book Company, New-York 1969, p. 67–72 81 STEWART, J. D., ROBERTS, V. A., THOMAS, N. R., GETZOFF, E. D., BENKOVIC, S. J., Biochemistry 33 (1994), p. 1994–2003 82 KIMBALL, A. S., LEE, J., JAYARAM, M., TULLIUS, T. D., Biochemistry 32 (1993), p. 4698–4701 83 VILADOT, J. L., de RAMON, E., DURANY, O., PLANAS, A., Biochemistry 37 (1998), p. 11332–11342 84 ADMIRAAL, S. J., SCHNEIDER, B., MEYER, P., JANIN, J., VERON, M., DEVILLE-BONNE, D., HERSCHLAG, D., Biochemistry 38 (1999), p. 4701–4711

33

34 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

85 LEE, Y. L., CHEN, J. C., SHAW, J. F., BioChem. Biophys. Res. Commun. 231 (1997), p. 452–456 86 STOOPS, J. K., HORGAN, D. J., RUNNEGAR, M. T., De JERSEY, J., WEBB, E. C., ZERNER, B., Biochemistry 8 (1969), p. 2026–2033 87 FASTREZ, J., FERSHT, A. R., Biochemistry 12 (1973), p. 2025–2034 88 SELZER, T., ALBECK, S., SCHREIBER, G., Nat. Struct. Biol. 7 (2000), p. 537–541 89 ERSOY, O., FLECK, R., SINSKEY, A., MASAMUNE, S., J. Am. Chem. Soc. 118 (1996), p. 13077–13078 90 JANDA, K. D., WEINHOUSE, M. I., SCHLOEDER, D., LERNER, R. A., J. Am. Chem. Soc. 112 (1990), p. 1274–1275 91 SUGA, H., ERSOY, O., TSUMURAYA, T., LEE, J., SINSKEY, A. J., MASAMUNE, S., J. Am. Chem. Soc. 116 (1994), p. 487–494 92 GRYNSZPAN, F., KEINAN, E., P.-, A., Chem. Commun. 21 (1998) 93 IWABUCHI, Y., KURIHARA, S., ODA, M., FUJII, I., Tetrahedron Lett. 40 (1999), p. 5341–5344 94 GOLINELLI-PIMPANEAU, B., GONCALVES, O., DINTINGER, T., BLANCHARD, D., KNOSSOW, M., TELLIER, C., Proc. Natl. Acad. Sci. USA 97 (2000), p. 9892–9895 95 LARSEN, N. A., HEINE, A., CRANE, L., CRAVATT, B. F., LERNER, R. A., WILSON, I. A., J. Mol. Biol. 314 (2001), p. 93–102 96 CROSS, S. S., BRADY, K., STEVENSON, J. D., SACKIN, J. R., KENWARD, N., DIETEL, A., THOMAS, N. R., J. Immunol. Methods 269 (2002), p. 173–195 97 STEVENSON, J. D., DIETEL, A., THOMAS, N. R., Chem. Commun. 1999 (1999), p. 2105–2106 98 WILLIAMS, N. H., TAKASAKI, B., WALL, M., CHIN, J., Acc. Chem. Res. 32 (1999), p. 485–493 99 WADE, W. S., ASHLEY, J. A., JAHANGIRI, G. K., MCELHANY, G., JANDA, K. D., LERNER, R. A., J. Am Chem. Soc 115 (1993), p. 4906–4907 100 BRUMMER, O., HOFFMAN, T. Z., JANDA, K. D., Bioorg. Med. Chem. 9 (2001), p. 2253–2257 101 HEDSTROM, L., Chem. Rev. 102 (2002), p. 4501–4524 102 GIGANT, B., CHARBONNIER, J. B., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., J. Mol. Biol. 284 (1998), p. 741–750

103 D’SOUZA, L. J., GIGANT, B., KNOSSOW, M., GREEN, B. S., J. Am. Chem. Soc. 124 (2002), p. 2114–2115 104 WADE, H., SCANLAN, T. S., J. Am. Chem. Soc., (1999), 121, p. 11935–11941 105 FINER-MOORE, J. S., SANTI, D. V., STROUD, R. M., Biochemistry, (2003), 42, p. 248–256 106 STROUD, R. M., FINER-MOORE, J. S., Biochemistry, (2003), 42, p. 239–247 107 SHI, J., RADIC, Z., TAYLOR, P., J. Biol. Chem., (2002), 277, p. 43301–43308 108 SHI, J., BOYD, A. E., RADIC, Z., TAYLOR, P., J. Biol. Chem. 276 (2001), p. 42196–42204 109 PADLAN, E. A., Mol. Immunol. 31 (1994), p. 169–217 110 FOOTE, J., MILSTEIN, C., Proc. Natl. Acad. Sci. USA 91 (1994), p. 10370–10374 111 LANCET, D., PECHT, I., Proc. Natl. Acad. Sci. USA 73 (1976), p. 3549–3553 112 WILSON, I. A., STANFIELD, R. L., Curr. Opin. Struct. Biol. 4 (1994), p. 857–867 113 JAMES, L. C., ROVERSI, P., TAWFIK, D. S., Science 299 (2003), p. 1362–1367 114 PELLEQUER, J. L., CHEN, S., ROBERTS, V. A., TAINER, J. A., GETZOFF, E. D., J. Mol. Recognit. 12 (1999), p. 267–275 115 ULRICH, H. D., MUNDORFF, E., SANTARSIERO, B. D., DRIGGERS, E. M., STEVENS, R. C., SCHULTZ, P. G., Nature 389 (1997), p. 271–275 116 YIN, J., MUNDORFF, E. C., YANG, P. L., WENDT, K. U., HANWAY, D., STEVENS, R. C., SCHULTZ, P. G., Biochemistry 40 (2001), p. 10764–10773 117 YANG, P. L., SCHULTZ, P. G., J. Mol. Biol. 294 (1999), p. 1191–1201 118 SCHULTZ, P. G., LERNER, R. A., Nature 418 (2002), p. 485 119 ROMESBERG, F. E., SPILLER, B., SCHULTZ, P. G., STEVENS, R. C., Science 279 (1998), p. 1929–1933 120 ARKIN, M. R., WELLS, J. A., J. Mol. Biol. 284 (1998), p. 1083–1094 121 JANDA, K. D., BENKOVIC, S. J., LERNER, R. A., Science 244 (1989), p. 437–440 122 TANAKA, F., KINOSHITA, K., TANIMURA, R., FUJII, I., J. Am. Chem. Soc. 118 (1996), p. 2332–2339 123 KURIHARA, S., TSUMURAYA, T., SUZUKI, K., KURODA, M., LIU, L., TAKAOKA, Y., FUJII, I., Chem. Eur. J. 6 (2000), p. 1656–1662

References 124 SHI, Z. D., YANG, B. H., ZHAO, J. J., WU, Y. L., JI, Y. Y., YEH, M., Bioorg. Med. Chem. 10 (2002), p. 2171–2175 125 WADE, H., SCANLAN, T. S., J. Am. Chem. Soc. 118 (1996), p. 6510–6511 126 FINN, M. G., LERNER, R. A., BARBAS, C. F., J. Am. Chem. Soc. 120 (1998), p. 2963–2964 127 GRIFFITHS, A. D., TAWFIK, D. S., Curr. Opin. Biotechnol. 11 (2000), p. 338–353 128 GIGANT, B., CHARBONNIER, J. B., ESHHAR, Z., GREEN, B. S., KNOSSOW, M., Proc. Natl. Acad. Sci., USA 94 (1997), p. 7857–7861 129 CHARBONNIER, J. B., GOLINELLI-PIMPANEAU, B., GIGANT, B., GREEN, B. S., KNOSSOW, M., Isr. J., Chem. 36 (1996), p. 171–176 130 BRAXTON, S., WELLS, J. A., J. Biol. Chem. 266 (1991), p. 11797–11800 131 PHILLIPS, M. A., FLETTERICK, R., RUTTER, W. J., J. Biol. Chem. 265 (1990), p. 20692–20698 132 MENARD, R., STORER, A. C., Biol. Chem. Hoppe-Seyler, 373 (1992), p. 393–400

133 NICOLAS, A., EGMOND, M., VERRIPS, C. T., DEVLIEG, J., LONGHI, S., CAMBILLAU, C., MARTINEZ, C., Biochemistry 35 (1996), p. 398–410 134 JANDA, K. D., WEINHOUSE, M. I., DANON, T., PACELLI, K. A., SCHLOEDER, D., J. Am. Chem. Soc. 113 (1991), p. 5427–5434 135 ANDREEVA, N. S., RUMSH, L. D., Protein Sci. 10 (2001), p. 2439–2450 136 NORTHROP, D. B., Acc. Chem. Res. 34 (2001), p. 790–797 137 KLEIFELD, O., FRENKEL, A., MARTIN, J. M. L., SAGI, I., Nat. Struct. Biol. 10 (2003), p. 98–103 138 PENNER-HAHN, J. E., Nat. Struct. Biol. 10 (2003), p. 75–77 139 LINDNER, A. B., Ph.D. Thesis, Feinberg 2002, graduate school, Weizmann Institute of Science, Rehovot, Israel 140 HOFFMANN, T., ZHONG, G. F., LIST, B., SHABAT, D., ANDERSON, J., GRAMATIKOVA, S., LERNER, R. A., BARBAS, C. F., J. Am. Chem. Soc. 120 (1998), p. 2768–2779

35

1

Chemical Microarrays for Small Molecule Ligand Screening G. Metz, H. Ottleben, and D. Vetter

Grafﬁnity Pharmaceutical Design GmbH, Heidelberg, Germany

Originally published in: Protein-Ligand Interactions. Edited by Hans-Joachim B¨ohm, Gisbert Schneider. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30521-6

1 Introduction

Since their introduction about a decade ago, combinatorial chemistry and highthroughput screening (HTS) have become indispensable tools in the drug-discovery process. The possibility to synthesize ever-increasing numbers of molecules through novel chemistries and automation is stimulating the development of higher screening capabilities through miniaturization and robotics. Robust biochemical assay development is providing the basis for large-scale screening of biological targets. While in the early days much effort and hope were directed towards managing a numbers game, the focus is shifting from quantity towards quality. For instance, the screening of compound mixtures is being replaced by screening of individual substances in a one-well–one-compound fashion. The design of general-purpose screening libraries as well as corresponding follow-up strategies has become a key aspect in small molecule discovery and optimization. Hits from high-throughput screening enter a selection process to become the subject of medicinal chemistry approaches in lead optimization. Strategies are employed to improve potency, selectivity, and physicochemical proﬁle. If possible, several compound series are generated to allow for alternative routes in case of failure of one. Syntheses of analogues for further exploration are guided by a combination of medicinal chemistry knowledge and intuition, as well as quantitative structure-activity relationships, if available. Such studies help to deﬁne particular pharmacophoric features within the hit or lead molecule that constitute the underlying molecular recognition motifs between the ligand and its target and provide a hypothesis for its mode of action. The focus in screening for biological activity assays is on detecting hits with activities in the low micromolar range. Compounds exhibiting this level of activity tend to be of molecular weight in the range of 300–600 Da and of substantial functional Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Chemical Microarrays for Small Molecule Ligand Screening

complexity. A conceptually different strategy can be envisioned that – instead of trying to identify and keep relevant features in a rather complex hit compound – aims at a stepwise discovery starting from molecular fragments. Such fragments need to be screened with techniques suitable for the detection of presumably weak interactions. Guided by early stage structure-afﬁnity information, more potent compounds can then be assembled either from a combination of low afﬁnity binders or by chemical modiﬁcation of the initial fragments. This article describes the underlying philosophy of fragment-based discovery as well as the experimental approaches suitable for this promising discovery concept. While fragment-based ligand discovery was ﬁrst adapted in computational methods, several experimental techniques have been introduced recently. Biophysical methods such as nuclear magnetic resonance (NMR) and X-ray crystallography have been successfully applied to fragment discovery. A novel screening technique based on chemical microarrays in combination with label-free afﬁnity detection has emerged and will be discussed in detail.

2 Fragment Approaches 2.1 Conceptual Ideas

Primary screening efforts in drug discovery aim at the identiﬁcation of hit molecules with the necessary characteristics to be developed into a promising lead molecule. The deﬁnition of favorable properties of the starting screening compounds has gained much attention. The design of libraries with drug-like characteristics generally follows the so-called “rule of ﬁve” which has been established by retrospective analysis of known drugs and allows a quick assessment based on simple properties, namely, preferred ranges for molecular weight and clogP as well as the number of hydrogen bond donors and acceptors [1]. However, it has been proposed that the ideal proﬁle for hit or lead compounds is different from that of the ﬁnal drug molecule [2], in particular because hit or lead compounds must be amenable for further optimization. Three categories of lead compounds have been deﬁned based on their physicochemical properties and typical afﬁnities. First, it has been pointed out that hits from drug-like libraries rarely show high (<100 nM) afﬁnities. Typical optimization schemes tend to increase both molecular weight and lipophilicity. Therefore, if applied to already large compounds, they would fall out of the preferred drug-like ranges. Second, non-drug-like hits with very high afﬁnity at an early stage, e.g., natural compounds, usually are rather complex, and optimization towards drug-likeness is difﬁcult. The third category, namely, small (100–350 Da) molecules with low clogP (1–3) is being described as a favorable type of lead compound, as it still allows exploitation of additional (hydrophobic) interactions during combinatorial optimization schemes. Overall, the value of active screening compounds is judged considering afﬁnity relative to units of molecular weight and

2 Fragment Approaches

lipophilicity. Teague et al. suggest that the quality of hits emerging from screening could be improved by tailoring screening libraries towards such lead-like characteristics [2]. In the case of hit identiﬁcation, various techniques of library design and compound selection are applied to maximize the likelihood of discovery. The relationship between the probability of discovering molecules in a screening assay and their structural complexity was studied using a model of receptor-ligand interactions [3]. The ligand and the active site are represented by a linear string of binary (“+” and “−”) features, and a good ﬁt is indicated by complementarity of the aligned patterns. While molecular recognition is determined by a delicate interplay of physicochemical and steric complementarity, the apparent simpliﬁcation still allows one to address some key aspects. Varying degrees of relative complexity between the active site and the ligand are represented by different lengths of the binary patterns. By calculating matching probabilities for varying ligand complexities, it is observed that the likelihood of ﬁnding any ﬁt exponentially decreases with ligand complexity and that the curve describing the chance of ﬁnding just one unique match between ligand and active site features reaches a distinct maximum. The theoretical complexity of such single-mode binding was found to peak at a binary pattern length of three when compared to a site complexity of 12 features. For less complex ligands, multiple binding modes start to dominate the probability curve. While this ﬁnding cannot be directly translated into simple molecular features, it points towards a higher overall chance of hit discovery for relatively small molecules. However, smaller molecules will probably exhibit weaker binding, and therefore chances to experimentally detect binding will depend on the screening technology. Taking this into account, the peak of preferred ligand complexity would shift towards somewhat larger compounds. In summary, the existence of an optimal ligand complexity and its relation to the detection probability deﬁnes a “range of useful events” (Fig. 1) and leads to the postulation that screening simpler molecules is advantageous from a probabilistic point of view [3]. Molecular recognition addresses the aspect of not only afﬁnity but also speciﬁcity, and the question of unique binding modes becomes fundamental for smaller molecules. In general, the multitude of energetically similar but structurally different binding modes increases with fewer interaction features of the ligand. The corresponding free energy of binding landscape is termed “frustrated,” and both native and non-native binding modes of small molecules must be considered [4]. Ideally, the native binding mode is separated from alternate positions in the binding site by an energy “stability gap,” which ensures that a rather speciﬁc recognition motif is present (Fig. 2). Small fragment-like compounds exhibiting such a preferred binding mode, so-called molecular anchors, may be more suitable to combinatorial optimization than stronger binders with iso-energetic multiple binding modes [5]. With anchors serving as a receptor-speciﬁc recognition motif, a native binding mode would imply that addition of structural features to a binding fragment will not dramatically alter its orientation. Such modular approaches rely on the observation that additivity if not synergy is obtained when preselected fragments are combined. A point in case is a detailed structural comparison of enzyme-ligand binding which

3

4 Chemical Microarrays for Small Molecule Ligand Screening

Fig. 1 The theoretical probability of molecular recognition based on a simple interaction model indicates that the likelihood of a unique binding mode decreases with increasing ligand complexity (A). The probability to experimentally detect a binding event is estimated to increase with com-

plexity (B). The product probability for a so-called useful event, namely, the detection of a ligand with a unique binding mode, reaches a maximum at a medium ligand complexity (C). (See citation in text for details and discussion).

showed that binding modes for individual fragments of the thymidylate synthase substrate correspond well with the whole substrate in the active site [6]. The relationship between the potential afﬁnity of a given molecule and its actually measured binding strength to a given target has been addressed by analyzing individual functional group contributions to drug-receptor interactions [7]. Average binding energies of 10 common functionalities have been estimated based on a dataset of 200 drugs and enzyme inhibitors with known free energies of binding. The average binding energy for a given ligand, often also referred to as Andrews energy, can then be calculated by adding up the average contribution of the functional

Fig. 2. Concept of a stability gap between different binding modes of a fragment within an active site. The assumed binding energy for two different fragments is shown as a simple scheme. Fragment A qualiﬁes as a molecular anchor, as there is a difference in free energy of binding of the best placement to alternative placements. Such a stability gap is not observed for the second fragment B. Originally, these considerations were made based on docked placements of fragments and free energies as estimated from a scoring function (see citation in text).

2 Fragment Approaches

groups present in the molecule and taking into consideration an entropic correction term. The comparison of experimental and theoretical binding energy may then be used to classify the binding as better or worse than average, in other words as an indicator of a good or bad ligand-receptor ﬁt. Poor binders were found to be large, ﬂexible, and rich in polar groups. A similar ﬁnding was reported for 3000 screening compounds, where a large proportion exhibited lower observed afﬁnities when compared to the average Andrews energy based on the number of functionalities present. The general non-linearity of binding afﬁnity and ligand size has been pointed out in comparing binding afﬁnities relative to the number of non-hydrogen atoms in known protein-ligand complexes [8]. While a correlation of binding energy with size was found for smaller ligands, there also is evidence for an upper limit of observed afﬁnities for larger ligands that therefore under-perform according to their Andrews energy. The authors note that as few as 7–10 atoms allow for nanomolar binding constants and also suggest to assess relative afﬁnity during a drug-discovery process using expansion approaches. In a more pictorial phrase, it is the “bang per dalton” that has to be kept in mind when evaluating the outcome of hit and lead ﬁnding efforts. This ratio tends to be better for smaller-sized compounds. 2.2 Choice of Screening Fragments

The quality and quantity of fragment collections are critical issues for all fragmentbased discovery methods. Some considerations are related to deconvolution strategies based on unique NMR shifts or unique mass or shape, while others are sensitive to the detection limits and noise level of the respective experiment. Besides these technical aspects, the goal is to identify those fragments that best suit the concept of molecular anchors or recognition motifs. On the one hand, a certain size and complexity make a deﬁned binding mode more likely. On the other hand, in order to qualify as a fragment, screening compounds would be limited to a molecular weight range between 100 and 300 Da. The total number of fragments in a screening collection is related to the experimental throughput and the necessity to cover the “accessible fragment space” in order to increase the chances of ﬁnding a hit in the initial screen. The sufﬁcient presence of functional groups in the screening fragments is of importance for subsequent hit expansion strategies such as combining or decorating the initial hits. Systematic procedures for the identiﬁcation of suitable fragments or substructures have been implemented that aim to analyze active molecules in order to identify biologically relevant motifs. Because the ﬁeld is expanding, a diverse nomenclature is evolving; synonyms for chemical fragments are needles, molecular anchors, biophores, molecular frameworks, MULBITs, or base fragments. In a procedure called RECAP [9], fragmentation is limited to predeﬁned bond types leading to virtual fragments with chemical functionalities that allow them to be used as building blocks for combinatorial chemistry. The method was applied to a number of databases, including the World Drug Index (WDI). By examination of fragment distribution across different therapeutic areas, speciﬁc motifs were identiﬁed for use in target-biased screening libraries. Typically, the molecular weight of most

5

6 Chemical Microarrays for Small Molecule Ligand Screening

fragments was around 200 Da and the cleavage rules kept ring systems intact. While the approach mainly addressed the question of building block selection for combinatorial libraries, the authors also pointed out its usefulness for generating a fragment database for (computational) ligand buildup approaches. Different variations of the basic approach have been implemented [10–12]. In a related work, databases of known drugs were the basis for breaking down molecules into rings, linkers, and side chains [13, 14]. In this way both common molecular frameworks and side chains were identiﬁed. Interestingly, it was found that rather few frameworks and side chains represent the majority of compounds in drug-related compound collections such as the Comprehensive Medicinal Chemistry (CMC) database. Incorporating this ﬁnding into a general-purpose screening library resulted in the so-called SHAPES library for NMR-based screening [15]. Natural product databases were also used for the search of interesting molecular scaffolds [11]. It has long been pointed out [16] that certain substructural motifs, named privileged structures, are capable of serving as a starting point for ligands for more than one protein. Systematic modiﬁcations of such structures with substitution patterns have been successfully applied in medicinal chemistry [17]. Often these attractive scaffolds are rigid polycyclic heteroatomic systems that allow us to present binding elements in various directions within the binding site. The concept has been adopted as a design strategy for combinatorial libraries [18], and corresponding building blocks are being marketed under the name “optimers” [19]. Experimental screening of compounds enriched with motifs based on such privileged fragments is thought to increase the chances of identifying promising hits for medicinal chemistry programs. Even more, fragment-based screening would allow us to directly use known privileged fragments or close analogues thereof. On the other hand, screening fragments that are not necessarily related to known privileged structures might help us to discover novel chemical motifs displaying properties that qualify them to be called “privileged” [20]. In summary, fragment collections for the experimental screening techniques that will be outlined below either fall into the category of “diversity-oriented” collections or are selected with some bias. The focus is on “drug-relevant” substructures or structures that are targeted against a certain class of proteins where prior knowledge exists. In addition to ﬁnding novel leads based on fragment screening, the technique can also be applied in the optimization of a known binder where a particular substructure needs to be replaced by a bio-isoster. A prime example is the quest for alternatives to charged basic groups occupying the thrombin S1 binding pocket in order to enhance bioavailability [21]. 2.3 Experimental Approaches

Fragment-based discovery methods have long been implemented in computational de novo drug design [22, 23]. In the multiple fragment-positioning methods, various functional groups are ﬁrst placed within the active site, and, after preferred

2 Fragment Approaches

placements have been identiﬁed, linking of the fragments provides molecules that can be ranked according a chosen scoring function. Alternatively, sequential buildup strategies start from one placed fragment and successively add functional groups in order to “grow” the ligand, guided by the target structure and a suitable energetic scoring function. A similar strategy is also used in docking programs using an incremental construction algorithm that ﬁrst performs a fragmentation of the ligand and then, starting with a base fragment placement, continuously builds up the ligand in the binding site. Clearly, these techniques are very much related to the molecular anchor concept discussed above. Besides problems associated with de novo computational approaches such as the synthesizability of proposed virtual compounds, the most critical issue is the fragment ranking, which is calculated by either energy-based methods or rule-based scoring functions. These predictive limitations in computationally derived binding conformation make experimental methods that are guided by either observed activity or binding afﬁnity very attractive. In a hybrid approach, termed “biased needle screening,” in silico prescreening of molecular fragments is followed by a high-concentration bioassay, biophysical hit validation, and structure-based optimization [24]. Virtual pharmacophore screening of 350,000 compounds resulted in the identiﬁcation of a 3000-member subset with molecular weights below 300 Da. These needle compounds were tested in an activity assay customized to pick up even weak binders. Several structural series could be conﬁrmed, and a 3-D structure-guided optimization based on NMR and X-ray data gave novel, potent inhibitors of DNA gyrase. The authors state that initial HTS on this target did not deliver suitable lead structures. This ﬁnding supports the concepts outlined above and demonstrates the usefulness of needle screening as a new entry point to explore the chemical space. The so-called “target-guided ligand assembly strategy” starts from a library of possible binding fragments where each member possesses a common chemical linkage group [25]. Monomer screening against the target is then performed in a bioassay at high concentrations in order to detect even weak binders that then can be connected with a set of ﬂexible linkers for a second round of screening. The utility of the method was demonstrated by using the tyrosine kinase c-Src and a microtiter-based ELISA assay. After screening at two concentrations (1 mM and 500 µM) and inspection of the hits, 37 reagents were selected out of 305 O-methyl oximes in the primary screening library. Using ﬁve different linkers, homo- and heterodimers of these reagents were synthesized in single wells, resulting in mixtures of compounds with different linker length. After identiﬁcation of wells showing inhibition in a second screening assay at higher concentration, deconvolution of the mixtures by single-compound re-synthesis allowed to identify individual active substances and highlighted the importance of the linker length. The most potent compound identiﬁed in this manner exhibits an IC50 of 64 nM, a very large increase compared with IC50 values around 40 µM for the individual reagents alone. As with other methods described above, the synergy achieved and the lower number of compounds needed for screening demonstrate the advantage of modular approaches in general.

7

8 Chemical Microarrays for Small Molecule Ligand Screening

In order to discover weakly bound, low-molecular-weight (approx. 250 Da) ligands, a “tethering” strategy has been suggested that relies on the formation of a disulﬁde bond between the ligand and a cysteine residue on the protein [26]. This cysteine would either be present in the wild type or be genetically engineered in order to target a speciﬁc site in the protein. A library of disulﬁde-containing molecules was prepared and a mixture of a few substances (8–15) was incubated with the target protein under conditions that allow reversible reactions. It is expected that the formation of disulﬁde bonds to the protein is entropically stabilized for those compounds with an inherent afﬁnity to the protein in proximity to the cysteine. Tethered complexes could be identiﬁed using mass spectrometry, provided that the library had been designed to contain molecules with unique molecular mass. The method was applied to thymidylate synthase containing a cysteine in the active site. Several library members showed binding corresponding to millimolar inhibition constants, as determined in an enzymatic assay. From a series of related ligands displaying different binding behavior, qualitative early SAR information was obtained. The exact binding mode was determined with the same ligand attached at different nearby sites through cysteine mutations; interestingly, the location of the tethered molecule was conserved, indicating little inﬂuence of the tether on the binding mode. Structure-based modiﬁcations in analogy to known substrates improved the initial hit to an inhibitor in the sub-micromolar range. Techniques such as X-ray crystallography and NMR, which combine low afﬁnity screening capability with structural information, are very powerful tools. NMR spectroscopy plays an increasingly prominent role among the biophysical screening methods, and the experimental schemes are continuously being improved to widen the scope of applications [27–32]. Different detection strategies based either on ligand or target resonance signals have evolved. The approaches share the ability to detect weak yet speciﬁc binders. Design principles for NMR screening compounds have been reviewed [33]. An early and frequently cited fragment-based experimental screening approach was termed “SAR by NMR” [34, 35]. Here, mixtures of small organic molecules together with 15 N-labeled protein were subjected to 2-D 1 H- 15 N NMR measurements. Protein chemical shift variations relative to spectra of the protein alone indicated a binding event. Based on the predetermined assignment of chemical shifts, both the location of the binding site in the protein and the binding molecule were identiﬁed. This initial binder was then used to saturate the protein in order to ﬁnd a second small molecule binding to a proximal site in a new round of screening. After neighboring small molecules were identiﬁed and optimized through an analogue approach, combinations with various linkers were synthesized and assayed. The linker design was supported by the 3-D structure of the protein-ligand complex. As a result of a successful linkage, binders were obtained with high afﬁnities even exceeding the product of the binding constants of the individual fragments due to linker-mediated entropic enhancement. The method has been successfully applied in a number of studies [36–40] elegantly combining the fragment-based strategy with use of structural information. However, the original method can be applied only if sufﬁcient (∼200 mg) N-labeled, soluble (at 2 mM) protein of limited size

2 Fragment Approaches

(< 40 kDa) is available, and it requires chemical shift assignment before the actual screening. The necessity for labeled proteins and the size limitation can be overcome by techniques that monitor not protein NMR signals but ligand resonances, either through line-broadening experiments, transferred NOE measurements, or relaxation- and diffusion-edited methods using pulsed ﬁeld gradients. The NMR SHAPES strategy utilizes 1-D line broadening and 2-D transferred NOE measurements to identify binders in a mixture of compounds [15]. Potential weakly binding scaffolds (µM to mM) are selected based on an analysis of known drugs and represent molecular frameworks recurrently found in active therapeutic molecules. In contrast to the SAR by NMR method, the strategy does not aim to ﬁnd highly potent ligands by NMR screening but instead empirically provides a basis for compound selection. In this way, libraries aimed at HTS screening may be biased by ﬁltering against the target-speciﬁc chemical motifs identiﬁed by the SHAPES approach. The use of X-ray crystallography for primary screening is conceptually related to the SAR by NMR technique. Again, the sensitivity of the method makes it well positioned for fragment-based discovery. Because organic solvent molecules contain functional groups representative of those found in screening compounds, it has been suggested to co-crystallize proteins with different solvents and to experimentally determine preferred locations of small organic molecules [41]. The position of several organic molecules was thought to provide initial templates for selection of screening compounds or rational ligand design efforts. In yet another fragment strategy termed CrystaLead [42], the electron density map of the protein was determined to identify protein and solvent densities in the unbound state. Next, the crystal was exposed to a mixture of small organic molecules in a soaking experiment, and binders were identiﬁed by their appearing electron densities. It was crucial for the identiﬁcation of the binding compounds in the screening mixture that they were of diverse and unique molecular shapes. Weak binding compounds (up to high µM) were detected and then optimized in a structure-directed process. The CrystaLead technology was demonstrated in a urokinase screen of 6–8 compound mixtures against 9 crystals. In this way, 61 fragments were exposed to the crystal soaking experiment, and 5 binders with afﬁnities in the low and high µM range were discovered. One compound was further optimized using additional structural knowledge from a known inhibitor and resulted in an optimized lead of 370 nM afﬁnity to urokinase. Crystallographic screening has gained signiﬁcant interest in the commercial setting where high-throughput crystallography laboratories and technological advances in both hardware and software have increased the rate of protein-ligand crystal structure determination. The advent of structural genomics initiatives has spurred the development of robotics and automated data interpretation [43, 44]. The latest developments have recently been reviewed [45–47]. These technological advances are now being paired with fragment-based approaches. Results have been reported [46] where deﬁned binding could be elucidated for very small fragments (< 200 Da), supporting the fact that despite small size and presumably weak interactions, fragments are able to make key interactions in order to act as molecular

9

10 Chemical Microarrays for Small Molecule Ligand Screening

anchor [4]. In a study with a protein kinase, such a weak initial fragment was optimized into a nanomolar inhibitor by the aid of structure-based design [45].

3 Chemical Microarrays 3.1 Background

Chemical microarrays can be deﬁned as collections of chemical compounds covalently immobilized on a carrier in a 2-D pattern. After the widespread adoption of DNA-based microarrays in basic and commercial research, there is a growing interest to extend the array concept beyond genetics applications. Exploratory work is underway to lay out proteins and cells in array formats [48–51]. In parallel, various routes are being taken to realize arrays of small molecules such as synthetic chemicals, peptides, or natural products. The principle of a regular 2-D arrangement of chemical diversity is well known from high-throughput screening, which currently serves as the paradigm in de novo small molecule discovery. In this highly industrialized undertaking, compounds are solubilized and deposited in wells of microtiter plates as either single compounds or mixtures. A recent modiﬁcation of HTS is the deposition of chemical-containing droplets on ﬂat surfaces [52] to reduce sample consumption. In the latter, the target protein is captured in a hydrogel that is subsequently added to cover the surface and resolubilize the screening compounds. It is an inherent problem of HTS that small molecule solubility can vary greatly and can hardly be accounted for. Therefore, the concentration of any given compound in the screening mixture is difﬁcult to predict and essentially unknown at the primary screening stage. This limitation is especially apparent in a fragment-based screening regime. Because higher compound concentrations are employed to facilitate the detection of binders below the usual micromolar cutoff, DMSO tolerance of the target protein or insolubility of the small molecules restricts this approach to robust assay systems and more hydrophilic fragment diversity. When we set out to develop a platform for chemical microarrays, compatibility with fragment-oriented diversity was a major design principle. We saw great promise in also solubilizing hydrophobic compounds by hydrophilic or amphiphilic spacer moieties. Such spacer groups are required for covalent tethering of the ligands to the array surface and for access to protein-binding pockets. The amphiphilic nature of the spacer aids hydration of small molecules associated with high clogP values. The ﬂexibility and the length of the spacer chain allow for the ligands to reach even so-called deep protein-binding sites. Covalent tethering of small molecules is generally seen as an obstacle for deriving structure-activity relationships. The compounds are not free to orient themselves for optimal ﬁt with the protein receptor, the spacer might sterically clash with the protein surface, and the spacer attachment itself might affect the electronic properties of the ligands. Consequently, in order to generate covalent modiﬁcations

3 Chemical Microarrays

of known drugs or other compounds known to be bioactive, the decision for regioselective spacer attachment should be based on a receptor-ligand complex structure if possible. Nevertheless, for de novo discovery of small molecule fragments, the spacer-mediated, structure-independent compound solubilization outweighs the above-mentioned potential disadvantage of restricted orientation. Furthermore, fragments can be conjugated to the spacer moiety by more than one type of coupling chemistry and through various functional groups present on the fragment itself. A third aspect of the small molecule “display” format is the additional information generated through knowledge of the tether site. The fact that possible orientations of a tethered compound within a binding site are more limited than in a homogeneous assay can facilitate building a hypothesis for a mode of action. The directionality can be used either in ligand-alignment procedures for SAR-type studies or as a bias in docking algorithms in order to evaluate possible binding modes. Other advantages of using chemical microarrays of peptides and organic compounds are in the minimization of biological sample consumption and the resulting increase in screening throughput. Array approaches hold the promise of mass production and industrialization. A simple format for reliably storing and reading chemical diversity is attractive compared to the current cumbersome fashion of operating refrigerated warehouse-type storage and dispensing systems. Chemical microarrays promise to enable function-blind screening of large numbers of novel targets. In general, chemical microarray approaches open up the opportunity to map interactions and discover small molecule binders for a given protein of interest even before understanding the protein’s function. Once the arrays are produced, the only prerequisite for array-based screening is the preparation of puriﬁed, homogeneous, and soluble protein. Advances in protein-production and -puriﬁcation methods, such as expression systems optimization, protein folding, and afﬁnity-tag development, complement array-based screening. It can be safely estimated that several thousands of novel, non-membrane-bound, putative target proteins will be derived from the knowledge of the human genome [53]. Chemical microarrays provide a potentially powerful alternative to high-throughput screening for small molecule de novo discovery on this wealth of novel targets. 3.2 On-array Synthesis

Chemical diversity for chemical microarrays can be accessed in two ways: importing compounds onto the chip surface or synthetically creating molecules directly on the support. In the pioneering work by Fodor and coworkers, a combination of solid-phase organic synthesis and photolithographic techniques was applied to the in situ creation of oligonucleotide and peptide microarrays [54, 55]. Technological challenges associated with this approach have limited it to the generation of oligonucleotide arrays. DNA consists of only four different nucleotides, and differences in reactivities among the four nucleotides on glass surfaces are well understood, and the solid-phase synthesis is highly optimized. Hence, highly

11

12 Chemical Microarrays for Small Molecule Ligand Screening

miniaturized synthesis of a vast diversity of different single-strand probes became feasible, and corresponding commercial products derived from photolithography are in widespread use. In contrast, the application of on-array synthesis techniques is more demanding for the generation of peptidic and even more challenging for combinatorial chemical libraries. In these cases, a much larger number of building blocks with greater ranges of reactivities is required. The greatest hurdle to highdensity, in situ array synthesis on glass is quality control of the numerous products generated. Techniques to infer physicochemical characteristics of monomolecular ﬁlms in micrometer-sized areas are just beginning to evolve and are still far away from becoming a routine application such as high-pressure liquid chromatographycoupled mass spectrometry (LC-MS), which is used for conventional combinatorial compound library analysis. A different way to produce chemical microarrays in situ is spot synthesis of combinatorial libraries on cellulose sheets [56]. Spot synthesis is conﬁgured as an open system to be operated at room temperature. Despite attempts to replace cellulose with polypropylene as a synthesis support [57], cellulose is still the support of choice for spot synthesis, and reaction conditions have to be compatible with the restricted chemical stability of cellulose. Due to the acid lability of such membranes, the diversity content of these arrays was initially restricted to the synthesis of peptides. Recently, a method was described that could widen the scope of spot synthesis arrays. Germeroth and coworkers [57] succeeded in the assembly of a library of 8000 cellulose-bound 1,3,5-triazines under mild reaction conditions. They employed a strategy that took advantage of a temperature-dependent, successive displacement of cyanuric chlorides by different nucleophiles in a ﬁrst report of the synthesis of small organic compounds on cellulose sheets. In addition to the limited range of cellulose-compatible synthesis protocols, two further drawbacks remain that are inherent to on-array approaches. Because synthesis takes place directly on the surface that is subsequently used for screening, quality control of the synthesis products is restricted to the surface-bound molecules. Standard cleave-and-characterize procedures involving LC-MS analytical techniques are not practical. This is a severe problem even for the synthesis of peptides where well-established protocols are available and becomes more pronounced when novel chemistries have to be employed. Secondly, each array produced is unique, which renders the production rather costly and prevents the generation of numerous copies of the same array for high-throughput applications. 3.3 Off-array Synthesis and Spotting

Some of the problems associated with on-array in situ synthesis can be overcome by a technology recently published by Stuart Schreiber’s group. In this work, the compound diversity was generated in solution or solid-phase formats different from the array layout, and chemical microarrays were subsequently produced by spotting pre-synthesized molecules [58–60]. In a proof-of-principle experiment, three

3 Chemical Microarrays

different organic compounds were immobilized on a chip surface [58]. For this purpose a silanated glass slide was derivatized to give a surface densely functionalized with thiol-reactive maleimide groups. Onto these surfaces, a high-precision robot delivered approximately 1 nL of a solution of the three different organic compounds, which were pre-synthesized to contain a spacer with a thiol group. In this way, an array of 10,800 spots each 200–250 µm in diameter was produced on a 2.5 × 7.5 cm glass slide. The chips were afterwards probed with respective ﬂuorescence-labeled proteins for selectively binding their ligands. The ﬂuorescence intensity recorded on the spot reﬂected the binding afﬁnity of the respective ligand. This ﬁnding demonstrates the feasibility of semi-quantitative measurements of ligands with binding afﬁnities in the low micro- to nanomolar range. In a second communication a similar technical setup was combined with a different chemical immobilization strategy [59]. The same set of organic probe molecules was presynthesized, this time containing primary alcohol groups, and spotted directly onto thionylchloride-activated glass slides. A small combinatorial library comprising 78 compounds was prepared in analogous fashion and probed with a target protein, and a new “hit” was identiﬁed in addition to the positive controls. Although the initial results of this approach were very encouraging, discovering new ligands for broader protein diversity would require a larger number of organic compounds. Moreover, the surface chemistries employed in both papers did not allow a tight control of the number and density of the compounds on each spot of every chip. This might become limiting when a more quantitative analysis of the results is required or when lower afﬁnity interactions are to be analyzed. In the approach cited above, both the nanoliter droplet deposition technique and the surface design of the array support were adopted from DNA microarray fabrication [61]. The presentation of oligonucleotides or DNA on surfaces as well as the readout of such chips in hybridization experiments are facilitated by the fact that both ligand and receptor molecules are relatively similar with respect to solubility, charge density, and pI. The physicochemical properties of nucleotides make it possible to design surfaces that show relatively little background binding under various conditions. However, the situation is quite different if proteins are the target receptors and protein binding rather than hybridization is of interest. Proteins differ dramatically in stability, solubility, and hydrophobicity and can be basic or acidic. The challenge here is to design a surface that resists the unspeciﬁc binding of a wide range of different proteins. In addition, probing a DNA microarray by hybridization is relatively simple, as DNA is rather stable and the binding constant for a double-strand formation is relatively high, allowing one to subject the molecules to stringent hybridization conditions. The interactions of small organic molecules or peptides with proteins are often much weaker and range mainly from milli- to micromolar binding constants. This requires high-performance surfaces to minimize unspeciﬁc binding. A common drawback of glass slides as supports for chemical microarrays, as well as of cellulose or polypropylene sheets, is the protein compatibility of their respective surface chemistries. It is inherently difﬁcult to render polypropylene or

13

14 Chemical Microarrays for Small Molecule Ligand Screening

silanated glass resistant to unspeciﬁc protein adsorption and is even more challenging to control a critical parameter such as ligand density on these polymers. For instance, engineering for biocompatibility of resins used in organic synthesis has become a major challenge for “bead-binding assay” screens [62]. In contrast, self-assembling monolayers (SAMs) of thiols on gold are not only among the best-characterized and best-deﬁned synthetic surfaces but also can be designed to exhibit very low background protein binding. SAMs give excellent control over surface properties at the molecular level. The basic principles of SAM formation and applications in protein binding to immobilized ligands have been reviewed [63, 64]. In general, an alkane thiol is chemisorbed onto a gold surface, and the packing of the hydrocarbon chains creates a dense monolayer on the gold. Attaching an oligo(ethylene oxide) to the hydrocarbon chain confers excellent resistance to nonspeciﬁc protein and DNA binding, biocompatibility, and nonfouling properties to the surface [65–67]. Reactive SAMs form uniform layers that contain reactive groups such as amines or carboxylic acids at their surface [68]. The ratio of alkane thiols in a mixture of molecules with and without a reactive group allows one to control the presentation of ligands in a deﬁned surface density (Fig. 3). Carboxylic acid-containing surfaces can be activated by pentaﬂuorophenyl or N-hydroxy succinimide esters. Reactive SAM technology has recently been used for peptide [69] and protein arrays [70] as well. Using the immobilized ligand benzene sulfonamide together with carboanhydrase, it was demonstrated that the unspeciﬁc binding can be minimized and

Fig. 3 Creating of a chemical microarray occurs in two steps. A self-assembled monolayer is formed on gold-coated glass from a diluent and an anchor molecule carrying a reactive group. Spotting of ligands attached

to a ChemTag linker results in covalent immobilization of organic molecules. The uniform density of the ligands at each spot is deﬁned by the ratio of anchor and diluent molecules.

3 Chemical Microarrays

that there was no irreversible adsorption. The measurements also were not complicated by mass transport [71, 72]. However, when rate constants for association and dissociation were analyzed, some lateral steric effects affected the binding kinetics. Ligands displayed on mixed self-assembled monolayers ligands in well-deﬁned surface densities offer control over critical parameters to minimize unspeciﬁc or irreversible binding or deviation of the binding kinetics from those observed in solution. In our laboratories, we developed a proprietary SAM technology comprising a functionalized anchor molecule diluted with unfunctionalized spacer molecules. The chemical functionality is used to capture spotted ligand molecules, which themselves are carrying a linear spacer and functionalization molecule, named ChemTag. The system was tested against numerous well-known pairs of interacting molecules such as receptor:ligand, enzyme:co-factor/inhibitor, and antigen:antibody for detection of speciﬁc interactions. Using a variety of proteins differing in hydrophobicity and molecular weight, the results showed that — irrespective of the protein chosen and even at high concentration — unspeciﬁc binding was minimized. These experiments proved that interactions between proteins and ligands of low molecular weight (<200 Da) as well as weak afﬁnities up to the millimolar range can be speciﬁcally detected, thus fulﬁlling a major requirement for the use of chemical microarrays for fragment-oriented screening applications. Building on the advantages provided by reactive mixed self-assembled monolayers, we designed both a compatible spacer chemistry and a miniaturized format for highly parallel solid-phase synthesis. Compounds are synthesized through combinatorial chemistry on a solid phase that is preloaded with the ChemTag spacer molecule. This linear spacer carries two functional groups, located at alpha and omega sites. The alpha group is used for transient conjugation to the synthesis support through a standard linker and remains protected in this fashion during the synthesis cycles. The omega group is available for ligand synthesis and can be functionalized in various ways to allow for different coupling chemistries. Cleavage from the solid phase by splitting the linker-alpha group bond yields free, spacermodiﬁed compounds in solution, with the alpha group free to be conjugated to the reactive sites on the microarray SAM surface. The common spacer group acts as a chemical shuttle to transport substances from one solid phase to another (Fig. 4). After solid-phase assembly, products are separated from the synthesis resin and stored in mother microtiter plates. At this stage, aliquots are subjected to quality control using LC-MS. After quality assessment, nanoliter amounts are transferred from the mother plates onto the surfaces of microarrays coated with the reactive mixed SAM. High-precision spotting of the substances in a custom-built automated environment ensures reproducible immobilization of the compounds onto the microarrays. A great variety of organic molecules can be attached through one of the different omega-site chemistries of the ChemTag spacer. The array content of our chemical microarrays ranges from the immobilization of single fragments to combinatorial libraries. As an example for a fragment array, up to 1536 individual monomers have been attached to the ChemTag and have been immobilized on a single array. Using

15

16 Chemical Microarrays for Small Molecule Ligand Screening

Fig. 4 The chemical microarray production starts by assembling combinatorial libraries or single fragments (A) to the ChemTag, which itself is attached to a solid-phase resin (B) resulting in compounds attached to the resin (C). After cleavage from the

resin, the tagged compounds are stored in mother plates (D) and LC/MS quality control can be performed (E). The ﬁnal step is spotting of the compounds onto a prefabricated microarray carrying a reactive selfassembled monolayer (F).

different chemistries, multifunctional fragments can be immobilized in more than one way. In addition, binary libraries with a batch size of 10,000 compounds have been synthesized on a nanoscale level (40 nmoles/compound) and then spotted on a single array. This rather low synthesis scale of chemical products is sufﬁcient for the production of hundreds of ready-to-screen arrays (Fig. 5).

Fig. 5 Chemical microarrays as they are used at Grafﬁnity. The array is based on a microtiter plate footprint and carries up to 10,000 individual compounds spotted in rows and columns onto an optical microstructure.

4 Screening on Microarrays

4 Screening on Microarrays 4.1 Detection Technology

Chemical microarrays that are based on glass or cellulose or other polymer surfaces were typically interrogated for protein binding by ﬂuorescence or chemiluminescence, respectively. Fluorophore-labeled protein samples were incubated with chemical microarrays, washed, and subsequently scanned by commercial microarray readers. The use of orthogonally labeled proteins was also described [58, 59]. Compound libraries on cellulose sheets are not readily subjected to ﬂuorescence imaging, but a multi-step process of target protein1 and target-directed antibody together with enzyme-conjugated secondary antibody binding serves to obtain high-sensitivity images of protein-binding patterns. In our approach towards chemical microarray screening, we wanted to exploit the fact that self-assembled monolayers readily lend themselves to label-free detection based on surface plasmon resonance (SPR). The key to this approach is a thin gold metal ﬁlm on a glass support that allows for both the physical effect of plasmon formation and the formation of high-quality self-assembled monolayers. Label-free imaging for chemical microarray readout is attractive because it reduces unspeciﬁc binding compared to chemiluminescence, as antibody and enzyme labels are not brought into contact with the array. The approach also allows us to observe the direct binding pattern of the unmodiﬁed target protein, as covalent modiﬁcations such as ﬂuorophore conjugations are obviated. Molecular recognition of an immobilized ligand and its solubilized binding partner can be detected by a physical phenomenon called surface plasmon resonance [73]. In general, the sensor chip comprises a glass prism covered with gold and coated with a SAM presenting the potential ligands (Fig. 6). Protein solution is added to a reservoir on top. Upon binding, an afﬁnity-dependent mass change occurs at the interface of the detector surface and the liquid above. This inﬂuences the dielectric properties of the gold layer. The readout is achieved by measuring the exact resonance condition for an energy transfer from the photons of a light beam to the electrons in the gold layer placed on top of a glass prism. An incident beam from the bottom is reﬂected at the gold surface and three factors, namely, the angle of incidence, the wavelength, and the refractive index at the interface, determine the resonance condition. For a ﬁxed angle, the wavelength-dependent minimum of the reﬂected light is quantitatively related to the mass change due to binding. Likewise, an angular dependence at a deﬁned wavelength can be recorded. Optical biosensors and in particular those based on SPR have gained importance in many areas. The technique itself, biological applications, and the availability of commercial instruments have been reviewed [74]. The most common application in the area of protein-ligand interactions is the detailed study of binding kinetics

17

18 Chemical Microarrays for Small Molecule Ligand Screening

Fig. 6 Basic setup (A) for surface plasmon resonance-based detection of molecular recognition comprises a gold-covered glass prism and the observation of the reﬂected light intensity in dependence of the angle and wavelength. Binding of a soluble protein to an immobilized ligand inﬂuences

the resonance condition at the gold interface. In the case of high-density chemical microarrays (B), the actual sensor ﬁelds are arranged in rows and columns to allow the parallel detection of up to 10,000 individual binding events using imaging technologies.

[75–79]. It relies on immobilization of one of the binding partners on a surface, and complex formation is optically detected in real time by adding the solubilized partner, e.g., by means of a microﬂuidic ﬂow chamber. The sensitivity of the technique allowed the detection of small molecules binding to immobilized protein receptors in a number of cases. However, sensitivity is at the limits and throughput is low if the protein is immobilized and the surface is exposed to individual ligands in a sequential fashion. These restrictions do not apply for the chemical microarray format, where a high number of small molecules are immobilized and binding to the target protein is detected in parallel (Fig. 6). Because SPR is sensitive to mass change, the binding of large proteins increases sensitivity as compared to the setup where the protein is immobilized. The need for higher throughput has led to the development of instruments that are capable of working either sequentially or parallel on several sensor ﬁelds [80]. However, parallel detection comes into play only when the sensor technology is combined with 2-D arrays opening the technique for screening applications. We designed and built a number of Plasmon Imager devices to fulﬁll this need and to enable us to perform label-free, simultaneous binding detection for up to 10,000 immobilized small organic molecules against a macromolecular receptor in solution. The instruments are based on imaging technology where the reﬂected light from the entire array is captured by a CCD camera. The resonance condition for surface plasmons is determined by stepwise variation of the wavelength of the incident light. For each sensor ﬁeld on the array, the light intensity (in percent of 100) of the reﬂected light is recorded against the wavelength (in nm), and for each spot a nm value for minimal light intensity is obtained. In order to generate an output that corresponds to protein-binding pattern recognition, two measurements are necessary. The difference between a measurement with buffer alone and after adding the target protein deﬁnes the SPR signal, i.e., the nanometer shift of the resonance condition.

4 Screening on Microarrays

4.2 Protein Afﬁnity Fingerprints

Chemical microarray screening data are conveniently displayed in a 2-D format of squares arranged in rows and columns. This representation reﬂects the actual spatially encoded ligand positions on the grid layout of the microarray. Therefore, each square represents a single spot on the physical array and thus also a single array compound. The SPR shift that is obtained after exposing the chemical microarray to a protein can be shown as a grayscale or in color codes. We developed a proprietary visualization tool for point-and-click interrogation of each data point for both the chemical structure and the measured afﬁnity associated with it. The visualized dataset is called an afﬁnity ﬁngerprint for the protein of interest (Fig. 7). In the case

Fig. 7 Afﬁnity ﬁngerprint of a target protein probed against a chemical microarray presenting 9216 immobilized binary compounds. The color range goes from red to orange to yellow to green to blue to code for decreasing SPR signal. Rows and columns each represent one of two monomers of the binary library and clearly

a pattern of prominent rows and columns points to the presence of certain monomer fragments being highly populated among the hits. The pop-up structure displays an array compound example, with the ChemTag attachment site indicated by a grayﬁlled circle.

19

20 Chemical Microarrays for Small Molecule Ligand Screening

Fig. 8 SPR signal of a chemical microarray presenting 1200 immobilized fragments shown as a function of molecular weight (A). Even small structures around 150 Da show detectable signal. The correspond-

ing afﬁnity ﬁngerprint of the target protein probed against the chemical fragment microarray is shown as a grayscale, with stronger binders being darker (B).

of binary libraries, constructed from two sets of monomers, visualization of the afﬁnity ﬁngerprints easily deconvolutes the presence of building blocks in hits. By arranging the compounds in rows and columns corresponding to the presence of a single monomer, the occurrence of many strong signals in one column indicates the signiﬁcant contribution or even dominance of a monomer in the combinatorial hit compound. Besides such prominent fragments, indicated by “strong” rows or columns, hits also can be found for certain binary combinations where each monomer has little or even no “tolerance” for the second building block in order to show afﬁnity. Typically, both types of hits are found. Overall, afﬁnity ﬁngerprints are the large-scale documentation of patterns of molecular recognition between small organic molecules and proteins. Immobilization of a collection of commercially available monomers on chemical microarrays represents the equivalent of the single fragment screening methods described above. Fig. 8 shows the afﬁnity ﬁngerprint of a chemical microarray containing 1200 individual fragments. The immobilized compounds all qualify as needle compounds, with molecular weights from 80 to 350 Da. Fragments can be selected that display diversity in pharmacophoric motifs and/or are enriched in drug-related chemical motifs as outlined above. The relationship between molecular weight and obtained SPR signal shows that the smallest fragments with detectable signals have a molecular weight of around 150 Da. Binding constants of chemical microarray hit fragments are typically observed to be in the micromolar range. For example, a well-known needle-type molecule, namely, amino-benzamidine with a K i of 80 µM against thrombin, is detectable within a fragment afﬁnity ﬁngerprint. After identiﬁcation of binders from the array screening, the ChemTag can be replaced by a series of small substructures, leading to soluble compounds based

5 Conclusion 21

around the original array hit structure. This so-called tag-replacement strategy makes use of the tether site to expand the array hits. The functional group originally connected to the ChemTag provides both a readily available linking chemistry and a bias towards an attachment point, with a higher chance of pointing towards additional interaction sites or subpockets within the protein. With tag replacement, areas are explored that are not yet occupied by the remainder of the ligand, and attachment sites are likely avoided that might result in clashes with the protein. Case studies showed that the afﬁnity of micromolar array hit compounds could be increased by one or two orders of magnitude. Such tag-replacement series link chemical microarray screening with follow-up medicinal chemistry efforts.

5 Conclusion

A number of technological advances in the area of surface chemistry and biophysical detection technologies have made fragment-based screening an attractive and promising tool for the early stages in drug discovery. The approach provides a new entry point for lead discovery, as it aims at ﬁnding drug fragments in a ﬁrst round of screening and using this information for an iterative buildup of chemical complexity. Such a procedure promises to address a number of issues encountered in the discovery and development of larger, more-complex screening compounds. Fragment-based screening allows us to investigate the relative contribution of the monomers in a reagent-based SAR evaluation, helping to avoid combinatorial explosion. Despite combinatorial chemistry, the number of accessible products by far exceeds the screening capabilities, and library diversity analysis tries to help reduce redundancy in compound collections to achieve a better coverage of “chemicals space.” Prescreening of monomers helps to better cover diversity by focusing combinatorial design around fragments that fared well in an early stage. Then, either smaller screening libraries for optimization can be synthesized based on these preferred monomers, or next rounds of testing can be based on fragment analogues. The experimental techniques that allow the detection of small and weak binders either are based on high-concentration bioassays or fall into the category of afﬁnity screening. Biological assays require careful setup and might not be available in general. NMR and crystallography have been used for some time now for fragmentscreening purposes, and elegant techniques have been developed to discover small fragments binding to target proteins. These techniques open the possibility of detailed insight into the binding mode of ligands, and their attraction lies in the seamless integration of structure-based optimization. Nevertheless, the experimental procedures are technically demanding and are limited either by crystallization conditions or NMR-related protein requirements. A novel emerging technology, label-free screening on chemical microarrays, brings together SPR, a well-known technique for studying molecular interactions, with self-assembled monolayer surface chemistry to display small organic

22 Chemical Microarrays for Small Molecule Ligand Screening

molecules in a miniaturized format. Progress in the area of label-free imaging now enables the parallel label-free afﬁnity ﬁngerprinting of 10,000 immobilized organic molecules against a target protein. The sensitivity of the method allows for the detection of weak interactions and makes routine, empirical fragment-based discovery a reality. The high-throughput, standardized screening format opens the possibility for chemical genomics applications aiming to move chemistry upstream in the discovery process. The goal is the early integration of chemical information either for target validation or for rapid identiﬁcation of novel leads on a genomic scale. Acknowledgments

The authors are grateful for discussions with the colleagues of Grafﬁnity Pharmaceuticals and want to emphasize the team effort that made SPR based screening on chemical microarrays possible. References 1 LIPINSKI, C. A., LOMBARDO, F., DOMINY, B. W., and FEENEY, P. J., Adv. Drug Deliv. Rev. 2001, 46, 3–26. 2 TEAGUE, S. J., DAVIS, A. M., LEESON, P. D., and OPREA, T., Angew. Chem. Int. Ed. 2001, 38, 3743–3748. 3 HANN, M. M., LEACH, A. R., and HARPER, G. J. Chem. Inf. Comput. Sci. 2001, 41, 856–864. 4 REJTO, P. A. and VERKHIVKER, G. M., Proc. Natl. Acad. Sci.U. S. A 1996, 93, 8945–8950. 5 REJTO, P. A. and VERKHIVKER, G. M., Pac. Symp. Biocomput. 1998, 362–373. 6 STOUT, T. J., SAGE, C. R., and STROUD, R. M., Structure. 1998, 6, 839–848. 7 ANDREWS, P. R., CRAIK, D. J., and MARTIN, J. L., J. Med. Chem. 1984, 27, 1648–1657. 8 KUNTZ, I. D., CHEN, K., SHARP, K. A., and KOLLMAN, P. A., Proc. Natl. Acad. Sci. U.S.A 1999, 96, 9997–10002. 9 LEWELL, X. Q., JUDD, D. B., WATSON, S. P., and HANN, M. M., J. Chem. Inf. Comput. Sci. 1998, 38, 511–522. 10 MAKINO, S., KAYAHARA, T., TASHIRO, K., TAKAHASHI, M., TSUJI, T., and SHOJI, M. J. Comput. Aided Mol. Des 2001, 15, 553–559. 11 SCHNEIDER, G., LEE, M. L., STAHL, M., and SCHNEIDER, P. J. Comput. Aided Mol. Des 2000, 14, 487–494. 12 LEWELL, X. Q. and SMITH, R. J. Mol. Graph. Model. 1997, 15, 43–46.

13 BEMIS, G. W. and MURCKO, M. A., J. Med. Chem. 1996, 39, 2887–2893. 14 BEMIS, G. W. and MURCKO, M. A., J. Med. Chem. 1999, 42, 5095–5099. 15 FEJZO, J., LEPRE, C. A., PENG, J. W., BEMIS, G. W., AJAY MURCKO, M. A., and MOORE, J. M., Chem. Biol. 1999, 6, 755–769. 16 EVANS, B. E., RITTLE, K. E., BOCK, M. G., DIPARDO, R. M., FREIDINGER, R. M., WHITTER, W. L., LUNDELL, G. F., VEBER, D. F., ANDERSON, P. S., CHANG, R. S., J. Med. Chem. 1988, 31, 2235–2246. 17 PATCHETT, A. and NARGUND, R., Annual Reports in Medicinal Chemistry 2000, 35, 289–297. 18 NICOLAOU, K., PFEFFERKORN, J., ROECKER, A., CAO, G., BARLUENGA, S., and MITCHELL, H. J. Am. Chem. Soc. 2000, 122, 9939–9953. 19 www.arraybiopharma.com. 2002. 20 HAJDUK, P. J., BURES, M., PRAESTGAARD, J., and FESIK, S. W., J. Med. Chem. 2000, 43, 3443–3447. 21 LUMMA, W. C., WITHERUP, K. M., TUCKER, T. J., BRADY, S. F., SISKO, J. T., NAYLER-OLSEN, A. M., LEWIS, S. D., LUCAS, B. J., and VACCA, J. P., J. Med. Chem. 1998, 41, 1011–1013. 22 APOSTOLAKIS, J. and CAFLISCH, A., Comb. Chem. High Throughput. Screen. 1999, 2, 91–104.

References 23 MURCKO, M. A. In: CHARIFSON, P. S., Editor, Practical Application of Computer-Aided Drug Design, Dekker, 1997. 24 BOEHM, H. J., BOEHRINGER, M., BUR, D., GMUENDER, H., HUBER, W., KLAUS, W., KOSTREWA, D., KUEHNE, H., LUEBBERS, T., MEUNIER-KELLER, N., and MUELLER, F., J. Med. Chem. 2000, 43, 2664–2674. 25 MALY, D. J., CHOONG, I. C., and ELLMAN, J. A. Proc. Natl. Acad. Sci.U. S. A 2000, 97, 2419–2424. 26 ERLANSON, D. A., BRAISTED, A. C., RAPHAEL, D. R., RANDAL, M., STROUD, R. M., GORDON, E. M., and WELLS, J. A., Proc. Natl. Acad. Sci.U. S. A 2000, 97, 9367–9372. 27 PELLECCHIA, M., SEM, D., and WUETHRICH, K., Nat. Rev. Drug Discov. 2002, 1, 211–218. 28 van DONGEN, M., WEIGELT, J., UPPENBERG, J., SCHULTZ, J., and WIKSTROM, M., Drug Discov. Today 2002, 7, 471–478. 29 DIERCKS, T., COLES, M., KESSLER, H., Curr. Opin. Chem. Biol. 2001, 5, 285–291. 30 BRADLEY, D., Modern Drug Discovery 2001, 29–34. 31 ROBERTS, G. C., Drug Discov. Today 2000, 5, 230–240. 32 MOORE, J. M., Curr. Opin. Biotechnol. 1999, 10, 54–58. 33 LEPRE, C. A., Drug Discov. Today 2001, 6, 133–140. 34 SHUKER, S. B., HAJDUK, P. J., MEADOWS, R. P., and FESIK, S. W., Science 1996, 274, 1531–1534. 35 HAJDUK, P. J., MEADOWS, R. P., and FESIK, S. W., Science 1997, 278, 497–499. 36 HAJDUK, P. J., DINGES, J., MIKNIS, G. F., MERLOCK, M., MIDDLETON, T., KEMPF, D. J., EGAN, D. A., WALTER, K. A., ROBINS, T. S., SHUKER, S. B., HOLZMAN, T. F., and FESIK, S. W., J. Med. Chem. 1997, 40, 3144–3150. 37 HAJDUK, P. J., ZHOU, M. M., and FESIK, S. W., Bioorg. Med. Chem. Lett. 1999, 9, 2403–2406. 38 HAJDUK, P. J., DINGES, J., SCHKERYANTZ, J. M., JANOWICK, D., KAMINSKI, M., TUFANO, M., AUGERI, D. J., PETROS, A., NIENABER, V., ZHONG, P., HAMMOND, R., COEN, M., BEUTEL, B., KATZ, L., and FESIK, S. W., J. Med. Chem. 1999, 42, 3852–3859.

39 HAJDUK, P. J., BOYD, S., NETTESHEIM, D., NIENABER, V., SEVERIN, J., SMITH, R., DAVIDSON, D., ROCKWAY, T., and FESIK, S. W., J. Med. Chem. 2000, 43, 3862–3866. 40 HAJDUK, P. J., GOMTSYAN, A., DIDOMENICO, S., COWART, M., BAYBURT, E. K., SOLOMON, L., SEVERIN, J., SMITH, R., WALTER, K., HOLZMAN, T. F., STEWART, A., MCGARAUGHTY, S., JARVIS, M. F., KOWALUK, E. A., and FESIK, S. W., J. Med. Chem. 2000, 43, 4781–4786. 41 ALLEN, K., BELLAMAINA, C., DING, X., JEFFERY, C., MATTOS, C., PETSKO, G., and RINGE, D., J. Phys. Chem. 1996, 100, 2605–2611. 42 NIENABER, V. L., RICHARDSON, P. L., KLIGHOFER, V., BOUSKA, J. J., GIRANDA, V. L., and GREER, J., Nat. Biotechnol. 2000, 18, 1105–1108. 43 ANDERSON, S. and CHIPLIN, J., Drug Discov. Today 2002, 7, 105–107. 44 GOODWILL, K., TENNANT, M., and STEVENS, R., Drug Discov. Today 2001, 6, S113–S118. 45 CARR, R. and JHOTI, H., Drug Discov. Today 2002, 7, 522–527. 46 BLUNDELL, T., JHOTI, H., and ABELL, C., Nat. Rev. Drug Discov. 2002, 1, 45–54. 47 STEWART, L., CLARK, R., and BEHNKE, C., Drug Discov. Today 2002, 7, 187–196. 48 BUSSOW, K., CAHILL, D., NIETFELD, W., BANCROFT, D., SCHERZINGER, E., LEHRACH, H., and WALTER, G., Nucleic Acids Res. 1998, 26, 5007–5008. 49 ARENKOV, P., KUKHTIN, A., GEMMELL, A., VOLOSHCHUK, S., CHUPEEVA, V., and MIRZABEKOV, A., Anal. Biochem. 2000, 278, 123–131. 50 ZHU, H., BILGIN, M., BANGHAM, R., HALL, D., CASAMAYOR, A., BERTONE, P., LAN, N., JANSEN, R., BIDLINGMAIER, S., HOUFEK, T., MITCHELL, T., MILLER, P., DEAN, R. A., GERSTEIN, M., and SNYDER, M., Science 2001, 293, 2101–2105. 51 MACBEATH, G. and SCHREIBER, S. L., Science 2000, 289, 1760–1763. 52 WARRIOR, U., BURNS, D., and KOFRON, J., Arrayed Compound Screening (ARCS): A Well-less Screening Platform with Advantages of Miniaturization. 2001. Eighth Annual HTT EXPO Advancing Drug Development 2001.

23

24 Chemical Microarrays for Small Molecule Ligand Screening

53 VENTER, J. C. et al., Science 2001, 291, 1304–1351. 54 FODOR, S. P.A., LEIGHTON, R. J., PIRRUNG, M. C., STRYER, L., LU, A. T., and SOLAS, D., Science 1991, 251, 767–773. 55 JACOBS, J. W. and FODOR, S. P., Trends Biotechnol. 1994, 12, 19–26. 56 FRANK, R., Tetrahedron 1992, 48, 9217–9232. 57 SCHARN, D., WENSCHUH, H., REINEKE, U., SCHNEIDER-MERGENER, J., and GERMEROTH, L., J. Comb.Chem. 2000, 2, 361–369. 58 MACBEATH, G., KOEHLER, A., and SCHREIBER, S. L., J. Am. Chem. Soc. 1999, 121, 7967–7968. 59 HERGENROTHER, P. J., DEPEW, K., and SCHREIBER, S. L., J. Am. Chem. Soc. 2000, 122, 7849–7850. 60 KURUVILLA, F. G., SHAMJI, A. F., STERNSON, S. M., HERGENROTHER, P. J., and SCHREIBER, S. L., Nature 2002, 416, 653–657. 61 SCHENA, M., SHALON, D., DAVIS, R. W., and BROWN, P. O., Science 1995, 270, 467–470. 62 LAM, K. S., LEBL, M., and KRCHNAK, V., Chem.Rev. 1997, 97, 411–448. 63 ULMAN, A., Chem. Rev. 1996, 96, 1533–1554. 64 MRKSICH, M. and WHITESIDES, G. M., Trends Biotechnol. 1995, 13, 228–235. 65 SIGAL, G. B., BAMDAD, C., BARBERIS, A., STROMINGER, J., and WHITESIDES, G. M., Anal. Chem. 1996, 68, 490–497.

66 MRKSICH, M., Journal of American Chemical Society 1995, 117, 12009–12010. 67 JUNG, L. et al., Langmuir 2000, 16, 9421–9432. 68 TSANG, S. K., CHEH, J., ISAACS, L., JOSEPH-MCCARTHY, D., CHOI, S. K., PEVEAR, D. C., WHITESIDES, G. M., and HOGLE, J. M., Chem. Biol. 2001, 8, 33–45. 69 HOUSEMAN, B. T., HUH, J. H., KRON, S. J., and MRKSICH, M., Nat. Biotechnol. 2002, 20, 270–274. 70 HODNELAND, C. D., LEE, Y. S., MIN, D. H., and MRKSICH, M., Proc. Natl. Acad. Sci.U. S. A 2002, 99, 5048–5052. 71 LAHIRI, J., ISAACS, L., GRZYBOWSKI, B., CARBECK, J., and WHITESIDES, G. M., Langmuir 1999, 15, 7186–7198. 72 LAHIRI, J., ISAACS, L., TIEN, J., and WHITESIDES, G. M., Anal. Chem. 1999, 71, 777–790. 73 MCDONNELL, J. M., Curr. Opin. Chem. Biol. 2001, 5, 572–577. 74 BAIRD, C. L. and MYSZKA, D. G., J. Mol. Recognit. 2001, 14, 261–268. 75 ABERY, J., Modern Drug Discovery 2001, 35–40. 76 RICH, R. L. and MYSZKA, D. G., J. Mol. Recognit. 2001, 14, 223–228. 77 RICH, R. L. and MYSZKA, D. G., J. Mol. Recognit. 2001, 14, 273–294. 78 MYSZKA, D. G., J. Mol. Recognit. 1999, 12, 390–408. 79 MORTON, T. A. and MYSZKA, D. G., Methods Enzymol. 1998, 295, 268–294. 80 COOPER M. A., Nat. Rev. Drug Disc. 2002, 1, 515–528.

1

Single Molecule Measurements of Molecular Motors Yoshiharu Ishii, and Toshio Yanagida

Single Molecule Processes Project, ICORP, JST, Osaka, Japan

Originally published in: Molecular Motors. Edited by Manfred Schliwa. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-52730594-0 Abstract

Single molecule detection was developed to measure the unique operation of molecular motors. The function of molecular motors has been ascribed to the movement of a single motor molecule along a partner protein track. Manipulating single protein tracks and single motors and imaging single molecules has allowed the behaviour of such unitary machines to be measured. The mechanical responses to the energy input from the hydrolysis of ATP molecules have been determined and the underlying mechanism explored. The results indicate that the movement of molecular motors is driven by thermal motion rather than structural changes occurring in the motor molecules. Thermal Brownian motion must be biased in one direction. In this article, we summarize the methods and results of single molecule measurements of molecular motors. 1 Introduction

Almost 30 years after the proposal of the sliding ﬁlament theory of muscle, the movement of actin ﬁlaments over myosin immobilized on coverslips was ﬁrst visualized under a microscope [1, 2]. This was accomplished by visualizing actin ﬁlaments labeled with phalloidin–tetramethylrhodamine by ﬂuorescence microscopy [3]. Phalloidin, a mushroom toxin, binds stoichiometrically to actin molecules and serves to stabilize the actin molecules in the ﬁlament form at low concentrations, at which point the motility assay and the single molecule measurements are performed. The in vitro assay has proven to be a model system in which both biochemical and mechanical properties of the interaction of myosin and actin can be studied using puriﬁed proteins. This system contrasts with conventional measurement systems which use single muscle ﬁbers and proteins in solution. The mechanical properties can be measured in a muscle ﬁber but the system is not Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Single Molecule Measurements of Molecular Motors

deﬁned biochemically. The biochemical properties can be measured in solution, while the mechanical properties cannot be studied. The sliding movement of actin and myosin can be basically ascribed to the operation of a unit machine that contains a single myosin molecule and an actin ﬁlament. In the in vitro assay, the smooth movement of an actin ﬁlament involves interactions with many myosin molecules. Thus, it is not possible to determine which myosin molecules interact with the actin ﬁlament and when and where ATP molecules are hydrolyzed. By manipulating single actin ﬁlaments and single myosin molecules, it has been possible to study the interaction between myosin and actin in a controlled manner. Examining the single unit machine offers many advantages over ensemble measurements. Myosin and actin function dynamically. Myosin undergoes a series of chemical reactions during ATP hydrolysis. It changes the interaction with actin cyclically in association with these chemical reactions. In addition, all biomolecules are inevitably exposed to thermal perturbations. These processes are stochastic. In ensemble measurements involving large number of molecules, only average values can be determined and the dynamic changes, which are important in relation to function, are hidden and obscured in the averaging process. Single molecule measurements have overcome these difﬁculties and have allowed the dynamic behavior of protein molecules to be measured [4].

2 Manipulation of Actin Filaments

Actin ﬁlaments can be captured and manipulated by a laser trap [5] or a glass microneedle [6]; Fig. 1. A laser trap is a technique used to manipulate protein molecules using the laser in a non-invasive manner. When a small bead is irradiated by a focused laser, the incident laser beam is deﬂected on the surface and radiation pressure is exerted. The total radiation pressure exerted on the entire sphere is always directed towards the focal point of the laser, so the beads are attracted to the focal point and follow the focal position of the laser when it is moved (laser trap). The beads also behave as if they are connected via a spring; i. e. the total trap force exerted on the beads is proportional to the distance from the focal point when their position deviates from the focal point (nanometry). Utilizing these properties of the laser trap, an actin ﬁlament can be manipulated via two beads attached at either end by a double trap, and the movement generated by myosin can be measured [5]; Fig. 1a. For the mechanical measurements, the actin ﬁlament is suspended taut and brought into contact with myosin molecules sparsely ﬁxed on the glass surface. The actin ﬁlaments with the beads at their ends move against the trap force in the presence of ATP. Thus, the displacement of the actin ﬁlaments generated by myosin can be measured by monitoring the changes in the position of the beads attached to them. The force generated can be estimated if the stiffness of the laser trap is known. The movement generated by single myosin molecules could be detected if the number of myosin molecules on the glass surface was reduced to one. The stiffness of the laser trap is dependent on the power of the laser. For a

3 Nanometry of Actin Filaments 3

Fig. 1 Manipulation of actin ﬁlaments. (a) An actin ﬁlament is manipulated via two beads attached at each end. (b) A microneedle is attached at one end of an actin ﬁlament to is attached at one end of an actin ﬁlament to manipulate it. For nanom-

etry, the displacement of the actin ﬁlament produced by single myosin molecules can be measured by recording the displacement of the beads or the tip of a microneedle with nanometer accuracy.

bead 1 µm in diameter, the trap force is several tens of pN and the stiffness is ∼0.1 pN nm−1 when the power of the laser is several hundred mW. This stiffness is sufﬁcient to measure the movement caused by a single myosin molecule. The displacement generated with a single ATP molecule is approximately 10 nm. Thus, the force will be less than 8 pN because the free energy released from the hydrolysis of a single ATP molecule is ∼80 pN nm−1 . Microneedles are another method used to manipulate actin ﬁlaments. Thin glass microneedles can be prepared and attached to the ends of the actin ﬁlaments [6] Fig. 1b. The premise of the measurement using microneedles is the same as that for the laser trap. When actin ﬁlaments move as a result of the interaction with myosin, the microneedle bends and at the same time a restoring force is applied to the microneedle resisting the movement. The strength of the force is proportional to the displacement, with the stiffness of the microneedle as a proportional constant. When the force caused by myosin is balanced with the restoring force applied by the microneedle, the movement ceases. The displacement of the actin ﬁlaments can be measured and the force can be estimated when the stiffness of the microneedle is known. The stiffness of a microneedle is dependent on its shape. Long, thin glass microneedles are more ﬂexible. A microneedle 50 µm in length and 0.3 µm in diameter, which is the typical length used for the measurements, gives a stiffness of 1 pN nm−1 .

3 Nanometry of Actin Filaments

Given that the displacement of the molecular motors is in the order of nanometers, the change in the position of the beads or the tip of the microneedle must be measured with nanometer accuracy. The image of the bead or the tip of the microneedle

4 Single Molecule Measurements of Molecular Motors

can be enlarged approximately 1000 times and projected onto a pair of photodiodes. The change in the position of the bead or tip of the microneedle results in a change in the difference of the intensities of each photodiode. Therefore, the difference in signals between multiple photodiodes is measured to estimate the displacement of the bead or microneedle. Thus the displacement can be determined with nanometer accuracy, which is much less than the diffraction limit. The diffraction limit is the minimum distance that two spots can be apart to be resolved as separate in the light microscope. As demonstrated in these measurements, it is possible to trace a moving spot with an accuracy greater than the diffraction limit. In these measurements, the displacement of the actin ﬁlaments is determined by measuring the displacement of a marker such as a bead or microneedle. If the connection between the myosin molecules and the beads or the microneedle is not tight and some compliant linkages exist between them, damping of the actual displacement of myosin and thermal ﬂuctuations can occur. In addition, the position of the bead or microneedle is subject to thermal ﬂuctuation. The mean square displacement x2 of a bead or microneedle can be related to the stiffness of the laser trap or microneedle, K, by the equation K x2 /2 = kB T/2. According to this equation, the root mean square distance is 6.43 nm for a stiffness of 0.1 pN nm−1 and 2.03 nm for a stiffness of 1 pN nm−1 . To measure the displacement with nanometer accuracy, it is important to make the system as stiff as possible. The beads or microneedle may respond slowly to the fast movements of the protein molecules. This response determines the time resolution. The response time of the system, τ , is related to the friction coefﬁcient β and the stiffness K as τ = β/K. The friction coefﬁcient depends on the shape and size of the probe. For a long and ﬂexible glass microneedle with a diameter of 1 µm and a length of 1 cm, (stiffness = 0.01 pN nm−1 ), for example, the temporal resolution is only 1 s. To increase the temporal resolution, a short thin glass microneedle is necessary. For a microneedle with a length of 50 µm and a diameter of 0.3 µm (which gives a stiffness of 1 pN nm−1 ), the temporal resolution can be increased to almost 1 ms. In contrast, the viscous drag on a bead is small because a bead is smaller than a microneedle. For a bead 1 µm in diameter, the temporal resolution is as good as 0.1 ms. In addition to this advantage of time resolution, the laser trap experiments are much easier to carry out than microneedle experiments. The stiffness of the laser trap can be easily changed by changing the laser power. In the microneedle measurements, microneedles must be individually handcrafted and the stiffness of each individual microneedle must be determined.

4 Movement of Actin Filaments Caused by Single Myosin Molecules

The actin ﬁlament repeats steps and returns to its original position, while myosin repeats the binding to and dissociation from the actin ﬁlament (Fig. 2). The proﬁle of the displacement trace is dependent on the ATP concentration. At high concentrations of ATP (in the mM range) the actin ﬁlaments showed rapid transient

4 Movement of Actin Filaments Caused by Single Myosin Molecules 5

Fig. 2 Time course of the displacement of actin ﬁlaments produced by single myosin molecules. Upper trace: the white line represents the raw data and the black line the same data after ﬁltration with a lowpath ﬁlter (20 Hz bandwidth). The stiffness was calculated from the variance of bead ﬂuctuations (bottom panel).

movement. As the ATP concentration decreased to the order of µM, the duration of the displacement became longer, while the size of the displacement remained the same. At very low ATP concentrations (in the nM range), individual events caused by a single myosin and ATP molecule can be identiﬁed. The duration time of the displacement increased from ∼0.2 s at 1 µM ATP to ∼20 s at 10 nM ATP. The ATP concentration-dependent trace of the displacement has been interpreted as the mechanical step in relation to the chemical reaction of ATP hydrolysis. The ATP binding step is dependent on the ATP concentration. The time for ATP molecules to bind is longer at low ATP concentrations and much shorter at high ATP concentrations. This is thought to be the mechanism by which single molecules ‘learn’ the ATP concentration of the system. The binding of ATP to myosin triggers the dissociation of myosin from the actin ﬁlament resulting in the bead returning to its original position. Myosin actually moves the actin ﬁlament when the products of ATP hydrolysis, ADP and Pi, are released. The relationship between the mechanical events and the chemical reaction has been conﬁrmed directly by simultaneously measuring the movement of the motor and the turnover of ATP molecules, as outlined below. One important parameter that the mechanical measurements of myosin have provided is the size of the myosin step (step size) caused during the hydrolysis of a single ATP molecule [7, 8]. The position of the bead ﬂuctuated thermally around a mean position, which was dependent on the stiffness of the trap as mentioned above [9]. Once myosin was strongly bound to the actin ﬁlament, the stiffness of the system increased and the thermal motion of the beads greatly decreased. By measuring the variance of the displacement trace, it is therefore possible to determine when myosin interacts with actin. However, the size of the step could

6 Single Molecule Measurements of Molecular Motors

not be determined for individual displacements, because the starting position of the displacements cannot be measured. The displacement relative to the mean position of the bead has a wide distribution, reﬂecting the thermal ﬂuctuation of the bead. Using this technique only mean displacements can be obtained. The size of the displacement caused by a single myosin molecule upon ATP hydrolysis varies among research groups, the reported values ranging from 5 to 20 nm [5, 9–11]. Many experimental factors can affect the results of the step size. In the measurements, myosin molecules are immobilized on the glass surface. These immobilization procedures may affect the activity of myosin. Direct attachment of the myosin head to the glass surface may alter its activity as suggested by the results of in vitro motility assays [12]. To avoid the direct artiﬁcial attachment of the head portion of the myosin to the glass surface, the head is placed in a myosin–myosin rod co-ﬁlament, which can be prepared so as to contain only one or two heads in a single ﬁlament 5–8 µm in length. The use of such ﬁlaments has an additional advantage. It has been reported that the step size is dependent on the angle between the myosin and the actin ﬁlament [11]. The step size is maximal when the two ﬁlaments are parallel, but approaches zero when the two ﬁlaments are positioned at right angles. In the myosin–myosin rod ﬁlament, the orientation of the myosin head relative to the actin ﬁlament is known. In the situation where the myosin head molecules are placed in a random orientation, the displacement of actin ﬁlaments would range between the maximum value at an angle of 0◦ and the minimum value at an angle of 90◦ [13]. The average value would be less than the maximum value.

5 Visualization of Single Molecules

Biomolecules including protein molecules and small chemical compounds such as ATP are much smaller than the diffraction limit. It is impossible to visualize them using conventional light microscopy. In order to visualize such small molecules they are labeled with ﬂuorescent probes and then the ﬂuorescence is monitored. In 1995 a single ﬂuorophore attached to a biomolecule in aqueous solution was ﬁrst visualized after the background noise was greatly reduced by eliminating the noise from different parts of the microscope [14]. Many microscopes have also been developed to decrease the background noise. The basic concept behind these microscopes is that only local regions are illuminated to excite the molecules of interest. To visualize the behavior or motion of biomolecules at work, realtime imaging in aqueous solution is also a prerequisite. Total internal reﬂection ﬂuorescence microscopy (TIRFM) is a technique which illuminates only the surface area between the solution and the coverslip, allowing both local illuminating and real-time imaging [14]; Fig. 3). When incident laser light is above the critical angle, the light is totally reﬂected at the interface with the aqueous solution, and an evanescent ﬁeld is created 100 to 200 nm from the surface of the aqueous solution. Thus, only the ﬂuorescent molecules positioned near the glass surface are excited

6 Visualization of ATP Turnover and Mechano-chemical Coupling

Fig. 3 Total internal reﬂection ﬂuorescence microscopy. An evanescent ﬁeld is created within 100–200 nm from the interface between the glass surface and the solution when the laser light strikes at an angle greater than the critical angle. Only molecules in the vicinity of the glass surface are excited and emit ﬂuorescence.

and emit ﬂuorescence. Large numbers of ﬂuorescent molecules outside the region are not excited and therefore the background noise is greatly reduced. A single ﬂuorophore is imaged as a spot, but the size of the spot appears to be enlarged to several hundreds of nanometers due to diffraction, even though the size of the molecule is in the order of a few nanometers. Because of this, spatial resolution is limited. While the ﬂuorophores are continuously illuminated, single ﬂuorophores are repeatedly excited and emit ﬂuorescence within nanoseconds. A single spot therefore represents the integration of a large number of photons emitted from a single ﬂuorophore. Single ﬂuorophores can suddenly cease to emit ﬂuorescence after a certain amount of exposure to irradiation, because they become irreversibly damaged as a result of the photochemical reaction. The average time for a photobleaching process to occur depends on the power of the laser. When a laser with higher power is used, photobleaching occurs in a short time, although the ﬂuorescence is initially brighter. The single stepwise drop of the ﬂuorescence is a good test to conﬁrm that the ﬂuorescent spot was from a single ﬂuorophore. Fluorescence intensity can also be used to determine whether the ﬂuorescence spots arise from single molecules. Histograms of the ﬂuorescence intensity from single spots show a large peak corresponding to multiples of the minimum intensity of a single molecule.

6 Visualization of ATP Turnover and Mechano-chemical Coupling

Fluorescent spots remain in the same position when single biomolecules are immobilized on a glass surface and their motion can be traced only when they move slowly (not greater than tens of µm s−1 ). Rapidly moving ﬂuorophores cannot be

7

8 Single Molecule Measurements of Molecular Motors

Fig. 4. Imaging of the turnover of individual ﬂuorescent ATP molecules produced by the interaction of single myosin molecules in a myosin ﬁlament with a single actin ﬁlament. (Top) a picture for an actin ﬁlament trapped by a laser via two beads attached at both ends and a myosin–myosin-rod co-ﬁlament. (Second) ﬂuorescence image of myosin and actin ﬁlament. The lower panels show sequential images of an association and dissociation event of ATP (ADP) with the same myosin head.

traced and only contribute to the background noise. The binding of ﬂuorescent ATP to the protein molecules can be visualized in single molecule imaging, because the mobility of bound ﬂuorescent ATP is different from that free in solution [14]; Fig. 4). ATP can be chemically modiﬁed with a ﬂuorescent probe. The ﬂuorescent ATP molecules can be visualized as spots when bound to immobilized myosin. However, ﬂuorescent molecules cannot be observed when they are free in solution due to the rapid Brownian movement. The time span between binding and dissociation has a characteristic distribution, reﬂecting stochastic behavior. The histogram of the time course can be ﬁtted to an exponential curve and the decay time (dissociation rate) can be compared with ATPase rates measured in solution at saturating concentrations of ATP. This assay allows the measurement of ATP binding at the single molecule level. However, the range of concentrations of ﬂuorescent molecules is limited, because all ﬂuorescent ATP molecules in solution contribute to the background noise even when evanescent illumination is used. Typically ﬂuorescent ATP is used at concentrations between 10 and 20 nM. Of course, the binding afﬁnity must be high enough to observe binding at such low concentrations. Given that both mechanical and chemical processes can be measured at the single molecule level, the coupling between them can be directly determined by simultaneous measurements [15]. The direct determination of the coupling is possible

6 Visualization of ATP Turnover and Mechano-chemical Coupling

Fig. 5 Direct determination of the coupling between chemical and mechanical events. The ATP turnover monitored by the ﬂuorescence of Cy3-ATP (bottom panel) and the movement of myosin (top panel) were simultaneously measured at the single molecule level. The dissociation of myosin

from actin ﬁlaments (decrease in the displacement) is coupled with the binding of ATP (increase in ﬂuorescence) and the generation of movement (increase in the displacement) is coupled with the dissociation of the products of the ATP hydrolysis reaction (decrease in ﬂuorescence).

only when single molecule detection techniques are used, as individual chemical and mechanical events are not averaged. The data of the simultaneous measurements of the biochemical and mechanical events at low time resolution (∼30 ms) have shown that the chemical reaction and the mechanical events appear to be coupled (Fig. 5). When ATP binds, myosin dissociates from actin and myosin generates the step movement upon ADP dissociation. The coupling between the binding of ATP and the dissociation of myosin has been conﬁrmed at higher time resolution (< 10 ms). However, a time difference between the ADP release and the generation of movement became apparent. This difference had a particular distribution at higher time resolution. In half the cases, the generation of the displacement occurred at the same time as the release of ADP, while in the other half the mechanical displacement was delayed by < 0.5 s after the release of ADP. It should be noted that it is not possible to experimentally distinguish photobleaching from the dissociation of ﬂuorescent ADP in a single event. In both cases, the ﬂuorescent spots disappear suddenly. The laser power can be decreased to

9

10 Single Molecule Measurements of Molecular Motors

decrease the rate of photobleaching and the probability that the photobleaching occurs before the dissociation takes place becomes lower. In these experiments the turnover of ﬂuorescent ATP by myosin and the decay time for the photobleaching of Cy3-ATP either directly attached to the glass or bound to myosin in the presence of vanadate, can be compared with the turnover time. Thus, the probability that photobleaching occurs prior to nucleotide release can be reduced to a few percent. The free energy released at the step where the products of ATP hydrolysis are dissociated, must be stored until the generation of the displacement. The storage of the energy, or a prolonged memory effect, can be explained with the existence of two or more different conformational states, between which the protein undertakes slow transitions. Evidence for these multiple conformational states and spontaneous slow transitions between them has come from the studies of single molecule spectroscopy for many proteins including myosin [16]. Protein molecules have several local minimal states with high-energy barriers between them.

7 Visualization of the Movement of Single Kinesin Motors

The visualization of single molecules also allows the observation of the movement of single ﬂuorescently-labeled motor molecules along a protein track. For visualization, the motor molecules must move for long distances without dissociating. Kinesin, a microtubule-based motor, is one such processive motor. The movement of single molecules of ﬂuorescently-labeled kinesin along microtubules has been visualized [17]. A single kinesin molecule moves for ∼500 nm in 1 ∼2 s. The velocity is similar to that of microtubules moving over kinesin molecules attached to a coverslip. The processive movement of kinesin has been explained on the basis of its double-headed structure; one of the two heads moves one step while the other head stays attached to the microtubule. The two heads operate cooperatively and the steps alternate. If this is the case, then single-headed kinesin should not be able to move for long distances. However, single molecule imaging showed that some of the single-headed kinesin molecules did in fact move along the microtubules for long distances, indicating that two heads are not essential for processive movement (Fig. 6). The data analysis of the time course of the displacement has shown that the movement of single-headed kinesin resulted from diffusion and was biased in one direction [18, 19]. For these single-headed kinesin molecules it has been suggested that additional interactions with microtubules occur so that the motor protein molecules are kept on the track of microtubules during movement. KIF1A, a single-headed motor of the kinesin superfamily, has a lysine-rich positivelycharged loop (the K loop) that is thought to interact with negative charges on the C-terminus of microtubules. Deletion mutants of single-headed kinesin also showed diffusion-driven movement and only traveled small distances of 50 nm in less than 1 s. The travel distance and interaction time were enhanced to 500 nm and 2 s, respectively, when they were fused with a peptide called biotin-dependent

7 Visualization of the Movement of Single Kinesin Motors 11

Fig. 6 Movement of single kinesin molecules along microtubules. The trace of the ﬂuorescently-labeled kinesin was recorded and the displacement along microtubules is plotted as a function of time. K351 is a kinesin construct which lacks the

C-terminal coiled-coil region (single headed kinesin) and biotin-dependent transcarboxylase (BDTC) is fused to K351 (K351BDTC). K411 is a double headed kinesin which shows processive movement over a long distance in one direction.

transcarboxylase (BDTC) which interacts with microtubules. BDTC does not affect the movement of two-headed kinesin. In the images of the movement of single kinesin molecules, we have frequently observed more than two kinesin molecules moving simultaneously along a single microtubule. The images have shown that there is a tendency for several kinesin molecules to move together suggesting that there is some communication between kinesin molecules via the microtubules [20]. Microtubules may be ‘activated’ by the binding of a single kinesin molecule to assume a high afﬁnity state, resulting in cooperative binding of kinesins. However, although the movement of single molecules has been visualized, it has not been possible to observe the unitary movement of kinesin due to limitations in the accuracy of single molecule imaging techniques. The experimental accuracy of imaging is restricted to several hundred nm due to the diffraction limits and the video rate of 1/30 of a second. These limitations have been overcome using a laser trap system where it has been possible to observe the stepwise movement

12 Single Molecule Measurements of Molecular Motors

of kinesin. The use of ﬂuorescent beads instead of single ﬂuorophores enhanced the contrast of the images. The laser trap ‘cooled’ the thermal motion of the beads and the use of a pair of photodiodes allowed the measurement of the displacement to be undertaken with nanometer accuracy [21]. Kinesin attached to a bead was brought into contact with microtubules in the presence of ATP. The number of kinesin molecules attached to a bead was estimated statistically. The distribution of kinesin molecules on beads was consistent with Poisson statistics as indicated by the probability of a moving bead at various molar ratios of kinesin to beads. The average number of kinesin molecules on a bead used for the experiments and the probability that more than two kinesin molecules attached to a bead can be estimated. Kinesin moves along microtubules in regular steps of 8 nm (Fig. 7). The steps are primarily in the forward direction and only occasionally in the backward direction. As kinesin molecules moved against an increased load produced by the laser, the average time of a step as well as the number of the backward steps increased. Eventually, at a stall force of 7–8 pN the number of steps in the forward and backward directions became equal. When this occurred the molecule dissociated from the microtubules and returned to its original position. Both forward and backward movements occurred at the same chemical state; the single cycle of the ATP hydrolysis was coupled to either forward or backward steps and the direction could be determined stochastically. Thus, the thermal movements, which have been clearly observed for single-headed kinesin, are the driving force even for the stepwise movement of kinesin.

Fig. 7 Stepwise movement of kinesin attached to beads. The displacement of kinesin can be monitored by the laser trap method. The displacement trace shows 8-nm steps. Backward movement was observed at high load.

8 Visualization of the Processive Movement of Single Myosin Motors

Substeps of the 8-nm step of kinesin have been uncovered using laser trap measurements with small beads [22]. Decreasing the size of the beads improved the time resolution, although the images of the bead on the detector became fainter. Dark ﬁeld illumination was used to brighten the images. The 8-nm step could be resolved into fast and slow substeps, each corresponding to a displacement of ∼4 nm.

8 Visualization of the Processive Movement of Single Myosin Motors

In the case of muscle myosin, it was impossible to detect the sliding movement with the imaging techniques described above, because muscle myosin readily dissociates from actin ﬁlaments at every cycle of the ATPase reaction. Recently, different types of myosin behavior have been discovered. Myosin V, VI and IX were found to move processively and it has been possible to visualize the movement of single myosin molecules when they are fused to green ﬂuorescent protein (GFP; Fig. 8). Myosin VI, for example, moves 240 nm along a single actin ﬁlament in 0.44 s [23]. Interestingly, the directionality of myosin VI is opposite to the direction of other myosins [24]. One of the ends of the actin ﬁlament (pointed end) was ﬂuorescently marked and the direction of the movement of myosin could be conﬁrmed for individual myosin molecules.

Fig. 8 Processive movement of myosin V and myosin VI. The movement of myosins V and VI was visualized by means of green ﬂuorescent protein (GFP). Sequential images indicate the movement of these

myosin molecules. Myosin V and myosin VI move in opposite directions. The pointed end of the actin ﬁlament was ﬂuorescently labeled.

13

14 Single Molecule Measurements of Molecular Motors

Given that myosin V and VI are processive motors, displacement records were obtained by trapping these myosins like kinesin, as well as by double trapping the actin ﬁlaments. A consecutive step movement was observed which was consistent with other processive motors. The advantage of processive myosins is that it is possible to determine the size of individual steps without disturbance from thermal ﬂuctuations, with the exception of the ﬁrst step. Measuring the step size of myosin V and VI allowed the testing of a model developed on the basis of structural studies of myosin heads. In this so-called lever arm model, a long α-helix (the neck domain) connecting the motor domain and the tail domain is thought to rotate relative to the motor domain, thus acting as a lever [25]. This model predicts that the size of the displacement would be proportional to the length of the neck domain, or lever arm. Myosin V has a long neck domain, which contains six repeats of the calmodulin or light chain binding site as compared to the two binding sites in muscle myosin. The step size of myosin V was shown to be 36 nm, while that of muscle myosin is 5 ∼20 nm, a ﬁnding in agreement with the lever-arm model [26]. However, a deletion mutant, in which the neck domain is shortened to only one calmodulin-binding site, performed the same large step as the original myosin V motor [27]; Fig. 9. A similar result has been obtained using myosin VI. Myosin VI in its natural form has only one calmodulin-binding site and also produces a large step [23, 28]. These observations of large steps in myosins with a short lever arm cannot be explained by the lever-arm model. A step size of 36 nm matches the helical pitch of the two-stranded actin ﬁlament, indicating that the architecture

Fig. 9 The step movement of myosin V and its deletion mutant. The displacement was measured by a double trapping method. (Top) wild type myosin V. (Bottom) the mutant was obtained by deleting a large part of the neck domain.

9 Manipulation of Single Myosin Molecules with a Scanning Probe and Nanometry 15

of actin ﬁlaments plays an important role in the movement of myosin. Thus, the relationship between structure and function can be studied by combining single molecule detection with cell biology and protein engineering. Even if the structural change, i. e. swinging of the lever arm, occurs as predicted based on the X-ray crystallographic studies, these studies indicate that the structural change does not drive the mechanical movement. The structural change of the neck domain may play another role, for example in regulating the kinetic steps as a strain sensor.

9 Manipulation of Single Myosin Molecules with a Scanning Probe and Nanometry

In order to measure the details of the displacement of motors, we have developed a new technique in which single muscle myosin molecules instead of actin ﬁlaments are captured and manipulated [29]; Fig. 10. A single myosin head fragment, S1, was attached to a scanning probe and the displacement of the probe along immobilized actin ﬁlaments was measured. The probe was brought into contact with actin bundles. The use of actin bundles instead of ﬁlaments makes the manipulation for the interaction easier and the system stiffer. The stiffness of this system was 1 pN nm−1 compared to a stiffness of 0.2 pN nm−1 for the laser trap system, given the root mean square displacement of 2.0 nm for the scanning probe system as compared to 4.5 nm for the laser trap system. The response time was < 0.4 ms. This improvement was critical for detecting the details of the movement of single myosin molecules. To capture single myosin S1 molecules, they are marked by attaching a ﬂuorescent dye to them. The ﬂuorescently-labeled S1 molecules were sparsely placed on a glass slide and scanned using a probe. As a scanning probe, a whisker made from ZnO and 10 nm in diameter with sharp edges, was attached to the tip of a thin glass needle. A single myosin S1 molecule was caught by the tip of the scanning probe.

Fig. 10. Measurement of 5.5-nm steps in the displacement generated by myosin S1. A single myosin S1 molecule was attached to the tip of a cantilever and brought into contact with actin ﬁlaments. Given that this system is stiffer than the laser trap system, steps in the rising phase of the displacement record are identiﬁed. Backward steps have been occasionally observed, as indicated by an arrow.

16 Single Molecule Measurements of Molecular Motors

Conﬁrmation that only a single S1 molecule had been captured, was achieved by monitoring the ﬂuorescence intensity and the photobleaching behavior. The possibility that non-ﬂuorescent myosin molecules and molecules which were labeled but had already been photobleached, were attached in addition to a single ﬂuorescent myosin molecule cannot be ruled out. However, the probability that additional non-ﬂuorescent molecules were attached was estimated to be marginal based on the labeling ratio of myosin after all experimental precautions had been taken. Myosin S1 was attached tightly to a probe via the biotin–avidin system. Given that this molecular glue was attached to the light chain of S1, which is a considerable distance from both the catalytic domain and the binding domain for actin on the myosin head, the attachment to the probe is thought not to interfere with motor functions. The displacement record of muscle myosin measured with this system was indistinguishable from the trace of actin ﬁlaments obtained from double trap measurements, if the data were plotted on the same time scale (Fig. 10). The rising phase of the displacement could be expanded, because the time resolution was considerably better for the scanning probe method. Within a single displacement during the hydrolysis of a single ATP molecule, there was stepwise movement. In a single displacement, there were one to ﬁve regular steps, giving rise to a total displacement of ∼5 to 30 nm. The average number of steps in a single displacement was three, corresponding to a total displacement of 13 nm. The size of a single step was 5.5 nm, coinciding with the interval between adjacent actin monomers on one strand of an actin ﬁlament.

10 Biased Brownian Movement

In addition to the variation in the total number of steps in a single displacement, evidence has been obtained to suggest that the rising phase of the displacement is stochastic. The interval time between steps also varied and was independent of the ATP concentration, whereas the time between the displacements was ATP concentration-dependent. The dwell time between steps was dependent on the temperature so that the 5.5-nm steps could be distinguished more clearly at lower temperatures. Approximately 10% of the total number of steps was in the backward direction. Backward movements increased with an increase in the load. In the experimental system used, the load applied could be increased by increasing the stiffness of the microneedle. At high loads, the number of steps in a single displacement was smaller and the interval time between the steps was larger, while the size of the steps remained the same. Thus, the reciprocal of the interval time between the steps decreased with an increase in the load. This relationship is basically similar to the force–velocity relationship in muscle, indicating that the unit machine consisting of a single myosin molecule and a single actin ﬁlament is sufﬁcient to explain the mechanical properties of muscle.

11 Conclusions 17

The available data indicate that the sliding movement of myosin is driven by thermal movement. However, thermal Brownian motion is random. The direction of the movement must be biased and the energy released from ATP hydrolysis is used to bias it. Otherwise, the second law of thermodynamics would be violated. In both myosin and kinesin, the directionality of the movement can be explained by a difference in the activation energy between forward and backward movement. The probability of movement is proportional to exp(−/kB T) when is the activation energy, kB is Boltzmann’s constant, and T is the absolute temperature. The ratio of the probability for forward and backward movement r = nforward /nbackward can be written by the difference in the activation energy between forward and backward movement, as r = exp(−/kB T), where = forward – backward . Analysis of the data shows that the difference in the activation energy was 2 ∼ 3 kB T for myosin and 5 kB T for kinesin at zero load. A model simulation further shows that the directionality ratio can reach 9:1 when the potential difference between neighboring actin monomers is only 10% of the energy barrier between neighboring actin monomers. The question arises as to how this direction-dependent potential proﬁle is created. One interesting idea is that the binding of a myosin molecule may activate several neighboring actin monomers in a direction-dependent manner, so that the myosin molecule is attracted more to one direction than to the other. In the case of the kinesin–microtubule system, such cooperativity in the binding was demonstrated based on the imaging of the binding of single kinesin molecules along microtubules, as described above. The binding of kinesin molecules was induced to a greater extent in front of the moving kinesin molecule. In this model, a statistically biased potential is assumed to cause the biased motion. Another explanation for biased Brownian movement is based on dynamic ﬂuctuations of the potential. Myosin and actin dynamically change their structure and the interactions that occur between them. It has been shown that the movement of Brownian particles under the asymmetrical potential is biased by ﬂuctuating potential unless the ﬂuctuation is random. There is an optimum frequency at which the movement is biased. In the model where a structural change results in movement, different structures of myosin are assumed to correspond to the different states of the ATPase cycle and the structural change is associated with the transition between these states. In contrast, in the biased Brownian model, the movement is thought to occur in one state (ADP.Pi state) in the cycle of ATP hydrolysis. Thus, it is possible to simulate the movement of myosin.

11 Conclusions

New ﬁndings in the ﬁeld of biosciences have been frequently initiated by the development of new techniques. Single molecule detection has allowed us to monitor the dynamic behavior of biomolecules, which is hidden in the average values of

18 Single Molecule Measurements of Molecular Motors

ensemble measurements where it is not possible to measure the sequence of the mechanical and biochemical events. Single molecule detection techniques have completely changed the style of research in this ﬁeld. Combining this technique with the techniques of protein engineering and cell biology, the possibility for new types of measurements has increased. However, many unanswered questions still remain and additional new questions have been raised. There is still room for further improvement of established techniques and the development of new ones. For example, ﬂuorescent dyes with greater stablity would be very useful. In addition, experimental systems that have increased stiffness would increase the experimental resolution for nanometry. Molecular motors are typical molecular machines. Although protein molecules of nanometer size are exposed to thermal agitation, they function very efﬁciently. There is increasing evidence to suggest that these molecules do not operate against thermal perturbation, but rather harness thermal energy to perform their functions. We have started measuring the behavior of molecular machines in which the biomolecules assemble and interact. Single molecule detection will be the technique of choice to unveil the underlying mechanisms.

References 1 HARADA Y., K. SAKURADA, T. AOKI, D. D. THOMAS, and T. YANAGIDA. 1990. Mechanochemical coupling in actomyosin energy transduction studied by in vitro movement assay J. Mol. Biol. 216: 49–68. 2 KRON, S. J. and J. A. SPUDICH. 1986. Fluorescent. actin ﬁlaments move on mysin ﬁxed to a glass surface. Proc. Natl. Acad. Sci. USA 83: 6272–6276. 3 YANAGIDA T., M. NAKASE, K. NISHIYAMA, and F. OOSAWA. 1984. Direct observation of motion of single F-actin ﬁlaments in the presence of myosin. Nature 307: 58–60. 4 ISHII Y. and T. YANAGIDA. 2000. Single molecule detection in life science Single Mol. 1: 5–16. 5 FINER J. T., R. M. SIMMONS, and J. A. SPUDICH. 1994. Single myosin molecule mechanics: Piconewton forces and nanometre steps. Nature 368: 113–119. 6 ISHIJIMA, A., H. KOJIMA, H. HIGUCHI, Y. HARADA, T. FUNATSU, and T. YANAGIDA. 1996. Multiple and single-molecule analysis of the actomyosin motor by nanometer-piconewton manipulation

7

8 9

10

11

12

with a microneedle: Unitary steps and forces. Biophys. J. 70: 383–400. ISHII, Y., S. ESAKI, and T. YANAGIDA. 2002. Ex perimental studies of the myosin–actinmotor. Appl. Phys. A 75: 325–330. SPUDICH J. A. 1994. How molecular motors work. Nature 372: 515–518. MOLLOY J. E., J. E. BURNS, J. KENDRICK-JONES, R. T. TREGEAR, and D. S. C. WHITE. 1995. Movement and force produced by a single myosin head. Nature 378: 209–212. GUILFORD, W. H., D. E. DUPUIS, G. KENNEDY, J. B. PATLACK, and D. M. WARSHAW. 1997. Smooth muscle and skeletal muscle myosin produce similar unitary forces and displacement fully equipped with factors essential for motor function. Biochem. Biophys. Res. Commun. 230: 76–80. TANAKA H., A. ISHIJIMA, M. HONDA, K. SAITO, and T. YANAGIDA. 1998. Orientation dependence of displacements by a single one-headed myosin relative to the actin ﬁlament. Biophys. J. 75: 1886–1894. IWANE, A. H., K. KITAMURA, M. TOKUNAGA, and T. YANAGIDA. 1997. Myosin

References

13

14

15

16

17

18

19

20

21

subfragment-1 is Biophys. J. 72: 1006–1021. YANAGIDA, T., K. KITAMURA, H. TANAKA, A. H. IWANE, A. ISHIJIMA, and S. ESAKI. 2000. Single molecule analysis of the actomyosin motor. Curr. Opin. Cell Biol. 12: 20–25. FUNATSU T., Y. HARADA, M. TOKUNAGA, K. SAITO, and T. YANAGIDA. 1995. Imaging of Single Fluorescent Molecules and individual ATP turnovers by single myosin molecules in aqueous solution. Nature 374: 555–559. ISHIJIMA A., H. KOJIMA, T. FUNATSU, M. TOKUNAGA, H. HIGUCHI, and T. YANAGIDA. 1998. Simultaneous measurement of chemical and mechanical reaction. Cell 92: 161–171. WAZAWA T., Y. ISHII, T. FUNATSU, and T. YANAGIDA. 2000. Spectral sluctuation of a single ﬂuorophore confugated to a protein molecule. Biophys. J. 78: 1561–1569. VALE R. D., T. FUNATSU, D. W. PIERCE, L. ROBMERG, Y. HARADA, and T. YANAGIDA. 1996. Direct observation of single kinesin molecules moving along microtubules Nature 380: 451–453. INOUE Y., A. H. IWANE, T. MIYAI, E. MUTO, and T. YANAGIDA. 2001 Motility of single one-headed kinesin molecules along microtubules. Bio phys. J. 81: 2838–2850. OKADA Y. and N. HIROKAWA. 1999. A processive single-headed motor: Kinesin superfamily protein KIF1A. Science 283: 1152–1157. MUTO, E. 2001. Is microtubule an active participant in the mechanism of motility? Biophys. J. 80: 513a. SVOBODA, K., C. F. SCHMIDT, B. J. SCHNAPP, and S. M. BLOCK. 1993. Direct observation of kinesin stepping by optical trapping interferometry. Nature 365 721–727.

22 NISHIYAMA, M., E. MUTO, Y. INOUE, T. YANAGIDA, and H. HIGUCHI. 2001. Substeps within the 8-nm step of the ATPase cycle of single kenisin molecules. Nature Cell Biol. 3: 425–428. 23 NISHIKAWA S., K. HOMMA, Y. KOMORI, M. IWAKI, T. WAZAWA, A. H. IWANE, J. SAITO, R. IKEBE, E. KATAYAMA, T. YANAGIDA, and M. IKEBE. 2002. Class VI myosin moves processively along actin ﬁlaments backward with large steps. Biochem. Biophys. Res. Commun. 290: 311–317. 24 WELLS A. L., A. W. LIN, L.-Q. CHEN, D. SATER, S. M. CALN, T. HASSON, B. O. CARRAGHER, R. A. MILLIGAN, and H. L. SWEENEY. 1999. Myosin VI is an actin-based motor that moves backwards. Nature 401: 505–508. 25 RAYMENT I., W. R. RYPNIEWSKI, K. SCHMIDS-BASE, R. SMITH, D. R. TOMCHICK, M. M. BENNING, WINKELMANN, D., A. G. WESENBERG, and H. M. HOLDEN. 1993. Three-dimensional structure of myosin subfragment-1: a molecular motor. Science 261: 50–58. 26 MEHTA, M., R. S. ROCK, A. D. M. RIEF J. A. SPUDICH, R. S. MOOSEKER, and R. E. CHENEY. 1999. Myosin-V is a processive actin-basedmotor. Nature 400: 590–593. 27 TANAKA H., K. HONMA, A. H. IWANE, E. KATAYAMA, R. IKEBE, J. SAITO, T. YANAGIDA, and M. IKEBE. 2001. The motor somain determines the large step of myosin-V Nature 415: 192–195. 28 ROCK R. S., S. E. RICE, A. L. WELLS, T. J. PURCELL, J. A. SPUDICH, and H. L. SWEENEY. 2001. Myosin VI is a processive motor with a large step size. Proc. Natl. Acad. Sci. USA 98: 3655–13659. 29 KITAMURA K., M. TOKUNAGA, A. H. IWANE, and T. YANAGIDA. 1999. A single myosin head moves along an actin ﬁlament with regular steps of 5.3 nanometres. Nature 397: 129–134.

19

1

Analysis of Receptor-Ligand Interaction M. M. H¨oﬂiger, and A. G. Beck-Sickinger Universit¨at Leipzig, Leipzig, Germany

Originally published in: Protein-Ligand Interactions. Edited by Hans-Joachim B¨ohm, Gisbert Schneider. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30521-6

1 Receptors

Receptors are the trigger molecules regulating a great variety of metabolic and physiological processes in cells as well as in more complex organisms. In the pharmacological sense, receptors are transducer proteins that selectively and reversibly bind an endogenous signaling molecule or its synthetic analogue, undergo a conformational change, and modify a cellular response as a consequence. This very general deﬁnition of the term receptor will be speciﬁed more precisely if we look at the different types of receptors that are present in mammalian cells. On one hand, there are the intracellular receptors of the nuclear receptor superfamily such as the steroid hormone receptors, the retinoid receptors, and the thyroid hormone receptors. The second category of receptor proteins is that of the membrane-bound receptors, which are by far more diverse. Those receptors serve the cells to transform a signal from the outside into a cellular answer. The superfamily comprises the ion-channel-coupled receptors, the kinase-coupled receptors, and the large family of G-protein-coupled receptors. In this chapter we focus on the characterization of the interaction between Gprotein-coupled receptors and their ligands. The presented techniques for the examination of ligand binding and receptor activation, however, are not limited to the analysis of this receptor group. 1.1 The G-Protein-Coupled Receptors

The G-protein-coupled receptors (GPCRs) are integral membrane proteins with the common characteristic of seven membrane-spanning helices. Their endogenous Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Analysis of Receptor-Ligand Interaction

ligands can be monoamine messengers (epinephrine, acetylcholine, serotonin, histamine, dopamine, etc.), lipids (prostaglandins, endogenous lipids, etc), neuropeptides (neuropeptide Y [NPY], substance P [SP], cholecystokinin [CCK], opioids, etc.), and peptide hormones (glucagon, angiotensin, bradykinin), as well as small proteins (chemokines) and large proteins (glycoprotein hormones, thrombin, etc.). All GPCRs transduce their signals to the interior of the cell through the interaction with heterotrimeric G-proteins. Additionally, important sensory proteins such as rhodopsin and the olfactory receptors belong to this family. The length of the different GPCRs varies considerably, from less than 300 amino acids for the smallest representatives, such as the adrenocorticotropin receptor, to more than 1100 amino acids for the metabotropic glutamate receptors. The receptors carry different posttranslational modiﬁcations such as glycosylation, which often leads to higher molecular weights of the receptors than expected from the amino acid sequence. Other modiﬁcations include disulﬁde bridges and palmitoylation at speciﬁc sites. Most frequently, the GPCRs are classiﬁed by primary sequence homology and subfamilies are named after well-characterized members. While only low homology is found in the loop segments, the seven transmembrane helices containing 20–25 hydrophobic amino acids show a higher degree of conservation [1]. So far our insight into the three-dimensional structure of GPCRs is rather limited, as it is very difﬁcult and time-consuming to crystallize such complex integral membrane proteins. Therefore, our knowledge about the 3-D structure of GPCRs is mainly based upon the crystal structures of bacteriorhodopsin and rhodopsin, the only homologous proteins with an elucidated crystal structure. Mainly, the crystal structure of bovine rhodopsin, solved by Palczewski et al. in 2000 [2], in combination with molecular modeling provides valuable knowledge for the understanding of other GPCRs. Computer models based on the sequence homology between rhodopsin and other receptor proteins can now be calculated. However, these models have to be investigated experimentally and crystal structures do not necessarily represent a receptor’s native state, which is to be considered as a dynamic equilibrium rather than a single solid state.

2 Ligand-binding Theory

Binding of a ligand can lead to agonistic, antagonistic, or inverse agonistic effects. Those effects are related to conformational changes in the receptor proteins and subsequent activation or deactivation of signal transduction cascades (Fig. 1). Due to a conformational change, receptors in the active state gain the ability to bind heterotrimeric G-proteins, a variety of which are displayed in Tab. 1. The G-proteins that are present in their inactive GDP-binding form then exchange GDP to GTP. Smaller subunits of the heterotrimeric G-protein complex dissociate. The GTPbinding subunit also dissociates from the receptor and activates a speciﬁc signal transduction cascade. This can mean activation or deactivation of the adenylate cyclase, ion channels, phospholipase C, or phosphodiesterase.

2 Ligand-binding Theory

Fig. 1 General scheme of the activation and signal transduction cascade by a GPCR. After binding of a ligand, the receptor is stabilized in the active conformation. It then can bind to the heterotrimeric G-protein. GDP bound by the "-subunit is exchanged

by GTP. The "-subunit dissociates from the $(-complex. The signal can be transduced by the activated "-subunit as well as by the $(-complex, which both then interact with their respective effector molecules.

So far, mainly two general models have been suggested that describe the interaction between a ligand and its receptor. The ﬁrst one, also known as the induced ﬁt or conformational induction hypothesis, postulates the receptor to be there in an inactive conformation. Agonist and antagonist are thought to bind to the receptor in a similar way. Binding of an agonist induces a conformational change in the receptor molecule, which then leads to the binding of G-proteins and activates the signal transduction cascade. In this model, antagonists bind to the same binding site as agonists, but do not induce the conformational change. The model can explain the action of agonistic or antagonistic ligands. However, some ligands have inverse agonistic effects, and, in fact, more and more substances formerly considered as antagonists are now classiﬁed as inverse agonists [3]. Their binding to a receptor not only inhibits the corresponding signal transduction pathway but even Table 1 G-protein subfamilies classiﬁed by their α-subunits

Subfamily

Effector protein

Gs

Adenylate cyclase ↑ Ca2+ channels↑ K+ channels↑ Ca2+ channels↓ cGMP speciﬁc phosphodiesterase↑ Adenylate cyclase↓ Phospholipase Cβ4 -

Gi

Gq G12

3

4 Analysis of Receptor-Ligand Interaction

reduces it below its basic level. For example, this is the case for some therapeutic drugs such as cimetidine (histamine H2 receptors), haloperidol (dopamine D2 receptors), prazosin (α 1 -adrenoreceptors), timolol (β 2 -adrenoreceptors), and clozapine (D2 receptors and 5-HT2c receptors) [4–8]. Furthermore, the phenomenon of constitutively active receptors has been observed where receptors are in an active state without ligand binding. This is, for instance, the case for the human β 2 adrenergic receptor [9] and the human calcitonin receptor [10]. GPCRs have been expressed at different levels in cell culture, and a direct relationship between the level of expression and basal signaling could be shown [11]. Constitutive receptor activity has been found mostly as a result of mutational changes in the sequence. Both inverse agonism and constitutive activity suggest that not all receptor-ligand interactions can be explained by the induced ﬁt hypothesis. The second model is that of conformational selection, which has been developed more recently. According to this hypothesis, GPCRs exist in at least two conformations. At least one of them binds to G-proteins and therefore is considered as the active state (R*). In other states the receptor cannot bind to G-proteins and is therefore referred to as the inactive, uncoupled receptor. There is an equilibrium between active and inactive conformations R* ↔ R. In the absence of a ligand, usually the inactive receptors represent the majority in this equilibrium. If a ligand is added, it may prefer a special conformation of the receptor for binding. In the case of an agonist, this will be the active state, whereas inverse agonists will prefer the inactive state. Binding of inverse agonists will therefore lower the level of basal signaling. Ligands that bind to active and inactive conformations of a receptor with the same afﬁnity will not inﬂuence the equilibrium. They are competitive antagonists, as they can displace other ligands from the binding site but do not lead to changes in basal signaling. Ligands with only little preference for the active state will lead only to a small shift in the equilibrium towards the active conformation. The consequence is an only moderate increase in signaling, also referred to as low efﬁcacy. Those compounds are partial agonists (Fig. 2). Between R and R* there are only small differences in energy. This can be seen by single amino acid mutations leading to constitutive receptor activity [12–14]. Conformational selection and conformational induction, however, are not necessarily in contradiction. Both can be seen as two parts of one mechanism in which the active conformation of a receptor is induced during the binding process, whereas the ligand preferably binds to the active conformation [15].

3 Characterization of the Receptor-Ligand Interaction

The characterization of new receptors usually starts with the discovery of a new receptor-ligand pair. This often starts with knowledge about a pharmacologically active compound, the mode of action of which is unknown, as was the case for neuropeptide Y [16], a 36-amino-acid neuropeptide extracted from porcine brain in 1982. On the other hand, it is also possible that a receptor is discovered for which no ligand is known (orphan receptor). This was the case for the orexin receptors

4 Receptor Material

Fig. 2 Model of the receptor-ligand interaction. Active (R*) and inactive (R) receptors are in a state of equilibrium. Whereas an agonist binds to the active conformation, an antagonist binds to the inactive conformation of the receptor. By stabilizing one

conformation, agonists and antagonists may shift the equilibrium to the respective side. Competitive antagonists will bind to both conformations with the same afﬁnity and therefore not inﬂuence the equilibrium.

OX1 and OX2 [17]. For neuropeptide Y (NPY) in humans, until now at least four functional receptors have been identiﬁed, the NPY-Y1 , NPY-Y2 , NPY-Y4 , and NPYY5 receptors. (For the NPY-Y4 receptor, the endogenous ligand is the pancreatic polypeptide.) In the case of the orexin receptors, we know two peptide ligands, orexin A and orexin B, that differ signiﬁcantly in their structure [18]. Once such a set of ligands and their corresponding receptors are known, the characterization of their interaction is necessary. On the one hand, it is desirable to understand how a ligand like NPY can distinguish between its receptors and whether there are differences in the binding mode. This will be especially important if it comes to the design of subtype-speciﬁc, pharmacologically active compounds. On the other hand, in cases like the orexin receptors — where we have the two ligands orexin A and B both binding to the OX1 and the OX2 receptor, respectively, but with different afﬁnities [17] — it is important to ﬁnd the causal connection between the ligand structure and its receptor selectivity.

4 Receptor Material

Usually, G-protein-coupled receptors and their ligands are ﬁrst identiﬁed from primary tissue. However, it is difﬁcult and also ethically questionable to get large enough amounts of primary tissue samples to closely investigate them. Furthermore, the stability of those samples and the accessibility of the embedded receptors can be a problem. There are different ways to circumvent those difﬁculties. Probably the easiest way is the isolation and cultivation of cancer cell lines that endogenously express the receptors that are of interest. This has been done with the SK-N-MC [19] cell line and the SMS-KAN cell line [20], which express the NPY-Y1 and the NPY-Y2 receptor, respectively. Another commonly used method is the cloning of

5

6 Analysis of Receptor-Ligand Interaction

the receptors into either cancer cell lines or bacterial or yeast cells. Cancer cells have the advantage that they come closest to mammalian tissue cells, are able to make posttranslational modiﬁcations, and do not have a cell wall like yeast cells do. A number of such cell lines have been established, such as BHK (baby hamster kidney), CHO (Chinese hamster ovary), HEK (human embryonic kidney), or COS (SV40 transformed African green monkey kidney cells) cells. Those cell lines can be transfected with the DNA of GPCRs of interest [21–24]. The advantage is that this method allows studies with genetically modiﬁed receptors in which single or different amino acids are mutated or in which a receptor is fused to another protein [25]. The expression of fusion proteins is often used in combination with ﬂuorescent proteins, such as green ﬂuorescent protein (GFP) [26], to make receptors visible and to perform ﬂuorometric assays (see Section 8). Usually, such transfections are transient. To make them stable, one can combine the receptor encoding DNA sequence with a sequence encoding for an antibiotic resistance. Cultivation of the respective cell line in an antibiotic-containing medium over a longer period of time then can lead to a stable transfection [27–29]. 5 Binding Studies

For new receptor-ligand pairs, usually the determination of binding parameters is the ﬁrst step. Also, for new, artiﬁcial compounds designed as new ligands for a receptor, it is important to determine whether they are binders. Therefore, a variety of binding assays have been established that can be divided into two groups: separation assays and direct assays [30]. In separation assays either whole cells or membrane preparations containing the receptor of interest are used. They are incubated with a radioactive- or ﬂuorescent-labeled ligand until the state of equilibrium is reached. The receptor-ligand complex is then separated by centrifugation or ﬁltration and the amount of bound ligand is determined. The more recently developed direct assays, based on surface plasmon resonance (SPR) [31] or ﬂuorescence correlation spectroscopy (FCS) [32], measure the receptor-ligand interaction in real time. 6 Binding Kinetics

Interactions between G-protein-coupled receptors and their ligands are reversible with the exception of rhodopsin. The binding parameters of a ligand can therefore be determined in a competition-binding assay with a labeled ligand. In such an assay, the displacement of labeled ligand from the receptor is measured in the presence of different concentrations of the examined ligand. The general equation for a bimolecular association between a receptor (R) and a labeled ligand (L*) is k12

R + L ∗ RL ∗ k21

(1)

6 Binding Kinetics 7

where k12 is the association rate (on-rate) and k21 is the dissociation rate (off-rate) of the receptor-ligand complex. The association constant K (not to be confused with the association rate k12 ) of the ligand binding reaction is K=k12 /k21 , whereas the dissociation constant K d is K d =1/K = k21 /k12 . When L*=K d =1/K, then RL* = Rt /2, with Rt being the total receptor concentration. In other words, if the free concentration of the labeled ligand L* reaches the value of K d , the receptor-binding sites will be half-saturated with ligand. The value of K d is one-half of the maximal speciﬁc binding Bmax . The free ligand concentration at 50% receptor saturation, the IC50 concentration, is a measure of K d (or 1/K). The K d and Bmax values of a ligand are determined in a saturation analysis. Cell membranes or whole cells are incubated with different concentrations of a labeled ligand. The resulting curve consists of speciﬁc binding to the receptorbinding sites and non-speciﬁc, non-receptor-binding sites. Each concentration of the labeled ligand should therefore be displaced by a 1000-fold excess of unlabeled ligand to distinguish between speciﬁc and non-speciﬁc binding. To get the values for the speciﬁc binding, the values for non-speciﬁc binding are subtracted from those for total binding. The resulting curve for speciﬁc binding should be saturable at sufﬁciently high concentrations and represent a hyperbolic function (Fig. 3a). As the K d value is equal to the concentration of labeled ligand occupying 50% of the Bmax value, it can be determined from the saturation curve. The data was formerly analyzed by Scatchard analysis where it is summarized in the Scatchard plot. The amount of bound ligand divided by the amount of free ligand in solution (y-axis) is plotted against the amount of bound ligand (x-axis) (Fig. 3b). In the case of a bimolecular interaction, this should lead to a straight line with a negative slope. The intercept point with the x-axis represents the Bmax value, whereas the absolute value of the slope represents the K d value (Eq. (2)). More recently, the Scatchard analysis is more and more replaced by computational non-linear regression analysis, a method that makes it possible to directly solve the equation of one-site ligand binding. y=

Bmax · x Kd + x

(2)

From the B max value, the number of binding sites per milligram membrane preparation or per cell, which corresponds to the number of receptors per cell, can be calculated. The IC50 concentration of an unlabeled ligand is determined in a competition assay with a labeled ligand L*. This has the advantage that not every new or uncharacterized ligand has to be labeled but can be tested against an already labeled and characterized compound. A constant concentration of L*, usually at or below its K d , is displaced by increasing concentrations of the unlabeled ligand. For a bimolecular reaction this results in a sigmoidal competition curve. The IC50 value is the concentration of unlabeled ligand, which displaces 50% of the speciﬁcally bound labeled ligand from the receptor-binding site (Fig. 4). The IC50 value, however,

8 Analysis of Receptor-Ligand Interaction

Fig. 3 (A) Saturation analysis for a radiolabeled ligand. The x-axis represents the concentration of the labeled ligand. The y-axis shows the dpm-values (decays per minute) for total binding, non-speciﬁc binding, and speciﬁc binding. The values for speciﬁc binding are obtained by subtraction of non-speciﬁc binding and total binding. (B) Scatchard plot of a saturation analysis

experiment. The x-axis shows the amount of bound ligand, whereas the y-axis represents the amount of bound ligand divided by the amount of free ligand in solution. Bmax is determined from the intercept point with the x-axis. K d is the absolute value of the slope (both graphs present artiﬁcial, idealized data).

is dependent on the concentration of labeled ligand and may vary between experiments performed under different conditions. With the constant value K i , which is independent from the concentration of the labeled ligand, it is easier to compare datasets from different experiments. K i is calculated according to the equation of Cheng and Prusoff [33]:

Ki =

IC50 1 + KLd

(3)

7 Binding Assays 9

Fig. 4 Competition analysis (idealized data). The labeled ligand, which is held at a constant concentration at or below its K d , is displaced by increasing concentrations of the examined ligand. The x value of the curve’s inﬂection point represents log IC50 .

7 Binding Assays 7.1 Separation Assays

In separation assays the receptors, usually in the form of membrane preparations or whole cells, are incubated with a labeled ligand. The incubation should be long enough to reach the state of equilibrium, which can be anything from seconds to hours. For thermodynamic reasons, the incubation should take place at room temperature or body temperature, which allows the receptors as well as the ligands to change conformation and therefore to interact with each other. As soon as the state of equilibrium is reached, the receptor-bound fraction of the labeled ligand is separated physically from the unbound fraction, which is still in solution. This can be done by either centrifugation or ﬁltration. To avoid disturbance of the established equilibrium during the separation process and following washing steps, the sample should be cooled during these procedures. This signiﬁcantly reduces the association and dissociation rate. Then the amount of receptor-bound labeled ligand can be determined [30]. 7.2 Radioligand-binding Assay

One of the most common separation assays to test for a ligand’s binding to a receptor is the radioligand-binding assay. Radioactively labeled compounds are used in this type of assay. The high sensitivity of the method allows for the detection and quantiﬁcation of very small amounts of ligand. Tritium (3 H) and iodine-125 (125 I) are the most commonly used isotopes. The selection of the respective label should, besides the availability, consider the radiochemical and safety properties. Tritium has a long half-life period (more than 12 years), which means that no correction is needed for the decay during the experiment and a labeled tracer can be used over a long period of time (provided that it is chemically stable). Furthermore, its incorporation

10 Analysis of Receptor-Ligand Interaction

into a molecule has no or only few sterical effects. Its properties as a beta radiator with a low radiant energy and only short radiant distance (0.6 cm in air) make it more convenient for handling than the gamma emitter iodine-125. However, the lower efﬁciency in measurement of 3 H (only 40% in liquid scintillation counting) may make it unsuitable for some applications. 125 I, with its higher radiant energy, higher maximum speciﬁc activity, and higher efﬁciency of measurement (75–90% in gamma counting), may be more suitable for assays in which very high sensitivity is required. One major disadvantage is, however, the size of the isotope 125 I, which can cause sterical changes in the tracer molecule and thereby inﬂuence its binding properties. Also, the low half-life period of 125 I makes it necessary to re-determine the speciﬁc activity of the labeled ligand before every assay. The radioactive binding assay can be used in both saturation and competition assays (see Section 6). In the saturation analysis, a variety of ligand concentrations are used to get a saturation curve. Each concentration is then competed by a 1000fold excess of the unlabeled ligand to determine the level of non-speciﬁc binding. In the competition assay, a single concentration of the radioligand is used. This concentration should preferably be at or below the K d value of the respective ligand. It is then competed by a variety of concentrations of the unlabeled ligand. The concentrations of the unlabeled competitor should be chosen in a way that the resulting competition curve covers the complete range from no displacement at all to complete displacement of the labeled ligand from the receptor [30, 34].

8 Fluorometric Assays 8.1 Fluorescence Labels

Examples of ﬂuorescence labels for ligands are carboxyﬂuorescein, Cy3TM , a commercially available ﬂuorescent marker based on a cyanine dye or tetramethylrhodamine. They are chemically introduced into a ligand. As with the radioactive labels, a possible inﬂuence of the labels on the binding behavior of the labeled ligands has to be considered, especially as the ﬂuorescent dyes are complex molecules. Furthermore, the receptors themselves can be ﬂuorescent labeled, which is done recombinantly. The respective receptors are expressed as fusion proteins with ﬂuorescent proteins, e.g., green ﬂuorescent protein (GFP) from Aequorea victoria, one of its mutant variants, or DSRed from Discosoma striata [26, 35]. 8.2 Fluorescence Correlation Spectroscopy (FCS)

For a number of assays, ﬂuorescent-labeled analogues of a ligand are used. Some of those assays are of a quantitative nature, such as in ﬂuorescence correlation spectroscopy (FCS), where the ﬂuorescent-labeled analogue is used to determine

8 Fluorometric Assays 11

Fig. 5 Schematic setup of a ﬂuorescence correlation spectrometer. A beam of parallel laser light is focused by an immersion objective onto a ﬂuorescent sample. The ﬂuorescence of molecules traversing the focus is collected by the same objective. It is ﬁltered, focused, and detected. Signal autocorrelation is carried out by the computer (PC).

binding kinetics (Fig. 5) [36]. FCS allows the direct detection of molecular interactions in solution. FCS monitors the random motion of a ﬂuorescent molecule in a deﬁned volume (1 fL). Thereby, the diffusion rate of a particle can be detected, which is directly dependent on the particle’s mass. Therefore, FCS can quantify the bound and the free fraction of the ﬂuorescent-labeled ligand and can be used to determine binding parameters. In contrast to the radioligand-binding assay, this has the advantage that the receptor-ligand interaction can be directly monitored over a longer period of time. Recently, FCS was applied to study the receptor diversity of the neuropeptide galanin in cultured cells [37]. In this case three different diffusion times for rhodamine-labeled galanin (Rh-GAL) could be detected: one short diffusion time for unbound Rh-GAL and two different longer diffusion times for membrane-bound Rh-GAL. Those ﬁndings suggest that FCS allows one not only to determine the amounts of bound and unbound ligand for a receptor-ligand complex but also to distinguish between different receptor subpopulations or different receptor subtypes. 8.3 Fluorescence Microscopy

Other ﬂuorometric assays will be of a more qualitative nature, if it comes to microscopic studies to characterize receptor expression on cells [38, 39] (Fig. 6) or receptor internalization upon the binding of a ligand [40]. 8.4 Fluorescence Resonance Energy Transfer (FRET)

The use of either pairs of differently ﬂuorescent-labeled receptors or ﬂuorescentlabeled receptors and ﬂuorescent-labeled peptides allows ﬂuorescence resonance

12 Analysis of Receptor-Ligand Interaction

Fig. 6 Fluorescence microscopy image of BHK cells expressing a human NPY-Y2 receptor-GFP fusion protein.

Fig. 7. Principle of FRET. A donor ﬂuorophore is excited at a certain wavelength and transfers its radiation energy non-radiatively to a closely located acceptor ﬂuorophore. Emission at the acceptor’s wavelength can therefore be measured to determine whether the two chromophores are co-localized or not.

energy transfer (FRET) studies. The prerequisite is that the used chromophores form a FRET pair. This means, when in close proximity, that the so-called donor, excited at a certain wavelength, transfers its radiation energy non-radiatively to the closely located acceptor chromophore. Emission at the acceptor’s emission wavelength can be detected (Fig. 7). A receptor-ligand FRET pair can be used to study the binding of a ligand to its receptor [41]. The co-transfection of cells with a receptor-receptor FRET pair can help one to decide whether a receptor is present in the cell membrane as a monomer or dimer [42, 43]. If two different types of receptors are used in a FRET assay, it may even help one to decide whether the receptors are present as heterodimers.

9 Surface Plasmon Resonance

Another technique used for the analysis of receptor-ligand interaction is surface plasmon resonance (SPR), with its ﬁrst commercially available application in the R BIAcore instruments [44] (Fig. 8). Like FCS, it allows the determination of kinetics by monitoring the association and dissociation of a receptor-ligand complex in real

9 Surface Plasmon Resonance 13

Fig. 8 Schematic setup of the surface plasmon resonance (SPR) detection unit in a R Bia-core instrument. One interaction partner is immobilized on a modiﬁed gold surface, whereas the other ﬂows by in solution. On the other side of the sensor chip, a beam of polarized light is reﬂected by the gold ﬁlm. The optical phenomenon SPR leads to a reduction in the intensity of re-

ﬂected light at a certain angle (A). This effect is dependent on the refractive index at the sensor surface and thereby on the mass bound to the chip. Therefore, the angle changes if a ligand is bound by the molecules immobilized on the chip (B). The response values are then displayed in resonance units (RU).

time. The interaction partners do not necessarily have to be labeled, which is an advantage of the technique. The principle of SPR measurements is based on an optical phenomenon. The core unit in this technique is a sensor chip consisting of a thin gold ﬁlm with a modiﬁed surface attached on one side. One reactant is attached to the modiﬁed sensor surface, whereas the other reaction partner ﬂows past this surface in solution. When the two interaction partners form a complex, the mass on the sensor chip surface increases; when the complex dissociates, the mass falls. This leads to changes in the refractive index in the aqueous layer close to the sensor chip surface that is measured by an optical detection unit on the dry side of the chip. The signal measured in arbitrary resonance units (RU) is approximately proportional to the change in mass with 1 RU=1 pg/mm2 for proteins [45]. A variety of sensor chips with modiﬁed surfaces is commercially available, most of them based on a carboxymethyldextrane matrix. This can be used to directly couple one interaction partner via its functional groups by deﬁned coupling chemistries for thiol, amine, or aldehyde coupling, or it can be further modiﬁed for capturing biotinylated or histidine-tagged interaction partners or for capturing liposomes. Finally, there is also a hydrophobic sensor surface available, composed of long-chain alkanethiol molecules, that can be used to construct lipid bilayers as membrane-like environments [46]. As recently described, liposome-capturing sensor chips can also be used to reconstitute G-protein-coupled receptors on them [47]. This was done with rhodopsin as the most frequently used model for G-protein-coupled receptors in general. Rhodopsin was immobilized and reconstituted in mixed micelles on the sensor chip surface. If the technique turned out to be suitable for other Gprotein-coupled receptors as well, it might become a valuable tool to test for the functionality of solubilized receptors, as well as to screen for new ligands and to determine binding kinetics.

14 Analysis of Receptor-Ligand Interaction

10 Molecular Characterization of the Receptor-Ligand Interaction 10.1 Antibodies

Antibodies can be used for a variety of applications in the molecular characterization of receptors and receptor-ligand interactions. Antibodies can be used for the detection of receptors in tissue slices, Western blot experiments [48], or ELISAs (enzyme-linked immunosorbent assays) [49]. They can also be used in competition experiments to map the binding epitope of a ligand [50]. Even though the use of antibodies is routine, there is no general protocol for their generation. Polyclonal antibody sera contain many different antibodies with different afﬁnities for the epitopes of an antigen. They are generated by immunization of animals, mostly chickens or rabbits [51], with the antigen (Fig. 9b). To get good immunization results, the animal species should not be too closely related to the species where the antigen comes from. In the case of a G-protein-coupled receptor, immunization may be done with the whole protein or peptide segments of the receptor. Immunization with a whole receptor protein will create a serum with a high diversity of antibodies directed against all parts of the molecule — extracellular, intracellular, or transmembrane segments [51]. The production of sufﬁcient amounts of puriﬁed receptor is often difﬁcult and is connected with considerable loss of material. One possibility is the separation on a polyacrylamide gel, followed by blotting onto a nitrocellulose membrane. The membrane can then be introduced under the skin of the animal. Even more drastic is the direct injection of a portion of polyacrylamide gel containing the receptor. However, these methods not only are very stressful for the animal but also are applicable only if other methods to identify the respective receptors are known. As antibodies are often generated against not very well characterized proteins, this may be a problem. The immunization of animals with synthetic peptides is a way to circumvent these problems. Furthermore, it has the advantage that it leads to antibodies directed against a speciﬁc and known part of the protein, a beneﬁt in some applications. Such antibodies can recognize parts of a receptor involved in ligand binding or serve as ligands with agonistic [52, 53] or antagonistic properties themselves. They can also displace other ligands, which might help to localize their binding site at the receptor. The peptide sequences are available from GenBank entries of the respective receptor or can be determined from the coding gene sequence. To allow good accessibility in cell or membrane assays, the peptide segments should preferably be chosen from parts of the protein exposed to the surface, especially the extracellular domain [49]. For previously unknown proteins, this information can be estimated from hydropathicity blots. Antibodies generated against intracellular parts of the receptor can be used to explore signal transduction processes or Gprotein binding [54]. To act as an antigen, the mass of a protein should be higher than 5–10 kDa. This often is not the case for the peptides of a receptor loop, so they have to be coupled to

10 Molecular Characterization of the Receptor-Ligand Interaction 15

Fig. 9 (A) Model of an immunoglobulin molecule. The molecule consists of the Fc portion, containing two disulﬁde bridges and two Fab portions with one disulﬁde bridge each. The Fab portions carry the variable domains of the antibody. (B) Immunization scheme for rabbits. Before administration of the antigen, a preliminary bleed should be taken. The animal is then immunized with the antigen. After 14 and 28 days, the ﬁrst and second booster immunizations are given. After 35 days a ﬁrst

bleed is taken from the animal. The third booster immunization follows after 56 days. Final bleeding is taken after 63 days. (C) Phage display for the generation of monoclonal antibodies. Different antibodies are displayed on the phages’ surfaces. They are then selected via an antigen. The phages that display antibodies with afﬁnity for the antigen are ampliﬁed in E. coli. Finally, the sequences of the respective antibodies are identiﬁed on the DNA level [51].

16 Analysis of Receptor-Ligand Interaction

a larger protein carrier. Because the immunogenic reaction will be directed against parts of the carrier as well, it should not be relevant for future assays. Bovine serum albumin (BSA) or keyhole limpet hemocyanin (KLH) is commonly used. Because BSA is often used as a blocking agent, for instance in Western blotting, KLH might be a better choice in many cases. Antibody sera obtained from immunized animals can in some cases be directly used for further experiments. Further puriﬁcation and enrichment, however, are often necessary to increase the speciﬁcity and overall afﬁnity of the puriﬁed antibodies. Simple enrichment of the antibodies can be achieved by precipitation with ammonium sulfate [55] or by chromatography with protein A [56] or protein G, bacterial cell wall proteins that speciﬁcally bind to the Fc portion of immunoglobulins. A more speciﬁc enrichment is achieved by afﬁnity chromatography with an antigen column. An alternative to the rather complex mixtures of polyclonal antibodies are monoclonal antibodies [51]. In the ﬁrst step, as for polyclonal antibodies, an animal is immunized. The antibody-secreting lymphocytes are then isolated from lymphoid tissue and fused with plasmacytoma cells of a similar differentiation state. The new hybridoma cells can then be cultivated and selected, and supernatants can be screened for activity against the antigen. This procedure can be repeated until a clone is found, which produces antibodies with good afﬁnity. Monoclonal antibodies have the advantage that they can theoretically be reproduced inﬁnitely. Another possibility for the generation of monoclonal antibodies is phage display [51] (Fig. 9c), an evolutionary technique in which antibody V genes are cloned for display of assembled heavy- and light-chain variable domains into ﬁlamentous bacteriophage. Phages binding to the antigen are selected and soluble antibody fragments are expressed by infected bacteria. 10.2 Applications of Antibodies 10.2.1 Receptor and Ligand Detection Because there are a huge variety of applications for antibodies, in this section we focus on those concerning receptor characterization and receptor-ligand interaction. Antibodies represent a valuable tool to detect receptors as well as their ligands in tissue samples by direct staining of tissue slices. Another method is the separation of cell membranes by SDS polyacrylamide gel electrophoresis (SDS-PAGE), followed by Western blotting onto a nitrocellulose or PVDF membrane (Fig. 10). In both cases the receptors are incubated with the antibodies, non-bound antibodies are washed away, and a second antibody speciﬁc for the Fc fragment of the ﬁrst antibody is added. The second antibody is labeled, either for direct detection with a ﬂuorescent dye or radioactivity or for detection by a staining reaction, with enzymes like alkaline phosphatase or horseradish peroxidase. Often nothing is known about new receptors but their amino acid sequence. In such cases antibodies can be used to screen for their production in various tissues either directly in tissue slices or after immunoblotting of membrane preparations

10 Molecular Characterization of the Receptor-Ligand Interaction 17

Fig. 10 Receptor characterization by SDSPAGE and Western blotting. A cell membrane sample is separated by SDS polyacrylamide gel electrophoresis. The proteins are then blotted onto a nitrocellulose membrane. The membrane is incubated with an antibody that is speciﬁc for the receptor. Af-

ter incubation with a second enzyme-linked antibody, the receptor band is stained with an enzymatic dye reaction. The parallel separation of a standard protein mixture allows one to determine the molecular mass of the receptor (MP: marker proteins; CM: cellular membrane) [51].

[48, 49]. Such immunohistochemical studies were performed for the angiotensin II receptor subtypes AT1 and AT2 in rat adrenal gland [48] and in the heart of rabbits [57], for the metabotropic glutamate receptor in rat piriform cortex [58], for serotonin 5-HT2A and 5-HT3 receptors in inhibitory circuits of the primate cerebral cortex [59], and for the dopamine D1 and D2 as well as M4 muscarinic receptor in striatonigral neurons [60]. 10.2.2 Receptor Characterization The detected receptor masses in immunoblotting experiments often differ significantly from those calculated from the amino acid sequence [48, 49]. Sometimes even several bands of different molecular masses are detected for one receptor. Higher receptor masses and different bands may be due to posttranslational modiﬁcations like glycosylation or lipid residues. However, such observations are often

18 Analysis of Receptor-Ligand Interaction

difﬁcult to explain and have to be checked carefully in each case. To distinguish whether a receptor double band is due to differences in the degree of glycosylation, due to different splice variants, or simply due to receptor degradation during the work-up process, a deglycosylation experiment can be performed. For this the receptors are treated with enzymes that release oligosaccharides from glycosylation sites, followed by immunoblotting and staining with antibodies. If a receptor is glycosylated, this should result in a single band, which should be detectable at the calculated molecular weight of the receptor. The most frequent posttranslational modiﬁcations for GPCRs are N-glycosylation at the N-terminus and external loop Asn-X-Ser/Thr sequences (human calcitonin receptor-like receptor [61] and α 1B -adrenergic receptor [62]), palmitoylation (human dopamine D1 receptor [63]), and phosphorylation (β 2 adrenergic receptor [64]). For many receptors different subtypes with a high degree of sequence homology are known. In cases such as the neuropeptide Y receptor family – with the Y1 , Y2 , Y4 , and Y5 receptors functionally expressed in humans — or the angiotensin II receptor family — with the receptors AT1A , AT1B , and AT2 in human — antibodies that distinguish between different receptor subtypes are a valuable tool to monitor the receptor expression on the protein level [48, 49]. 10.2.3 Functional Characterization of the Receptor-Ligand Interaction Antibodies generated against a peptide segment of a receptor can be used to map the binding site of the endogenous ligand at the respective receptor. This is especially the case if a variety of antibodies generated against different receptor peptides are available. The receptors are then simultaneously incubated with ligand and antibodies. In an ELISA the amount of antibody bound in the presence or absence of the ligand can be determined. Competition is expected for antibodies that have binding sites at the receptor that are overlapping with the ligand’s binding site. In addition, competition assays with radiolabeled ligand can be performed. In that case the amount of ligand bound to the receptor is determined after simultaneous incubation with the antibody. The assay is performed in the same way as the radioactive competition assay (see Section 7.2). Finally, if photoactivatable analogues (see Section 10.5) of the ligand are available, they can be cross-linked to the receptor. Afterwards, the cross-linked receptor-ligand complex can be incubated with the anti-receptor antibodies to see whether their binding site is blocked. A combination of these techniques was used to characterize the binding site of NPY at the Y 1 receptor [50]. 10.3 Aptamers

Aptamers (Latin aptus=ﬁt) are RNA or DNA molecules isolated from combinatorial nucleic acid libraries by in vitro selection experiments termed SELEX: systematic evolution of ligands by exponential enrichment. A few binders with good afﬁnities for the target are selected from up to 1015 different oligonucleotide sequences. The selection is performed by column chromatography or other enrichment techniques

10 Molecular Characterization of the Receptor-Ligand Interaction 19

that are suitable to separate binders from non-binders. Non-binders are washed away, whereas the binders are regained, ampliﬁed, and subjected to a new cycle of selection. After several cycles the stringency of the binding conditions is increased, which allows the selection of good binders only [65]. Developed in 1990 [66, 67], until now aptamers have been isolated against more than 150 target molecules, among them small organic molecules, amino acids, peptides, and proteins [68]. The mechanism of target recognition by aptamers is adaptive [69]. Whereas they are predominantly unstructured in solution, aptamers fold upon association with their ligands. The ligand becomes an intrinsic part of the nucleic acid structure, and the 3-D structures of aptamer complexes form highly optimized scaffolds for speciﬁc ligand recognition. Thus, aptamers, like receptors, seem to be able to distinguish not only between different ligand molecules but also between different conformations of a single molecule. This was nicely demonstrated for an aptamer selected against the 36-amino-acid peptide NPY [70]. NPY is the endogenous ligand for the Y1 , Y2 , and Y5 receptors. Receptor-selective, conformationally constrained, synthetic analogues of the peptide are known, suggesting that NPY binds to its receptors in different conformations (see Section 10.4.2). The generated aptamer was tested against a variety of these peptides and showed preference for Y2 -receptorselective analogues of NPY. Furthermore, a competition experiment was performed in which aptamer and radiolabeled NPY were simultaneously incubated with the respective receptors. Again, the aptamer showed higher competition for NPY at the Y2 receptor. Recently, intramers, aptamers that can be expressed inside cells and retain their function [71], were developed. Such intramers might also be generated against biomolecules that are part of a signal transduction pathway. In the future they might be used to elucidate signal transduction cascades triggered by a receptor. 10.4 Receptor Mutation and Ligand Modiﬁcation

A way to gain insight into the interaction between a receptor and its ligand is to mutate single or several positions in the receptor sequence or to modify the ligand. Whereas in the receptor molecule an amino acid is usually substituted by another, the modiﬁcations of the ligand can be various. Different ligands, endogenous or synthetic, for a receptor do not necessarily have the same chemical structure. They do not even have to be members of the same chemical class; therefore, the determination of a pharmacophore (deﬁned orientation of functional groups being the basis of biological action) may be difﬁcult. Another problem is the determination of the ligand’s binding site at the receptor, especially because membrane receptors are difﬁcult to crystallize and our structural models of G-protein-coupled receptors are based mainly on the X-ray structure of rhodopsin. 10.4.1 Receptor Mutagenesis One way to locate domains of a receptor that are involved in ligand binding is the creation of chimeric receptors. Chimeric receptors are fusion proteins in which

20 Analysis of Receptor-Ligand Interaction

one part originates from one receptor and the other part originates from another receptor. In most cases, chimeric receptors are used to obtain an initial picture of the location of interesting segments involved in ligand binding [72]. Usually, such chimeric constructs are made of two closely related receptors, such as the neurokinin NK1 and NK2 receptor [73], to obtain functional receptors. There are, however, examples of chimeric constructs from two distantly related receptors, such as the muscarinic M3 and the adrenergic α 2 receptor [74, 75]. In the respective chimeric constructs, the amino terminal ﬁve transmembrane domains (TMs) originated from the α 2 adrenergic receptor and the carboxy terminal two TMs originated from the M3 receptor (or vice versa). Whereas the single chimeric constructs were not functional, the co-expression of both constructs leads to functional activity corresponding to both an adrenergic and a muscarinic receptor. Another way to gain more detailed information on important residues for ligand recognition is site-directed mutagenesis of single amino acid residues of a receptor sequence. Before receptor mutations are introduced, one has to select amino acids that are likely to be of relevance for receptor function and ligand binding. One approach is to search for conserved residues of a receptor. This can be done by alignment either of sequences of the same receptor from different species or of sequences of different subtypes of a receptor family. Conserved amino acids, especially in the extracellular regions but also in the transmembrane regions, are likely to play a role in the mechanism of ligand binding. Amino acids in a transmembrane helix often interact with amino acids from another helix in a way that stabilizes the inactive conformation of a receptor [76]. Disruption of such an interaction by an agonistic ligand or by mutation of one interaction partner leads to a change in conformation and to activation of the receptor. This was demonstrated for the α 1B -adrenergic receptor in which residues Asp125 in transmembrane domain 3 and Lys331 in transmembrane domain 7 apparently form a salt bridge that holds the receptor in the inactive conformation [77]. The mutation of Lys331 to Ala led to a six-fold increase in recognition of the endogenous ligand epinephrine without inﬂuencing the binding behavior of selective antagonists. Furthermore, the mutation led to an increased level of basal receptor signaling. This explains why some receptors upon mutation of one single amino acid residue are constitutively active. In combination with modeling studies, an assumption for the second interaction partner can be made, which then has to be conﬁrmed by creating the respective mutant. Conserved residues in the intracellular parts of a receptor are more likely to be involved in G-protein binding and signal transduction. This is the case for the DRY motif, a highly conserved amino acid sequence that, with the exception of some conservative mutations, is present in all rhodopsin-like receptors [1]. Another way to identify residues that might be important for ligand binding is to take amino acids from the putative binding region of a receptor and to search for possible interaction partners in the ligand. This approach was performed with the human NPY-Y1 receptor, where a number of negatively charged aspartic acid and glutamic acid residues in the extracellular domains of the NPY-Y1 receptor were systematically mutated to alanines [78]. Those residues are possible partners for electrostatic interactions with positively charged amino acids (arginine, lysine,

10 Molecular Characterization of the Receptor-Ligand Interaction 21

and histidine) in the N- and C-termini of NPY. In radioligand-binding assays, a number of residues were shown to be essential for ligand binding (D104, D194, D200, D287), whereas others had no or only moderate effects. 10.4.2 Ligand Modiﬁcation To investigate which residues of the ligand are important for receptor recognition, analogues of a known ligand often are synthesized and tested for binding at the respective receptors. This can be done by rational or combinatorial approaches or by a mixture of both. In combinatorial methods the ligand is modiﬁed systematically, and up to several hundred slightly different compounds are prepared. Such methods have become possible with the introduction of robotic techniques for chemical synthesis that allow the simultaneous synthesis of many compounds at the same time. Such a set of chemical entities is also called a library. The single compounds are then tested for binding and for their biological activities. Thus, more important residues can be distinguished from less important ones. If analogues with antagonistic properties are detected, structural comparison to the endogenous ligand may help to detect elements that are necessary for receptor activation. For instance, aromatic side chain residues like Tyr, Trp, His, and Phe can be found to play an important role in receptor activation [79]. Also, the reduction of backbone amide bonds converts some peptide agonists to antagonists. This was demonstrated for the C-terminal tetrapeptide of gastrin, where the respective pseudo-peptide analogues had antagonistic properties [80]. Other approaches are more rational. For peptide ligands, a fast way to locate the binding site on the ligand is to create truncated analogues, representing only parts of the original molecule. The truncation does not necessarily have to be N- or C-terminal; also, centrally truncated analogues have been reported to act as ligands. In the case of NPY, a highly Y2 -receptor-selective analogue was reported in which the central amino acids 5–24 have been replaced by a single aminohexa-noic acid (Ahx) molecule [81]. This leads to the assumption that a conformation in which the N- and the C-termini of the peptide are closely associated is relevant for binding at this receptor. As with the receptors, for peptide ligands, chimera can be created in which one part of the molecule originates from one peptide, whereas the other part originates from a second peptide. If the original peptides are ligands of the same receptor family, the chimera can be used to determine structural characteristics that are the precondition for subtype speciﬁcity. Chimeric peptides have been used to study the interaction between galanin and its receptors [82]. A more rational approach is the alanine scan (Fig. 11), a method frequently used to screen the amino acid sequence of peptide ligands for the contribution of each residue to the receptor-ligand interaction. In an alanine scan, every single amino acid of a peptide’s natural sequence is replaced by L-alanine. Alanine residues in the natural sequence are usually replaced by glycine. For example, a complete alanine scan of neuropeptide Y revealed residues that are important for NPY’s interaction with the NPY-Y1 and NPY-Y2 receptors [83]. Testing of the analogues at the respective receptors revealed that parts of the ligand, such as the C-terminal pentapeptide amide and especially R33, were necessary for the recognition of both

22 Analysis of Receptor-Ligand Interaction

Fig. 11 l-Alanine scan of CGRP Y0 -28–37, a selective antagonist for the human calcitonin gene-related peptide 1 receptor (CGRP1 ). The analogues were tested in binding-competition assays against (125 Iiodohistidyl)-CGRP. Whereas the replace-

ment of some residues leads to complete loss of binding (P29, T30, V32, and G33), other replacements have a more moderate effect on the receptor recognition (n.d.=no displacement; (A36) was not replaced) [100].

receptors. Other residues were needed only for the binding to one receptor (P2, the NPY loop and R33 for the Y1 receptor and the C-terminal helix, and Y36 for the Y2 receptor), whereas their exchange was tolerated by the other receptor. Based on the ﬁndings from an alanine scan, further peptides can be synthesized in which residues involved in the receptor recognition are substituted by homologous amino acids. Interesting in this context is also the use of conformationally constrained analogues in which amino acid residues such as the helix breaker proline or the turn-inducing motive alanine-aminoisobutyric acid are introduced into the natural peptide sequence. In the case of NPY, a ligand that has a distinct conformation in solution [84, 85], it is questionable whether the observed effects are due to direct interactions between residues of the ligand and side chains of the receptor or whether they are caused by an altered conformation of the ligand. Thus, instead of solely determining the binding afﬁnities of modiﬁed ligands, one has to investigate their conformation as well. Usually, it is easier by far to gain structural information on the ligands than on G-protein-coupled receptors. Many ligands can be synthesized in sufﬁcient quantity and purity, most of them are water-soluble, and some of them can also be crystallized. A number of techniques allow structural insight into ligand conformation, such as X-ray crystallography, solid-state and solution NMR, and circular dichroism studies for peptides. Well-characterized sets of ligands, especially when containing agonists as well as antagonists for a speciﬁc receptor, can be used for computer modeling and structure afﬁnity relationship studies. Ligands for receptors that are not members of the same chemical class and therefore do not share structural similarities at ﬁrst sight are especially helpful in creating a pharmacophore hypothesis.

10 Molecular Characterization of the Receptor-Ligand Interaction 23

10.4.3 Combination of Receptor Mutation and Ligand Modiﬁcation Some amino acids of a receptor’s sequence, when mutated, may have effects on the recognition of an agonist but not on the recognition of an antagonist and vice versa, a fact that supports the conformational selection hypothesis for the interaction between ligands and their receptors (see Section 2). Based on the mentioned mutagenesis study at the NPY-Y1 receptor and the results of the alanine scan of NPY, a model for the interaction of NPY with the hY1 receptor was designed [86]. In the meantime, the non-peptide compound BIBP 3226 was created and shown to act as a competitive, speciﬁc, and selective Y1 receptor antagonist [87, 88]. A second receptor mutation study at the NPY-Y1 receptor, based on those ﬁndings, showed that the agonist NPY and the antagonist BIBP 3226 share an overlapping, but not identical, binding site [89]. Whereas some mutations affected the binding of the endogenous ligand NPY only, others lead to decreased binding of both NPY and BIBP 3226, and one mutation affected the binding of the antagonist BIBP 3226 only. A similar approach was done with the NK-1 receptor and its peptide ligand substance P [90]. Important residues for the recognition of substance P were identiﬁed in the N-terminal extension, just outside the transmembrane domain 1 (TM-I), in the ﬁrst extracellular loop outside TM-II and at the top of TM-III. Substitutions of these residues lead to a dramatic loss of binding of substance P but did not affect non-peptide antagonist binding. The residues important for non-peptide antagonist binding were identiﬁed in the outer parts of TM-IV and TM-V. As in the case of the NPY-Y1 receptor, substance P and the antagonists are competitors at the NK-1 receptor but seem to bind differently. Strikingly, one of the antagonist compounds CP96,345, seems to share no interaction point at the receptor with substance P, despite being a competitive antagonist. This again may be explained by the hypothesis of conformational selection, with the antagonists stabilizing an inactive conformation of the receptor and thereby inhibiting the recognition of substance P. The introduction of some mutations interestingly did not affect the ability of radiolabeled substance P to bind to the mutant receptors but impaired its ability to compete with the radiolabeled non-peptide antagonists. The combination of receptor mutagenesis and ligand modiﬁcation helps to elucidate speciﬁc interactions between residues of the ligand and residues within the receptor sequence. 10.5 Cross-linking

Modiﬁcation of the ligand and receptor mutagenesis studies are indirect approaches for the analysis of molecular interactions between a receptor and its ligand. It can be difﬁcult to determine whether a change in afﬁnity is caused by an altered conformation of either the receptor or the ligand or whether it is due to changed direct molecular interactions. An alternative method to circumvent this problem is to covalently link a ligand to its receptor after incubation. The interaction site can then be determined. In principle two different proceedings are known for

24 Analysis of Receptor-Ligand Interaction

Fig. 12 (A) Model of photo-cross-linking of a peptide ligand to its receptor by the photoactivatable amino acid p-(3triﬂuoromethyl)-diazirinophenylalanine (Tmd(Phe)). In this example Tmd(Phe) is used to replace its homologue natural

amino acid tyrosine. (B) Reaction scheme of Tmd(Phe). After irradiation with UV light, Tmd(Phe) forms a reactive carbene, which then reacts with residues of the receptor sequence that are closely located in the receptor-ligand complex.

cross-linking. One is the use of an additional bifunctional photoactivatable linker molecule. This has the advantage that the ligand does not have to be modiﬁed. However, a limitation of this method is that bifunctional reagents often cross-link the ligand with the receptor at 14–16 Å from the binding site [1]. One way to circumvent this problem is to introduce photoactivatable groups into the ligand itself (Fig. 12a). For peptide ligands a number of photoactivatable amino acids are known that upon irradiation with UV light form highly reactive species, for example, a carbene, a nitrene, or a diradical [1]. The binding behavior of the respective photoactivatable analogues of a native ligand of course has to be characterized before performing a cross-linking experiment, e.g., in a radioligandbinding assay [91]. Provided the photoactivatable ligand shows binding behavior similar to the native ligand, it can be used for cross-linking studies. To enable detection of successful cross-linking, the introduction of a second label is favorable [92]. This can be a radioactive isotope, which has the advantage of a very low detection limit, a ﬂuorescent

10 Molecular Characterization of the Receptor-Ligand Interaction 25

dye, or a group with a deﬁned interaction pattern such as biotin or a histidine-tag. The latter ones have the advantage that they allow the speciﬁc puriﬁcation of the cross-linked receptor-ligand complex [93]. A standard procedure is to incubate the photoactivatable ligand with the receptor as in a standard binding assay [91]. The ligand’s photoactivatable group is then activated by irradiation with UV-light and thus is covalently cross-linked to close residues in the receptor sequence [91]. At this point it is of crucial importance to check for the speciﬁcity of the cross-linking, as most photoactivatable groups will react with any residue in their close environment [92]. Therefore, displacement assays with unlabeled ligands should be performed in parallel [92]. If the cross-linking was speciﬁc, it should be signiﬁcantly competed by an unlabeled ligand. Because photoafﬁnity labeling studies are usually performed with whole cells or membrane preparations, puriﬁcation of the cross-linked complex is necessary before further characterization of the interaction site is possible. This can be done by SDS-PAGE or via speciﬁc interactions of additional labels such as biotin or a histidine-tag. If radio or ﬂuorescent labeling is used for detection, the cross-linked receptors will be directly identiﬁed from the gel by phosphorimaging [94]. If labels such as biotin or antibodies are used for detection, the gel will have to be blotted onto a membrane ﬁrst, followed by staining with enzyme-coupled streptavidin or second antibodies (see Section 10.2.1). The corresponding bands on a silver or coomassie stained gel can be identiﬁed via their mass. The identiﬁed bands of the cross-linked receptors can then be excised from the gel, de-stained, and submitted to in-gel digestion by a proteolytic enzyme with speciﬁc cleavage sites, for example, trypsin [95, 96]. Trypsin speciﬁcally cleaves after lysine and arginine and therefore leads to deﬁned peptide fragments of the photoafﬁnity-labeled receptors. Before trypsin digestion, sulfur atoms of the receptor protein can be reduced and alkylated, e.g., with iodoacetamide, to avoid the formation of peptide dimers via disulﬁde bonds. Among the digestion fragments should then be a peptide that is cross-linked to the ligand. It can be identiﬁed by mass spectrometry (MS) analysis, either of the crude peptide mixture (MALDI-TOF-MS) [95] or after further puriﬁcation (ESI-MS), [97] or other methods. To facilitate the identiﬁcation of digestion products, a number of online devices for in silico digestion are available (e.g., at www.expasy.org). The experimentally found fragments can then be compared to the theoretical ones. The cross-linked fragment can be identiﬁed by its mass, which is the mass of one receptor fragment plus that of the label. A number of photoactivatable ligands have been used to identify binding sites of ligands at their receptors [92]. The choice of the photoactivatable group is strongly dependent on the ligand into which it is introduced. It should alter the binding properties and the conformation of the ligand as little as possible. Examples of photoactivatable amino acids are p-(3-triﬂuoromethyl)diazirinophenylalanine (Tmd(Phe)) and p-benzoylphenylalanine (Bpa), two analogues of the amino acid phenylalanine [98]. They can be used to replace Phe or its homologue Tyr, as was shown in cross-linking experiments with the NPY-Y2 receptor [91]. Whereas Tmd(Phe) reacts via formation of a carbene, Bpa forms a biradical upon irradiation with UV light (Fig. 12b).

26 Analysis of Receptor-Ligand Interaction

Examples for the identiﬁcation of G-protein-coupled receptor binding sites by photoafﬁnity labeling are the renal V2 vasopressin receptor [99] and the human brain cholecystokininB receptor [97]. 11 Conclusion

Despite the lack of crystallographic structural data for all G-protein-coupled receptors except rhodopsin, the information collected from the various experiments described here can be used to create models how GPCRs function and how the interaction between a ligand and its receptor takes place on the molecular level. The processing of these data not only requires careful evaluation by the scientist but also would be impossible without advanced computing technology. In combination, they lead to an evolving process of the formulation of new hypotheses and their experimental proofs. Nevertheless, direct structural determination of the receptor-ligand interaction remains a great aim for the future. The crystallization of rhodopsin was one step in this direction, and most likely other receptors’ crystallization will follow. Maybe other techniques for the elucidation of transmembrane receptors’ structures will be developed. However, as the interaction between a receptor and its ligand is always a dynamic process, techniques that monitor different aspects of this interaction will retain their importance. References 1 BECK-SICKINGER, A. G., Drug Discovery Today, 1996, 1, 502–513. 2 PALCZEWSKI, K., KUMASAKA, T., HORI, T., BEHNKE, C. A., MOTOSHIMA, H., FOX, B. A., LE TRONG, I., TELLER, D. C., OKADA, T., STENKAMP, R. E., YAMAMOTO, M. and MIYANO, M., Science, 2000 289, 739–745. 3 STRANGE, P. G., Trends Pharmacol. Sci., 2002, 23, 89–95. 4 SMIT, M. J., LEURS, R., ALEWIJNSE, A. E., BLAUW, J., Van NIEUW AMERONGEN, G. P., Van De VREDE, Y., ROOVERS, E. and TIMMERMAN, H., Proc. Natl. Acad. Sci. USA, 1996, 93, 6802–6807. 5 HALL, D. A. and STRANGE, P. G., Br. J. Pharmacol., 1997, 121, 731–736. 6 ROSSIER, O., ABUIN, L., FANELLI, F., LEONARDI, A. and COTECCHIA, S., Mol. Pharmacol., 1999, 56, 858–866. 7 CHIDIAC, P., HEBERT, T. E., VALIQUETTE, M., DENNIS, M. and BOUVIER, M., Mol. Pharmacol., 1994 45, 490–499.

8 WESTPHAL, R. S. and SANDERS-BUSH, E., Mol. Pharmacol., 1994, 46, 937–942. 9 MILANO, C. A., ALLEN, L. F., ROCKMAN, H. A., DOLBER, P. C., MCMINN, T. R., CHIEN, K. R., JOHNSON, T. D., BOND, R. A. and LEFKOWITZ, R. J., Science, 1994, 264, 582–586. 10 COHEN, D. P., THAW, C. N., VARMA, A., GERSHENGORN, M. C. and NUSSENZVEIG, D. R., Endocrinology, 1997, 138, 1400–1405. 11 ARVANITAKIS, L., GERAS-RAAKA, E. and GERSHENGORN, M. C., Trends Endocrinol. Metabol., 1998, 9, 27–31. 12 TURNER, P. R., BAMBINO, T. and NISSENSON, R. A., Mol. Endocrinol., 1996, 10, 132–139. 13 REN, Q., KUROSE, H., LEFKOWITZ, R. J. and COTECCHIA, S., J. Biol. Chem., 1993, 268, 16483–16487. 14 WESTPHAL, R. S., BACKSTROM, J. R. and SANDERS-BUSH, E., Mol. Pharmacol., 1995, 48, 200–205.

References 15 TUNG, M. F., Cellular Signalling, 1996, 8, 217–224. 16 TATEMOTO, K., CARLQUIST, M. and MUTT, V., Nature, 1982, 296, 659–660. 17 SAKURAI, T., Regul. Pept., 1999, 85, 25–30. 18 S¨OLL, R. and BECK-SICKINGER, A. G., J. Pept. Sci., 2000, 6, 387–397. 19 BIEDLER, J. L., HELSON, L. and SPENGLER, B. A., Cancer Res., 1973, 33, 2643–2652. 20 REYNOLDS, C. P., BIEDLER, J. L., SPENGLER, B. A., REYNOLDS, D. A., ROSS, R. A., FRENKEL, E. P. and SMITH, R. G., J. Natl. Cancer Inst., 1986, 76, 375–387. 21 ALALUF, S., MULVIHILL, E. R. and MCILHINNEY, R. A. J., J. Neurochem., 1995, 64, 1548–55. 22 JORDAN, B. A. and DEVI, L. A., Nature, 1999, 399, 697–700. 23 SAITO, Y., NOTHACKER, H. P., WANG, Z., LIN, S. H., LESLIE, F. and CIVELLI, O., Nature, 1999, 400, 265–269. 24 DUROCHER, Y., PERRET, S., THIBAUDEAU, E., GAUMOND, M.-H., KAMEN, A., STOCCO, R. and ABRAMOVITZ, M., Analytical Biochemistry, 2000, 284, 316–326. 25 GUJER, R., ALDECOA, A., BUEHLMANN, N., LEUTHAEUSER, K., MUFF, R., FISCHER, J. A. and BORN, W., Biochemistry, 2001, 40, 5392–5398. 26 MILLIGAN, G., Br. J. Pharmacol. 1999, 128, 501–510. 27 FUKUNAGA, K., ISHII, S., ASANO, K., YOKOMIZO, T., SHIOMI, T., SHIMIZU, T. and YAMAGUCHI, K., J. Biol. Chem. 276, 43025–43030. 28 TARASOVA, N. I., STAUBER, R. H., CHOI, J. K., HUDSON, E. A., CZERWINSKI, G., MILLER, J. L., PAVLAKIS, G. N., MICHEJDA, C. J. and WANK, S. A., J. Biol. Chem., 1997, 272, 14817–14824. 29 MOSER, C., BERNHARDT, G., MICHEL, J., SCHWARZ, H. and BUSCHAUER, A., Can. J. Physiol. Pharmacol., 2000, 78, 134–142. 30 CHO, W., BITTOVA, L. and STAHELIN, R. V., Anal. Biochem., 2001, 296, 153–161. 31 JOENSSON, U., FAEGERSTAM, L., IVARSSON, B., JOHNSSON, B., KARLSSON, R., LUNDH, K., LOEFAAS, S., PERSSON, B., ROOS, H., SJOLANDER, S., STAHLBERG, R., STENBERG, E., URBANICZKY, C., MALMQUIST, M.,

32

33 34 35

36

37

38

39

40

41

42 43 44 45

46

47 48

OSTLIN, H. and RONNBERG, I., Bio-Techniques, 1991, 11, 620–627. HESS, S. T., HUANG, S., HEIKAL, A. A. and WEBB, W. W., Biochemistry, 2002, 41, 697–705. CHENG, Y. and PRUSOFF, W. H., Biochem. Pharmacol., 1973, 22, 3099–3108. LAZARENO, S., J. Recept. Signal Transduct. Res., 2001, 21, 139–165. SCHULZ, R., WEHMEYER, A. and SCHULZ, K., J. Pharmacol. Exp. Ther., 2002, 300, 376–384. RIGLER, R., PRAMANIK, A., JONASSON, P., KRATZ, G., JANSSON, O. T., NYGREN, P., STAHL, S., EKBERG, K., JOHANSSON, B., UHLEN, S., UHLEN, M., JORNVALL, H. and WAHREN, J., Proc. Natl. Acad. Sci. USA, 1999, 96, 13318–13323. PRAMANIK, A., OLSSON, M., LANGEL, U., BARTFAI, T. and RIGLER, R., Biochemistry, 2001, 40, 10839–10845. INGENHOVEN, N. and BECK-SICKINGER, A. G., J. Recept. Signal Transduct. Res., 1997, 17, 407–418. FABRY, M., CABRELE, C., HOCKER, H. and BECK-SICKINGER, A. G., Peptides, 2000, 21, 1885–1893. FABRY, M., LANGER, M., ROTHEN-RUTISHAUSER, B., WUNDERLI-ALLENSPACH, H., HOCKER, H. and BECK-SICKINGER, A. G., Eur. J. Biochem., 2000, 267, 5631–5637. TURCATTI, G., NEMETH, K., EDGERTON, M. D., MESETH, U., TALABOT, F., PEITSCH, M., KNOWLES, J., VOGEL, H. and CHOLLET, A., J. Biol. Chem., 1996, 271, 19991–19998. OVERTON, M. C. and BLUMER, K. J., Current Biology, 2000, 10, 341–344. DEVI, L. A., Trends Pharmacolog. Sci., 2000, 21, 324–326. MYSZKA, D. G., Curr. Opin. Biotechnol., 1997, 8, 50–57. STENBERG, E., PERSSON, B., ROOS, H. and URBANICZKY, C., J. Colloid Interface Sci., 1991, 143, 513–526. PLANT, A. L., BRIGHAM-BURKE, M., PETRELLA, E. C. and O’SHANNESSY, D. J., Anal. Biochem., 1995, 226, 342–348. KARLSSON, O. P. and LOFAS, S., Anal. Biochem., 2002, 300, 132–138. FREI, N., WEISSENBERGER, J., BECK-SICKINGER, A. G., H¨OFLIGER, M.,

27

28 Analysis of Receptor-Ligand Interaction

49

50

51 52

53

54

55 56

57

58

59 60 61 62

63

64

65

WEIS, J. and IMBODEN, H., Regul. Pept., 2001, 101, 149–155. ECKARD, C. P., BECK-SICKINGER, A. G. and WIELAND, H. A., J. Recept. Signal Transduct. Res., 1999, 19, 379–394. WIELAND, H. A., ECKARD, C. P., DOODS, H. N. and BECK-SICKINGER, A. G., Eur. J. Biochem., 1998, 255, 595–603. ECKARD, C. P. and BECK-SICKINGER, A. G., Curr. Med. Chem., 2000, 7, 897–910. LEBESGUE, D., WALLUKAT, G., MIJARES, A., GRANIER, C., ARGIBAY, J. and HOEBEKE, J., Eur. J. Pharmacol., 1998, 348, 123–133. LEIBER, D., HARBON, S., GUILLET, J. G., ANDRE, C. and STROSBERG, A. D., Proc. Natl. Acad. Sci. USA, 1984, 81, 4331–4334. KUSAKABE, Y., ABE, K., TANEMURA, K., EMORI, Y. and ARAI, S., Chem. Senses, 1996, 21, 335–340. DUNBAR, B. S. and SCHWOEBEL, E. D., Methods Enzymol., 1990, 182, 663–670. EY, P. L., PROWSE, S. J. and JENKIN, C. R., Immunochemistry, 1978, 15, 429–436. FU, M. L., SCHULZE, W., WALLUKAT, G., ELIES, R., EFTEKHARI, P., HJALMARSON, A. and HOEBEKE, J., Receptors and Channels, 1998, 6, 99–111. BENITEZ, R., FERNANDEZ-CAPETILLO, O., LAZARO, E., MATEOS, J. M., OSORIO, A., ELEZGARAI, I., BILBAO, A., LINGENHOEHL, K., Van der PUTTEN, H., HAMPSON, D. R., KUHN, R., KNOPFEL, T. and GRANDES, P., J. Comp. Neurol., 2000 417, 263–274. JAKAB, R. L. and GOLDMAN-RAKIC, P. S., J. Comp. Neurol., 2000, 417, 337–348. INCE, E., CILIAX, B. J. and LEVEY, A. I., Synapse, 1997, 27, 357–366. KAMITANI, S. and SAKATA, T., Biochim. Biophys. Acta, 2001, 1539, 131–139. BJOERKLOEF, K., LUNDSTROEM, K., ABUIN, L., GREASLEY, P. J. and COTECCHIA, S., Biochemistry, 2002, 41, 4281–4291. JIN, H., ZASTAWNY, R., GEORGE, S. R. and O’DOWN, B. F., Eur. J. Pharmacol., 1997, 324, 109–116. FRASER, I. D. C., CONG, M., KIM, J., ROLLINS, E. N., DAAKA, Y., LEFKOWITZ, R. J. and SCOTT, J. D., Current Biology, 2000, 10, 409–412. KLUG, S. J. and FAMULOK, M., Mol. Biol. Rep., 1994, 20, 97–107.

66 TUERK, C. and GOLD, L., Science, 1990, 249, 505–510. 67 ELLINGTON, A. D. and SZOSTAK, J. W., Nature, 1990, 346, 818–822. 68 FAMULOK, M. and MAYER, G., Curr. Top. Microbiol. Immunol., 1999, 243, 123–136. 69 HERMANN, T. and PATEL, D. J., Science, 2000, 287, 820–825. 70 PROSKE, D., H¨OFLIGER, M., S¨OLL, R. M., BECK-SICKINGER, A. G. and FAMULOK, M., J. Biol. Chem., 2002, 277, 11416–11422. 71 FAMULOK, M., BLIND, M. and MAYER, G., Chem. Biol., 2001, 8, 931–939. 72 SCHWARTZ, T. W., Curr. Opin. Biotechnol., 1994, 5, 434–444. 73 YOKOTA, Y., AKAZAWA, C., OHKUBO, H. and NAKANISHI, S., EMBO J., 1992, 11, 3585–3591. 74 MAGGIO, R., VOGEL, Z. and WESS, J., FEBS Lett., 1993 319, 195–200. 75 MAGGIO, R., VOGEL, Z. and WESS, J., Proc. Natl. Acad. Sci. USA, 1993, 90, 3103–3107. 76 LEURS, R., SMIT, M. J., ALEWIJNSE, A. E. and TIMMERMAN, H., Trends Biochem. Sci., 1998, 23, 418–422. 77 PORTER, J. E., HWA, J. and PEREZ, D. M., J. Biol. Chem., 1996, 271, 28318–28323. 78 WALKER, P., MUNOZ, M., MARTINEZ, R. and PEITSCH, M. C., J. Biol. Chem., 1994, 269, 2863–2869. 79 MARSHALL, G. R., Biopolymers, 2001, 60, 246–277. 80 MARTINEZ, J., BALI, J. P., RODRIGUEZ, M., CASTRO, B., MAGOUS, R., LAUR, J. and LIGNON, M. F., J. Med. Chem., 1985, 28, 1874–1879. 81 CABRELE, C. and BECK-SICKINGER, A. G., J. Pept. Sci., 2000, 6, 97–122. 82 BARTFAI, T., LANGEL, U., BEDECS, K., ANDELL, S., LAND, T., GREGERSEN, S., AHREN, B., GIROTTI, P., CONSOLO, S., CORWIN, R. et al., Proc. Natl. Acad. Sci. USA, 1993, 90, 11287–11291. 83 BECK-SICKINGER, A. G., WIELAND, H. A., WITTNEBEN, H., WILLIM, K. D., RUDOLF, K. and JUNG, G., Eur. J. Biochem., 1994, 225, 947–958. 84 MONKS, S. A., KARAGIANIS, G., HOWLETT, G. J. and NORTON, R. S., J. Biomol. NMR, 1996, 8, 379–390.

References 85 BADER, R., BETTIO, A., BECK-SICKINGER, A. G. and ZERBE, O., J. Mol. Biol., 2001, 305, 307–329. 86 SAUTEL, M., MARTINEZ, R., MUNOZ, M., PEITSCH, M. C., BECK-SICKINGER, A. G. and WALKER, P., Mol. Cell. Endocrinol., 1995, 112, 215–222. 87 RUDOLF, K., EBERLEIN, W., ENGEL, W., WIELAND, H. A., WILLIM, K. D., ENTZEROTH, M., WIENEN, W., BECK-SICKINGER, A. G. and DOODS, H. N., Eur. J. Pharmacol., 1994, 271, R11– R13. 88 DOODS, H. N., WIENEN, W., ENTZEROTH, M., RUDOLF, K., EBERLEIN, W., ENGEL, W. and WIELAND, H. A., J. Pharmacol. Exp. Ther., 1995, 275, 136–142. 89 SAUTEL, M., RUDOLF, K., WITTNEBEN, H., HERZOG, H., MARTINEZ, R., MUNOZ, M., EBERLEIN, W., ENGEL, W., WALKER, P. and BECK-SICKINGER, A. G., Mol. Pharmacol., 1996, 50, 285–292. 90 ROSENKILDE, M. M., CAHIR, M., GETHER, U., HJORTH, S. A. and SCHWARTZ, T. W., J. Biol. Chem., 1994, 269, 28160– 28164.

91 INGENHOVEN, N., ECKARD, C. P., GEHLERT, D. R. and BECK-SICKINGER, A. G., Biochemistry, 1999, 38, 6897–6902. 92 DORMAN, G. and PRESTWICH, G. D., Trends Biotechnol., 2000, 18, 64–77. 93 BAYER, E. A. and WILCHEK, M., J. Chromatogr., 1990, 510, 3–11. 94 BOLT, M. W. and MAHONEY, P. A., Anal. Biochem., 1997, 247, 185–192. 95 SHEVCHENKO, A., WILM, M., VORM, O. and MANN, M., Anal. Chem., 1996, 68, 850–858. 96 DIHAZI, H., KESSLER, R. and ESCHRICH, K., Anal. Biochem., 2001, 299, 260–263. 97 ANDERS, J., BLUGGEL, M., MEYER, H. E., KUHNE, R., TER LAAK, A. M., KOJRO, E. and FAHRENHOLZ, F., Biochemistry, 1999, 38, 6043–6055. 98 WEBER, P. J. and BECK-SICKINGER, A. G., J. Pept. Res., 1997, 49, 375–383. 99 KOJRO, E., EICH, P., GIMPL, G. and FAHRENHOLZ, F., Biochemistry, 1993, 32, 13537–13544. 100 RIST, B., ENZEROTH, M. and BECK-SICKINGER, A. G., J. Med. Chem., 1998, 41, 117–123.

29

1

Protein Microarrays: Surface Chemistry and Coupling Schemes Michael Sch¨aferling Universit¨a Regensburg, Regensburg, Germany

Dev Kambhampati Alameda, USA

Originally published in: Protein Microarray Technology. Edited by Dev Kambhampati. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30597-1

1 Introduction

This review will focus on various strategies for constructing protein microarrays on a wide range of substrates. The way in which proteins are immobilized on solid substrates will determine the functional properties of a protein microarray. While protein interactions are currently conducted on a wide variety of formats, it is still a major challenge to obtain high quality biomolecular interaction data that can be used to predict how proteins behave in their biological environments. Two approaches for developing protein microarrays are currently underway. In the first approach the properties of the microarray surface are tailored to functionalize unmodified (pristine) biomolecules and in the second biomolecules are modified with tags to effectively anchor them to existing microarray or sensor substrates. Irrespective of which approach is chosen for a particular application, fundamental challenges still remain when it comes to interpreting the data. Extrapolation of protein immobilization strategies from one system to another for different classes of proteins and applications is difficult since proteins have a wide range of structural, chemical, and biological properties. Even within a single protein array application system, normalization studies (corresponding to probe surface concentration and activity) of a wide range of probes with different functional properties have to be considered. Sample preparation and quality control issues plague almost all microarray formats that are currently available. These processes are extremely time consuming and counteract the overall claims and advantages of such microarrays in functional high throughput screening.

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Protein Microarrays: Surface Chemistry and Coupling Schemes

A “one-approach-fit-all” scheme for screening all the various types of proteins is improbable. The current (and the most practical) approach is to generate surfaces that can be used for broad subsets of protein families without compromising their functional properties. In this article, protein chips constructed by using different immobilization chemistries on glass, gold, polymer and other special formats (semiconductor surfaces and lab-on-a-chip) will be discussed in detail. Various strategies for increasing the degree of probe immobilization and preventing protein denaturation by the use of effective tags will also be described. 1.1 Background and Current State of Biomolecule Libraries

Most large-scale biotechnology, pharmaceutical and research laboratories currently have existing libraries of both pristine proteins and those conjugated with various tags (biotin, His-Tag, Strep-Tag, Avi-Tag, etc.). Since these molecules have been extensively characterized, such laboratories will be reluctant to alter them to suit the surface immobilization requirements of microarray platforms. As a result, surface immobilization strategies on microarrays have to be designed on the basis of the existing state of biomolecule libraries where there is limited flexibility for operation. Most of the microarray schemes that will be described here will focus on the abovementioned existing biomolecule libraries. Almost all kinds of proteins have reactive functional groups in their side chains (Table 1) which can be used as a starting point for anchoring these molecules on microarrays. In the case of biomolecules conjugated with tags, the immobilization approach proceeds with a higher surface coupling efficiency (when compared to pristine probes). The drawback in this case is that the inherent biomolecular interaction properties of the conjugated probes Table 1 Various methods of coupling proteins on a microarray platform

Functional side group on peptide

Available surface for derivatization

Type of binding

Natural –COOH (carboxylic acid) Asp

Amino

Electrostaticy Covalent amide (after carboxy activation) Electrostatic

–NH2 (amino) Lys, Gln, Arg –SH (thiol) Cys –OH Ser, Thr Synthetic His-Tag Strep-Tag Biotin

Carboxylic acid, active ester, epoxy, aldehyde Maleimide

Covalent amide Covalent thio ether

Epoxy

Covalent ether

Ni-NTA complex Strep-Tactin Streptavidin

Coordination complex Supramolecular complex Supramolecular complex

2 Microarrays Based on Glass Substrates

might be altered, thereby affecting their kinetic and thermodynamic properties. Irrespective of the nature of the probes, the overall goal is to target reactive sites which do not denature the molecules or affect the complementarity-determining regions (CDRs) of the probes during surface immobilization. Numerous surface chemistry approaches are currently available to address such issues.

2 Microarrays Based on Glass Substrates

Most of the current analysis of protein interactions on microarrays are based on surfaces and formats that have been used for DNA arrays. Glass slides coated by epoxy- or aldehyde-terminated alkylsilanes or by polylysine can also be used to immobilize proteins, antibodies and antigens. Such activated glass (microscope slide format) slides can be spotted with biomolecules by the end-user in any format and density. The detection mode for such systems is based mainly on optical techniques such as fluorescence. Table 2 lists some examples of commercially available glass slides with reactive surfaces. 2.1 Surface Modification

The reactive coatings of the glass surface are mainly based on self-assembly techniques. Self-assembled mono-layers (SAMs) of alkylchlorosilanes or alkylalkoxysilanes require hydroxylated surfaces as substrates for their formation. The driving force for this self-assembly is the in-situ formation of polysiloxane, which is connected to surface silanol groups (SiOH) via Si-O-Si bonds (Scheme 1). These thin organic films enhance the biocompatibility of the surface and protect the proteins from denaturation and structural changes during the immobilization step. A great variety of silanization conditions pertaining to temperature and reaction time, which are also strongly influenced by the stability of the terminal functional groups have been described in the literature [1]. Monolayers of aminopropylsilanes, Table 2 Examples of commonly available glass-based microarray platforms

Coating

Supplier

Advantages

Disadvantages

Poly-L-Iysine

Sigma, Fisher

Cheap, easy to handle

Only electrostatic absorption, low stability

Highly reactive, great versatility

Low spot homogeneity, susceptible to contamination

Aminopropylsilane Corning, Erie, Xomics Aldehyde-, NHSGeneScan, Xomics ester- or epoxysilane GeneScan, Xomics Genetix NoAb BioDiscoveries Telechem, Quantifoil

3

4 Protein Microarrays: Surface Chemistry and Coupling Schemes

Scheme 1. 2D schematic description of a polysiloxane monolayer on a glass surface (X = terminal functional)

for example, can be formed under mild conditions (room temperature for 30 min) [2]. Scheme 2 shows some commonly-used silane-reagents used for the derivatization of glass surfaces [3]. Substrates on which these organosilicon monolayers have been successfully prepared include silicon oxide, aluminium oxide, quartz, glass, mica, zinc selenide and germanium oxide [4, 5]. Silicon substrates can also be covered with silane monolayers [6], resulting in a new approach for tuning the electronic properties of semiconductor chips [7] and for challenging new interfaces in the field of bioelectronics.

Scheme 2 Reagents for derivatization of glass surfaces. 1 APTES = aminopropyltriethoxysilane; 2 MPTS = 3mercaptopropyltrimethoxysilane; 3 GPTS = glycidoxypropyltrimethoxysilane; 4 TETU = triethoxysilane

undecanoic acid; 5 HE-APTS = bis(hydroxyethyl)aminopropyltriethoxysilane); 6 4-trimethoxysilylbenzaldehyde; 7 GPTS/HEG = glycidoxypropyltrimethoxysilanehexaethylene glycol; 8 poly(lysine).

2 Microarrays Based on Glass Substrates

2.2 Current State of Glass-Based Protein Microarrays

As in the case of DNA-arrays, complex issues pertaining to spot morphology, homogeneity of signal across the spot, microarray smearing and non-specific adsorption are not adequately addressed in the case of protein microarrays. In addition to the above challenges, it is difficult to compare the experimental results obtained in two different laboratories using the same set of arrays. Reproducibility of data is a major concern and if this current trend continues, it will become extremely difficult to accurately validate the biomolecular interactions that form the basis of many drug discovery and research studies. Preliminary studies [8] pertaining to the use of polylysine arrays in monitoring the interactions of 115 antigen/antibody pairs using complex protein mixture solutions was recently reported. While the success rate of the interaction data was between 20 and 50%, concerns pertaining to microarray smearing and background still remain (Figure 1a). Microarray smearing is a common artifact found in most glass-based arrays and it is one of the methods by which cross-talk between the adjacent probes occurs, leading to false results. In addition to microarray smearing, the electrostatic charges present on the glass can

Fig. 1 (a) Smearing effects on antibody-immobilized polylysine arrays [8]. (b) Snapshot of microarray spots with poor morphology and surface chemistry (within the individual spots). Differences in spot homogeneity can result in varying signal intensities within the same spot [9].

5

6 Protein Microarrays: Surface Chemistry and Coupling Schemes

also lead to the denaturation of proteins, although the exact level of electrostatic charge is difficult to estimate. In another study, large-scale protein–protein interaction studies were also successfully conducted on aldehyde-coated glass slides [9]. While the slides were successful in discriminating between the specific and non-specific interactions, issues pertaining to “quenching” or neutralizing the remaining reactive groups on the slides still remain. BSA was used to effectively block the free surface of the chip to minimize non-specific interactions. This could pose a problem on such slides if small molecules are used which can potentially be masked by the bulky BSA molecules. While spot morphology is not a major concern in normal laboratoryscale research studies, it becomes critical when investigating several thousands of protein samples spotted on a single microarray slide. Fluorescence intensities emanating from the spots (Figure 1b) and their adjoining areas (if the spots are not homogeneous) can affect the data acquisition process. In addition, if the surface chemistry within the spots is not homogeneous, varying intensities of fluorescence signal can be observed within the same spot. Unless the data analysis software is highly sophisticated, such spots can lead to poor averaging effects during the analysis step which will in turn result in misleading protein expression levels for the protein of interest. In another study, Joos et al. [10] reported the direct parallel detection of autoantibodies in human sera by using a single immunoassay experiment in which 18 different antigens were spotted onto aldehyde-coated slides. After activation of a 96-well teflon-coated glass plate (Erie Scientific) with aminopropylsilane, protein antigens were attached to the pre-defined spots to form a high-throughput microarray-based enzyme-linked immunosorbent assay (ELISA) [11]. Inside the wells, the authors spotted arrays of 144 (4 36) elements each using a 36-capillary based print head. This array format thus appears as a compromise between high-density microarrays and conventional micro-titerplates. Antibodies can also be arrayed on aminosilated glass surfaces using a micro-molded hydrogel stamper [12]. Many approaches use micro-contact printing with PDMS stamps [13] or photoreactions in combination with masks [14] for the patterning of glass surfaces with proteins. The profiling of cancer cells from a cellular lysate was carried out with aldehyde- or polylysine-coated glass microarray containing 146 distinct antibodies. The target proteins were labeled with Cy5 or Cy3 dyes and no amplification methods such as ELISA were needed for monitoring the interactions [15]. In another interesting study, Schweitzer et al. designed microarrays containing 75 antibodies against cytokines on chemically-derivatized glass slides to study cytokine secretion from human dendritic cells induced with lipopolysaccharide or tumor necrosis factor-α [16]. The superiority of nickel complex-coated surfaces over conventional aldehyde slides was reported by Snyder in his study on a proteome chip containing 6566 GST yeast proteins spotted onto a glass slide (Figure 2). In this particular case, the fusion proteins attach via their His-Tags and it is believed that the binding sites orient away from the surface [17]. This example shows that the use of tags can help to control the orientation of the immobilized biomolecules on the surface.

3 Microarrays based on Gold Substrates

Fig. 2 GST yeast protein analysis. (A) 60 samples were examined by immunoblot analysis using anti-GST; 19 representative samples are shown. (B) 6566 protein samples representing 5800 unique proteins were

spotted in duplicate onto a single nickelcoated microscope slide which was then probed with anti-GST. (C) Enlarged image of one of the 48 blocks.

3 Microarrays based on Gold Substrates

The use of gold-based substrates for monitoring protein interactions has a relatively long history. Instead of using polycrystalline gold surfaces, thin gold films (10– 200 nm) can easily be deposited on solid supports similar to glass or silicon wafers by evaporation or sputtering in an ultrahigh vacuum (UHV) [18]. Since the adhesion of gold on glass or silicon is poor, pre-coating of a thin layer of Cr or Ti (1–10 nm) is generally required prior to gold deposition. Since the work of Allara [19] and Whitesides [20] in the mid-1980s, the spontaneous adsorption of organosulfur compounds onto gold has been a widely used method for the preparation of self-assembled monolayers (SAMs). If the alkyl chain is of sufficient length, the resulting molecular architecture is very stable and orients itself nearly perpendicular to the surface [21]. The close proximity of the adsorbed molecules causes van der Waals forces to play an important role in intermolecular stabilization. The thickness of these monolayers is typically in the nanometer range in the vertical direction and as a result, they can be termed nanoassemblies. It is important to note however, that in the horizontal plane, i.e. over the substrate surface, these monolayers can extend to a macroscopic length. Although SAMs have been constructed on a wide range of surfaces, monolayers based on alkanethiols on gold have been investigated extensively for the following reasons: (1) ease of fabrication, (2) degree of perfection, (3) chemical stability under laboratory

7

8 Protein Microarrays: Surface Chemistry and Coupling Schemes Table 3 Examples of commercially available gold-based microarray platforms

Coating

Supplier

Advantages

Disadvantages

Functionalized alkanethiols

Zyomyx

Good control over surface quality, good biocompatibility

More expensive than glass or polymer arrays

Graffinity

conditions, (4) availability of materials, and (5) flexibility in introducing various chemical functional groups to give different surface properties. They provide excellent control over the surface properties by individually tailoring/engineering the building blocks. Using these principles, customized surfaces with good biocompatibility properties can be developed to meet the complex needs of proteomic studies. Unlike glass- or polymer-based arrays, SAMs on gold also have the added advantage of the surface properties being efficiently characterized by optical, mechanical and electrochemical analytical tools. Many commercial suppliers offer gold-coated microscope slides with sufficiently high surface quality. In spite of the above advantages, gold-based microarrays still are not in widespread use due to their cost and stability (chemical and thermal). A major limitation to the use of SAMs as an immobilization matrix is their low temperature stability due to the weak bonding (interactions) between sulfur and gold. While this is not a concern in the case of protein arrays (where experiments are conducted at low temperatures), such microarrays are found to be unsuitable for DNA array applications where high stringent wash and temperature conditions are employed. Because of the excellent electronic properties of gold, these surfaces can be used in more advanced and label-free detection technologies such as Surface Plasmon Resonance (SPR, Graffnity, Table 3. A reference to Biacore technology is provided in Section 4.1) and mass spectrometry (refer to Section 5). 3.1 Surface Modifications

Natural proteins contain a wide range of functional groups on the side chains of their polypeptide backbone (refer to Table 1). In principle, all of them can be used during the direct chemical coupling reaction on specially prepared SAM surfaces with appropriate terminal reactive groups. Scheme 3 shows the various terminal functional groups of alkanethiols that are potentially available for biosensor applications. Hydroxy terminal alkanethiols (oligo ethylene glycol units [22] and mannitol groups [23]) undergo low non-specific interactions with proteins and enzymes, and as a result are excellent candidates for making mixed monolayer-based protein array platforms. For other surface modifications which resist protein adsorption, refer to the extensive review by Whitesides [24].

3 Microarrays based on Gold Substrates

Scheme 3 Schematic respresentation of long-chain alkanethiol monolayers (e.g. 16 mercapto-hexadecanoic acid) on gold with different terminal functional groups. 1, biotin; 2, NHS-ester (NHS, Nhydroxysuccinimide); 3, epoxy-ethylene glycol; 4, maleimide; 5, diaminohexane; 6 oligo (ethylene glycol).

3.2 Current State of Gold-based Protein Microarrays

The most widely used gold-based protein microarrays are based on biotin–streptavidin technology. As a result, the following discussion will focus primarily on this technology. One of the reasons for the rapid growth in the application of biotin–streptavidin technology is the ease with which it is possible to biotinylate various biomolecules. The following two premises are, however, inherent when we talk about biotin-conjugated molecules: (a) the biochemical and physical properties of the biomolecules are not affected significantly by the biotinylation step and (b) the properties of the biotin moiety are not affected by the derivatized biomolecule. The relative ease of attachment of biotin to organic molecules substituted with a thiol makes it possible to use gold substrates as the underlying platform for constructing protein arrays. The streptavidin surface (formed on top of the underlying biotin-terminated SAM) can be used for immobilizing a wide range of biotinylated molecules (nucleic acids, proteins, saccharides, etc.). Single component SAMs containing biotin tethered at the terminal end are known to generate poor streptavidin monolayers due to steric constraints imposed

9

10 Protein Microarrays: Surface Chemistry and Coupling Schemes

by the tight biotin layer. The close-packed biotin layer shields the individual biotin units and also prevents the effective docking of the bulky streptavidin molecules. This problem can be obviated by using either spacer molecules or via surface coupling schemes that involve lower yields. The research groups of Knoll [25] and others [26] have actively investigated the interaction of biotin and streptavidin monolayers on a variety of sensor formats. By exploring many different combinations of biotinylated thiols and various spacer molecules, Knoll and co-workers have shown that an optimum monolayer of streptavidin can be generated by using a 1:9 mole ratio of biotin and hydroxyl-terminated thiols. The latter have a dual function: first, they act as efficient spacers between the biotinylated components and second, they minimize the non-specific interaction of streptavidin with the sensor substrate. Other research groups [27], using various streptavidin mutants, have indicated that an optimum streptavidin monolayer can be obtained by using approximately 10–25% biotin on the surface. Another method of generating a well-packed streptavidin surface is by using a surface reaction between a hydrophobic interface (Figure 3) comprising carboxy groups [28] and biotin-terminated linker units attached to amino groups. A denselypacked streptavidin surface is obtained when the surface coupling yield of biotin is between 25 and 30%. This value is in the same range as that of the mixed SAM layers. An application of this surface in screening patients affected with Lyme Borreliosis is shown in Figure 4. No blocking solutions were required during this assay and highly reproducible results with high signal-to-noise ratios were observed because the assay surface was quite homogeneous (i.e. uniform probe packing density) [29].

Fig. 3 Schematic respresentation of a steptavidin sensor surface assembled on a reaction-controlled biotinylated SAM [28].

3 Microarrays based on Gold Substrates

Fig. 4 Applacation of stretavidin sensor platform (refer to Figure 3) for the diagnosis of Lyme Borreliosis. a-e, different patients suffering from the disease; f and g, control experiments using healthy subjects. The difference in the above biomolecular in-

teraction curves can be attribued to patients at various stages of infection. Irrespective of the stage of infection, good signal-to-noise ratios can be observed between infacted patients and healthy subjects using the streptavidin microarray platform [29].

While both the above methods can be used in assembling a streptavidin surface, the mixed SAM approach is relatively easy to construct and can thus accelerate the production process. The drawback is that these molecules are not readily available and as a result, need to be specially synthesized. The synthesis of these molecules is not trivial and the synthesis steps usually have poor success (yield) rates. These problems can be overcome by using the readily available carboxy-terminated thiols for constructing the streptavidin sensor surface. It is important to note that both these techniques result in a high quality streptavidin surface. The biocompatibility of the SAM interface is likely to make such sensor surfaces interesting candidates for the screening of antibodies, patient sera and other complex proteins. Several reports dealing with surface patterning of proteins on gold-coated surfaces using particle lithography [30], photoimmobilization [31], chemical selectivity of mixed monolayers [32] and microcontact printing have been published [33]. Atomic Force Microscopy (AFM) [[33], 26b] or Scanning Tunneling Microscopy (STM) [34] techniques can also be used for direct visualization of such protein patterns on gold surfaces. In the latter case, a thiolated protein was directly adsorbed onto the surface. Issues pertaining to protein denaturation still remain as the study involved the direct contact of the proteins with the metal surface. Gold-coated electrodes can also serve as amperometric biosensors in the form of enzyme electrodes. The enzymes on such biosensors are immobilized via functionalized SAMs [35], conducting polymers such as polypyrrole or polythiophenes [36] or by direct absorption onto the electrode surface [37]. Some prominent examples include the glucose sensor based on the electrochemical detection of enzymaticallyproduced hydrogen peroxide [38] or the screening of neurotoxins with immobilized acetylcholinesterase in the form of competitive assays (using acetyl-thiocholine)

11

12 Protein Microarrays: Surface Chemistry and Coupling Schemes

[39]. However, none of these approaches is likely to be used in a microarray format for HTP studies in the near future.

4 Microarrays based on Polymer Substrates

Most of the above discussions have focused primarily on two-dimensional (2-D) surface architectures. Since the functional properties of a protein are highly dependent on its conformation and structure, it is possible that delicate proteins may not be able to survive the harsh conditions of 2-D surfaces. As a result, other formats based on three-dimensional (3-D) surfaces which provide a fluidic environment that closely mimics the biological environments of various proteins, have been proposed. Some prominent examples of 3-D-based systems include hydrogels and polymer membranes. The basic underlying tenet of such 3-D systems is that a fluidic environment preserves the conformation and, as a result, the activity of the protein. Additional benefits of 3-D architecture include larger surface area and greater probe density. Polymer membranes such as aminated nylon [40] or nitrocellulose [41] were the first commonly used substrates for the preparation of microarrays. For over two decades, these platforms have been widely employed in microbiology laboratories as blotting supports. Detection on these supports has been carried out mainly by radioimaging, although a combination of other detection technologies such as fluorescence is also possible. Table 4 provides an overview of the polymer microarray platforms that are currently available on the market. Gel-based studies in combination with downstream operations such as mass spectrometry, dominate most proteomic investigations. Since it is possible to transfer proteins and antibodies directly onto the blotting membranes after electrophoresis, these platforms definitely have certain advantages when it comes to sample preparation and in terms of the overall cost. Hydrogel-based arrays based on gold substrates have dominated the field of real-time protein interaction studies for the past decade. This has been mainly possible due to the widespread application of SPR Table 4 Examples of common polymer and hydrogel substrates

Polymer substrate

Supplier

Nylon (aminated) membrane

Pall, Macherey & Nagel Cheap, versatile, high biocompatibility, applicable to blotting Schleicher and Schuell Pierce, Bio World Nunc BiaCore Perkin Elmer Prolinx

Nitrocellulose membrane Polystyrene Dextran-Hydrogels Polyacrylamide Polymer brush

Overall advantages

General limitations Only electrostatic absorption, low spot homogeneity and reproducibility, mass transport limitations, high background smearing and non-specific interactions

4 Microarrays based on Polymer Substrates

technology by Biacore. Polymer brushes and hydrogels on glass substrates (conventional microarray format) have also been developed (refer to Table 4), although their usage is still limited. 4.1 Surface Modifications

The immobilization of biomolecules on functionalized polymer membranes such as nylon or cellulose occurs via electrostatic absorption. This is also the case for hydrogels based on polyacrylamide platforms. As a result, it is difficult to obtain reliable, high quality data from such systems for potential application in microarray studies. During the past decade, self-assembled monolayers modified with dextran have become the system of choice for many biomolecular interaction studies. The widespread use of this 3-D architecture was mainly a result of the enormous success of the commercial SPR instrument maker, Biacore. Well-established protocols of self-assembly techniques on gold were utilized by researchers at Biacore to synthesize these dextran surfaces [42]. Epoxy-terminated thiol monolayers were used for coupling dextran polymers to the sensor surface. The immobilization of ligands was achieved by using carboxy-methyl-modified dextran containing extensive reactive handles for activation and covalent attachment of the molecules of interest. The resulting layer is a hybrid material composed of a 2-D monolayer on gold and a 3-D hydrogel which is highly flexible, non-crosslinked and extends 100–200 nm from the coupling surface under physiological buffer conditions (Scheme 4). While such 3-D surfaces have numerous benefits, they are still plagued by problems relating to mass transport effects and high background signals emanating from non-specific interactions. The 3-D dextran surfaces are known to impose mass transfer limitations that can significantly alter the inherent kinetics of the interaction. Diffusion effects play an important role within these 3-D structures, and can lead to the formation of several local concentration gradients. During realtime kinetic studies using surface plasmon resonance measurements, such local gradients can result in the calculation of false kinetic rate constants. Protein immobilization with the help of functionalized polymer brushes or by photochemically-activated polymers will be discussed in Sections 4.2 and 6. 4.2 Current State of Polymer-based Microarrays

Numerous protein chip surfaces can be created by activating the above-mentioned dextran surface. Using the dextran as the base, surfaces incorporating thiol, amine, (strept)-avidin or aldehyde groups can be generated (Scheme 5). These surface modifications have resulted in the commercial success of numerous sensor chips [43] which are routinely used in a plethora of applications ranging from HIV analysis using various monoclonal antibody–antigen interactions to the area of small molecule drug discovery operations and quality control. Since these sensor chips are quite robust and can withstand stringent acidic or basic regeneration

13

14 Protein Microarrays: Surface Chemistry and Coupling Schemes

Scheme 4 Reaction schemes for preparing a hydrogel dextran matrix [41].

steps, they can be re-used several times without any substantial loss of the surface density/activity of the ligands. Another new approach that combines self-assembled monolayer techniques with a 3-D microenvironment is the synthesis of functionalized dendrimers on solid supports such as glass. The activated dendritic structures are terminated by NHS esteror isothiocyanate- groups [44]. Emerging technologies use the (co)polymerization of methacrylate, styrene or vinyl alcohol monomers on the surface of a glass chip followed by subsequent functionalization using photografting [45]. The optical properties of the chip surface can be tailored to the reflectance of light or scattering effects. The tuning of the refractive index of the surface can also be accomplished by effective implementation of high-refractive index materials (TiO2 ) or low-refractive layers (polymers, SiO2 ) [46]. This approach has been used to make highly reflective microarray surfaces at the applied wavelength (emission and excitation) for potential applications in highly sensitive luminescent-based assay systems. A comparative study involving the screening of 11 different array surfaces on polymer gel and silane-coated glass slides was carried out by applying five different monoclonal antibodies in a 20 × 18 matrix. All the important aspects of the procedure are discussed in this work, including ease of chip preparation and handling, storage, signal intensities, reproducibility and detection limits [47]. Based on these investigations, reflective slides with a mirror-like surface (3-aminopropyltriethoxysilane, Amersham Biosciences [48]) were found to provide high levels of

5 Special Formats: Microfluidic Devices and Integrated Semiconductor Chips

Scheme 5 Reaction schemes for generating different protein chip surfaces using the dextran matrix as the underlying platform.

uniformity and reproducibility, whilst the polyacrylamide-coated slides were characterized by their high levels of sensitivity. The authors conclude by stating that the selection of the array support material strongly depends upon the special requirements of the experiment and the properties of the probe biomolecules. Currently, many groups have reported on the fabrication of custom-made antibody arrays on nitrocellulose or nylon membranes for the detection of cytokines in conditioned media or human serum using a multiplexed sandwich assay [49].

5 Special Formats: Microfluidic Devices and Integrated Semiconductor Chips

The lab-on-chip technology involving microfluidics has evolved rapidly over the last decade. This technology allows for the parallelization and integration of multiple sample processing approaches in the field of diagnostics or proteomic research. These devices are however, more complex than conventional microarrays (Figure 5), and while they are suitable for small-scale sample preparation applications, their throughput is still relatively low. Nevertheless, as these systems continue to move from academic laboratories to commercialized technology, the user-friendliness of such microfluidic systems is slowly improving [50]. In spite of the above limitations, this technology has certain advantages such as the use of very small volumes of human sera or urine samples for conducting rapid diagnostic assays. This technology is poised to make rapid advances in the area of nucleic acid-based testing, a field that is set to explode during this decade. The lab-on-a-chip SNP genotyping systems by

15

16 Protein Microarrays: Surface Chemistry and Coupling Schemes

Fig. 5. Schematic diagram of a microfluide system for the analysis of DNA samples. Image by courtesy of US DOE Genomes to Life Program

Caliper [51] and Nanogen [52] are clear illustrations of the scope of such technologies. Whilst most of the early microfluidic formats have focussed primarily on DNA interactions, such systems can also be used for the detection of protein–protein or antibody–antigen interactions if appropriate surface immobilization strategies are employed. While Nanogen uses a plastic cartridge in their silicon wafer biochip unit, many microfluidic systems are based on glass or silicon using classical etching and processing technologies of the semiconductor industry for the structuring of chips with microchannel systems. The planar waveguide glass chips from Zeptosens use integrated microfluidics combined with high refractive index material (Ta2 O5 ) deposited on the glass support for monitoring biomolecular interactions using highly sensitive fluorescence detection methods [53]. The Flow-thru ChipTM from MetriGenix is also based on a three-dimensional microchannel glass chip [54]. CombiMatrix [55] attaches a proprietary porous three-dimensional reaction layer to the active side of semiconductor surfaces for the immobilization of biomolecules. Yet another microfluidic protein biochip platform based on silicon is currently under development at Zyomyx [56]. Microchannels in polymeric materials such as poly-(methyl methacrylate) (PMMA) can also be used to make microfluidic devices and at the same time the methyl ester groups on the surface of the PMMA can be chemically derivatized with reactive groups to immobilize biomolecules [57].

6 Chemical Immobilization Techniques for Proteins

Some of the potential advantages of miniaturization can also be combined with other detection technologies for effective analysis of protein purification and interactions. Two examples of this are the coupling of microfluidics with ionization mass spectroscopy (ESI-MS) [58] or with micro-capillary electrophoresis [59]. Another technology that has been borrowed from the semiconductor industry in the field of integrated microarrays is the fabrication of microelectrode structures on silicon wafers. These gold, silver or platinum electrodes can be coated with functionalized monolayers or conducting polymers that are biocompatible and reactive towards biomolecules. On such electrodes, a voltage can be applied for the selective immobilization of the physiologically charged biomolecules. The microarray system from Nanogen is one example of such an approach, while the chip readout is still undertaken using conventional optical techniques [60]. Other approaches are one step ahead and use electrochemical detection of DNA hybridization in combination with redoxactive labels [61] (such as Motorola’s eSensorTM DNA detection system) or intercalators [62]. Currently these technologies are primarily focused on DNA arrays although in future, additional solutions for monitoring protein–protein or antibody–antigen interactions can also be envisaged by using such techniques. Technologies such as impedance spectroscopy or surface capacitance measurements [63] can potentially be the key to these application (e.g. at BIONCHIP [64]). In analogy to luminescence-based assays, electrochemical detection methods can also be amplified by ELISA techniques. Enzymes such as horseradish peroxidase or alkaline phospatase can be conjugated to the target molecules, and the reaction products can be measured by impedance spectroscopy [65] or with amperometic methods [66] if a suitable substrate has been added. A fully integrated electronic CMOS (Complementary Metal Oxide Silicon) biochip (Figure 6) based on enzyme-linked sensor techniques has been recently launched by Infineon [67].

6 Chemical Immobilization Techniques for Proteins

Generally, all the functional surface coatings described above can act as biocompatible interfaces during the chemical coupling of proteins to any microarray substrate, either via covalent binding, complex coordination chemistry or supramolecular interactions. In addition to the above methods, proteins also have a tendency to interact with the surface via electrostatic interactions. This principle is used in the case of simple electrostatic adsorption of proteins onto nylon, nitrocellulose, polyL-lysine or aminopropylsilane-coated surfaces. These interactions constitute what is widely known as non-specific interactions. Special SAMs based on polyethylene glycols [68] or poly(L-lysine)-g-poly(ethylene glycol) [69] are known to resist such interactions and are thus widely used in the design of most biosensor and microarrays. In the following sections, various strategies for coupling proteins using covalent, photochemical tags and site-directed immobilization approaches will be addressed.

17

18 Protein Microarrays: Surface Chemistry and Coupling Schemes

Fig. 6 CMOS-based biochip platform with integrated electronic microstructure (gold on silicon) suitable for an electrochemical or optical chip read-out by Infineon (top) and Bionchip (bottom). Images by coutesy of Infineon and Bionchip BV.

6.1 Covalent Chemical Coupling

In order to achieve covalent chemical coupling to microarray substrates, proteins should have the following chemical functionalities (see also Table 1) in the side chains of their polypeptide backbone: SH (Cysteine), NH2 (Lysine), COOH (aspartic acid, glutamic acid), OH (Serine), Ph-OH (Ph = phenyl, Tyrosine), imidazole (Histidine). In principle, all of them can be used during the direct chemical coupling reaction on specially prepared SAM surfaces. NHS (N-hydroxysuccinimide ester), aldehyde or epoxy groups form stable amide bonds with the free amino groups of the peptide (Scheme 6). The activation of carboxyl groups with EDC (N-3-dimethylaminopropyl-N-ethylcarbodiimide) and NHS, as well as further coupling schemes with NHS-modified surfaces are shown in Scheme 5. In another approach, the carboxylic acid groups in the side chains of the polypeptide can be activated by carbodiimides or NHS and can be covalently coupled to free amino groups on the sensor surface.

6 Chemical Immobilization Techniques for Proteins

Scheme 6 Surface coupling reaction of NHS-esters with the amino residues of the side-chains of polypeptides (lysine units). R, protein.

Other chemicals which are currently available for the covalent coupling of biomolecules to microarray surfaces include maleimides, pyridyl disulfide and vinyl sulfone, all of which will react with thiol groups [70] (Scheme 7). Epoxymodified surfaces can also be used for coupling hydroxy groups (see Scheme 4) and isothiocyanates for binding with amino groups [71] (Scheme 8). 6.2 Photochemical Cross-Coupling

In principle, there are many approaches available for the photochemical crosscoupling of biomolecules to specially functionalized surfaces. For example, the binding of biomolecules to epoxy- or aldehyde-coated slides can also be induced by irradiation with UV light. But there are also some typical photoreactive groups

Scheme 7 Surface modifications for the attachment of thiol residues (which are present in the side chains) of polypeptides (cysteine units). R, protein, 1, maleimide; 2, disulfides; 3, vinyl sulfone.

19

20 Protein Microarrays: Surface Chemistry and Coupling Schemes

Scheme 8 Immobilization of proteins and enzymes on aminoterminated monolayers using the bifunctional reagent DIDS (4,4 diisothiocyanato-trans-2,2-disulfonic acid disodium salt).

that are currently available. A prominent example is the azidophenyl group. Photoinduced crosslinking of proteins with azidophenyl-functionalized layers can be achieved by irradiation with UV or visible light. The excitation wavelength of this reaction can be changed by the use of additional substituents on the azidophenyl ring [72] (Scheme 9). Another strategy for the photochemical attachment of biomolecules has been developed by researchers at the commercial firm, Exiqon. They have taken advantage of an anthrachinone linker that can be coupled to a wide variety of solid polymeric supports (e.g. polystyrene, polyethylene, polypropylene, PMMA, nylon) by irradiation (Scheme 10) and can be conjugated with either biomolecules, ligands or reactive groups (e.g. carboxylic acids or amines) [73]. Yet another versatile method for the photochemical cross-linking of proteins to surfaces is the photoaptamer technique used by SomaLogic. However, in contrast to the above methods where the capture probes are photochemically immobilized, here the molecular recognition event is stabilized by consecutive photocrosslinking. Photoaptamers are chemically synthesized single-stranded DNA molecules that can be labeled at their 5 -end with acrydite, amine, thiol or biotin. These labels facilitate the covalent attachment of the photoaptamers to solid supports

Scheme 9 Photo-induced cross-linking of proteins R and azidophenylfunctionalized layers (8 = 265–275 mm; if the hydroxyl substituent is replaced with a nitro group, then 8 = 300–460 mm).

6 Chemical Immobilization Techniques for Proteins

Scheme 10 Exiqon’s patented anthrachinone (AQ) covalent attachment photochemical method. R, polymeric solid; P, specific ligand (biomolecule) or reactive group.

including beads, multi-well plates and activated slides. Photoaptamers derived from the PhotoSELEXTM process which substitutes a brominated deoxyuridine (BrdU) for the thymidine base (T) normally found in DNA, are high-affinity ssDNA compounds that can function as highly specific capture agents for proteins. The photocross-linking of the BrdU of the photoaptamer to an electron-rich amino acid side chain of the protein adds a second dimension of specificity to the normal protein binding event. Since photoaptamers can be covalently bound to their target analytes before signal detection, such arrays can be vigorously washed to remove non-specifically bound proteins, yielding superior signal-to-noise ratios [74]. The Versalinx Protein Microarray Technology by Prolinx utilizes a synthetic chemical affinity (Scheme 11) based on the reversible complexation of phenyldiboronic acid (PDBA) with salicylhydroxamic acid (SHA). The microarray surface is then coated with 3-D polymer brushes which have SHA as their terminal units [75]. This surface permits the reversible binding of PDBA-conjugated proteins. 6.3 Tagged Proteins

To prevent protein denaturation, it is preferable to design surface modifications that target protein functional groups which are furthest from their CDRs. It is important that the epitope mapping domains of the proteins are not affected or modified during the surface immobilization step. By understanding the chemical nature of the proteins of interest, smart surfaces can be constructed for use in highquality protein expression studies. Several protein or antibody samples currently exist in conjugated forms having so-called tags such as biotin, His-Tag, Strep-Tag, Avi-Tag, etc. Tags (Table 5) enhance the efficiency of the surface-coupling step but at the same time have a tendency to alter the properties of pristine proteins. Most research laboratories have tools that are adapted to these tags and if protein

21

22 Protein Microarrays: Surface Chemistry and Coupling Schemes

Scheme 11 Reversible formation of a complex of PDBA (phenyldiboronic acid)-conjugated proteins with SHA (salicyl hydroxymid acid)terminated polymer brushes.

chips have to be integrated into routine test procedures, then they also have to be compatible with such tagged molecules. The use of biotin–streptavidin technology in generating biocompatible surfaces has been highlighted in a large number of research articles [76]. Streptavidin surfaces can be used for immobilizing a wide range of biotinylated molecules (nucleic acids, proteins, saccharides, etc.) and biotin–streptavidin architectures have been widely employed as sensor supports for studying numerous biomolecular interactions. One of the reasons for the rapid growth in the application of biotin–streptavidin technology is the ease with which it is possible to biotinylate various biomolecules. In addition to conventional procedures, new developments are continuously being established in the area of biotinylation of biomolecules. In this area, it is worth mentioning a new method for labeling proteins with biotin by using the so-called AviTagTM technology [77]. Many commercial suppliers offer both biotinylated microwell plates/microarray slides and biotin-conjugated biomolecules Table 5 Examples of commercially available tagged proteins and their corresponding immo-

bilization systems Protein tag

Manufacturer of tagged proteins or antibodies Immobilization matrix

Biotin

Roche, MorphoSys; Biotinylation kits by Molecular Probes or Avidity IBA Novagen, ClonTech

Strep-Tag His-Tag

Streptavidin (Greiner, Perkin Elmer, Xenopore) Strep-Tactin (IBA) Ni-NTA complex (Greiner, Qiagen, Xenopore)

6 Chemical Immobilization Techniques for Proteins

Scheme 12 (A) Structure of N-(5-amino-1-carboxypentyl)iminodiacetic acid. (B) The quadridentate nitrilotriacetic acid (NTA) ligand forms a complex with four binding sites on the nickel metal which is present in the center. The two remaining binding sites can be coordinated with histidine ligands (L).

for biomolecular screening experiments. Another interesting tag for coupling to streptavidin-related surfaces is the Strep-Tag. The Strep-Tag consists of eight amino acids and has a high specific affinity for Strep-Tactin, a synthetically-engineered streptavidin derivative. Recombinant proteins or enzymes can be labeled with such a Strep-Tag. This technology is currently used for protein purification, although microplates and microarrays coated with Strep-Tactin can also be used as a platform for high-throughput screening assays [78]. A well-known label in protein chemistry is the widely used His-Tag. Proteins with six histidine amino acid units at the C- or N-terminus of the peptide chain can be expressed in E. coli. By analogy to affinity chromatography columns, nickel chelate complexes [79] can be used as immobilization matrices on biochip surfaces for His-tagged proteins. An example of a chelate complex is Ni-NTA (NTA = nitrilotriacetic acid). For this purpose, N-(5-amino-1-carboxypentyl)iminodiacetic acid can be synthesized by methods similar to those reported in the literature [80] or by solid phase peptide synthesis coupled to an activated carboxylic acid monolayer and subsequent treatment with a Ni2+ solution. The two free binding sites of the central nickel metal can be coordinated to two histidine units of the tag (Scheme 12). One of the main advantages of such systems is their re-usability. The immobilized proteins can be removed after the screening experiments by the addition of a stong nickel-complexing agent such as EDTA. Although the binding of a histidine tag to nickel is relatively weak compared to the biotin–streptavidin system, Snyder et al. [17] have obtained relatively good results from their nickel-coated protein chips. There are several commercial suppliers who provide Ni-based products for biomolecular screening. Nickel chelate-coated glass slides are, for example, provided by Greiner [80]. As part of their LiquiChip System, Qiagen also offers NiNTA-coated beads. This bead-based microplate platform can be used for a wide range of protein-based assays for proteomics and drug discovery [81]. Whitesides and others have also been granted a patent that covers the use of alkanethiol SAMs

23

24 Protein Microarrays: Surface Chemistry and Coupling Schemes

Scheme 13 Coupling of aldehyde residues of glycoproteins to hydrazide-terminated monolayers.

on gold surfaces which are terminated with Ni-chelate complexes for the coordination of polyamino acid-tagged biomolecules [82]. 6.4 Site-Specific Immobilization of Antibodies

Antibodies are known to bind reversibly to protein A-coated surfaces. Antibodies of the IgG type can be attached to protein A surfaces for high-throughput immunoassay applications by means of their Fc-domains of the heavy chains. Under such conditions, the Fab-domain (fragment of antigen binding) is oriented away from the surface and is easily available for the antigens in the sample solution. In situations where monoclonal IgG antibodies do not bind to protein A surfaces, protein G can be used for antibody immobilization. A review of various currently used immunoassay strategies for protein interactions has been recently reported by MacBeath [83]. Another covalent-linking approach involves the use of hydrazide-terminated monolayers. These functionalities bind exclusively to aldehyde groups that appear only in the glycosylated part of the heavy chains of antibodies, but not in the protein chains of the Fab-domains (Scheme 13).

7 Conclusions

This article has reviewed the numerous strategies which are currently available for immobilizing proteins and related biomolecule families. Since proteins are not as homogeneous as nucleic acids, a wide range of challenges will continue

References

to arise depending upon the nature of the end application. Protein denaturation will continue to be the rate-limiting step in most protein microarray platforms. However, selection of smart chemistries coupled with a sound knowledge of the structural and chemical properties of the probes (which are to be immobilized) will ultimately result in the development of reliable strategies which can be used in the near future to develop functional screening platforms for biomolecules.

References 1 [x1] A. ULMAN, Chem. Rev. 1996, 96, 1533–1554. 2 N. ZAMMATTEO, L. JEANMART, S. HAMELS, S. COURTOIS, P. LHEVESI, J. REMACLE, Anal. Biochem. 2000, 280, 143–150. 3 M. C. PIRRUNG, Angew. Chem. Int. Ed. 2002, 41, 1276–1289. 4 A. ULMANN, Chem. Rev. 1996, 1533–1554. 5 J. GUN, J. SAGIV, J. Colloid. Interface Sci. 1986, 112, 457. 6 K. BIERBAUM, M. GRUNZE, Adhes. Soc. 1994, 213. 7 R. COHEN, N. ZENOU, D. CAHEN, S. YITZCHAIK, Chem. Phys. Lett. 1997, 279, 270–274. 8 B. B. HAAB, M. J. DUNHAM, P. O. BROWN, Genome Biol. 2001, 2, 4.1–13. 9 G. MACBEATH, S. L. SCHREIBER, Science 2000, 289, 1760–1763. 10 T. O. JOOS, M. SCHRENK, P. H¨OPFL, K. KRO¨ GER, U. CHOWDHURY, D. STOLL, D. SCHO¨ RNER, M. D¨URR, K. HERRICK, S. RUPP, K. SOHN, H. H¨AMMERLE Electrophoresis 2000, 21, 2641–2650. 11 L. G. MENDOZA, P. MCQUARY, A. MONGAN, R. GANGHADHARAN, S. BRIGNAC, M. EGGERS, BioTechniques 1999, 27, 778–788. 12 B. D. MARTIN, B. P. GABER, C. H. PATTERSON, D. C. TURNER, Langmuir, 1998, 14, 3971–3975. 13 ] A. BERNARD. E. DELAMARCHE, H. SCHMID, B. MICHEL, H. R. BOSSHARD, H. BIEBUYCK, Langmuir 1998, 14, 2225–2229. 14 S. K. BHATIA, J. L. TEIXEIRA, M. ANDERSON, L. C. SHRIVER-LAKE, J. M. CALVERT, J. H. GEORGER, J. J. HICKMAN, C. S. DULCEY, P. E. SCHOEN, F. S. LIGLER, Anal. Biochem. 1993, 208, 197–205. 15 A. SREEKUMAR, M. K. NYATI, S. VARAMBALLY, T. R. BARRETTE, D. GHOSH, T. S. LAWRENCE, A. M. CHINNAIYAN, Cancer Res. 2001, 61, 7585–7593.

16 B. SCHWEITZER, S. ROBERTS, B. GRIMWADE, W. SHAO, M. WANG, Q. FU, Q. SHU, I. LAROCHE, Z. ZHOU, V. T. TCHERNEV, J. CHRISTIANSEN, M. VELLECA, S. F. KINGSMORE, Nature Biotechnol. 2002, 20, 359–365. 17 H. ZHU, M. BILGIN, R. BANGHAM, D. HALL, A. CASAMAYOR, P. BERTONE, N. LAN, R. JANSEN, S. BIDLINGMAIER, T. HOUFEK, T. MITCHELL, P. MILLER, R. A. DEAN, M. GERSTEIN, M. SNYDER, Science 2001, 293, 2101–2105. 18 L. BERTILSSON, B. LIEDBERG, Langmuir 1993, 9, 141–149. 19 a) R. G. NUZZO, D. L. ALLARA, J. Am. Chem. Soc. 1983, 105, 4481–4483. b) M. D. PORTER, T. B. BRIGHT, D. L. ALLARA, C. D. CHIDSEY, J. Am. Chem. Soc. 1987, 109, 3559–3568. 20 a) G. M. WHITESIDES, C. D. BAIN, Science 1988, 240, 62–63. b) L. STRONG, G. M. WHITESIDES, Langmuir 1988, 4, 546–558. 21 a) L. H. DUBOIS, B. R. ZEGARSKI, R. G. NUZZO, Proc. Natl. Acad. Sci. USA 1987, 84, 4739–4742. b) C. D. BAIN, G. M. WHITESIDES, J. Am. Chem. Soc. 1988, 110, 3665–3666. 22 K. L. PRIME, G. M. WHITESIDES, Science 1991, 252, 1164–1167. 23 Y. Y. LUK, M. KATO, M. MRKSICH, Langmuir 200, 16, 9604–9608. 24 E. OSTUNI, R. G. CHAPMAN, R. E. HOLMLIN, S. TAKAYAMA, G. M. WHITESIDES, Langmuir, 2001, 17, 5605–5620. 25 W. KNOLL, M. LILEY, D. PISCEVIC, J. J. SPINKE, M. J. TARLOV, Adv. Biophys. 1997, 34, 1–15. 26 a) L. S. JUNG, K. E. NELSON, C. T. CAMPBELL, P. S. STAYTON, S. S. YEE, V. PEREZ-LUNA, G. LOPEZ, Sensors Actuators B 1998, 54, 137–144. b) M. SCHA¨ FERLING, M. KRUSCHINA, F. ORTIGAO, M. RIEPL, K.

25

26 Protein Microarrays: Surface Chemistry and Coupling Schemes

27

28 29

30 31

32 33

34

35 36 37

38 39

40 41 42 43 44

ENANDER, B. LIEDBERG, Langmuir 2002, 18, 7016–7023. L. S. JUNG, K. E. NELSON, P. S. STAYTON, C. T. CAMPBELL, Langmuir 2000, 16, 9421–9432. M. MECKLENBURG, B. DANIELSSON, F. WINQVIST, Patent SE, PCT/EP 97/03317. P. PAVLICKOVA, N. MEJLHEDE JENSEN, H. PAUL, M. SCHA¨ FERLING, C. GIAMMASI, M. KRUSCHINA, W.-D. DU, M. THEISEN, M. IBBA, F. ORTIGAO, D. KAMBHAMPATI, J. Prot. Res. 2002, 1, 227–231. J. C. GARNO, N. A. WADU-MESTHRIGE, G.-Y. LIU, Langmuir 2002, 18, 8186–8192. E. DELAMARCHE, G. SUNDARABABU, H. BIEBUYCK, B. MICHEL, C. GERBER, H. SIGRIST, H. WOLF, H. RINGSDORF, N. XANTHOPOULOS, H. J. MATHIEU, Langmuir 1996, 12, 1997–2006. M. VEISEH, M. H. ZAREIE, M. ZHANG, Langmuir 2002, 18, 6671–6678. F. MORHARD, J. PIPPER, R. DAHINT, M. GRUNZE, Sensors Actuators B 2000, 70, 232–242. G. J. LEGGETT, C. J. ROBERTS, P. M. WILLIAMS, M. C. DAVIES, D. E. JACKSON, S. J. B. TENDLER, Langmuir 1993, 9, 2356–2362. I. WILLNER, E. KATZ, Angew. Chem. Int. Ed. 2000, 39, 1180–1218. W. SCHUHMANN, Mikrochim. Acta 1995, 121, 1–29. a) T. TATSUMA, T. WATANABE, Anal. Chem. 1991, 63, 1580–1585. b) M. MCRIPLEY, R. A. LINSENMEIER, J. Electroanal.Chem. 1996, 414, 235–246. A. RIKLIN, I. WILLNER, Anal. Chem. 1995, 67, 4118–4126. a) S. ANDREESCU, L. BARTHELMBS, J.-L. MARTY, Anal. Chim. Acta 2002, 464, 171–180. b) G. JEANTY, A. WOJCIECHOWSKA, J.-L. MARTY, M. TROJANOWICZ, Anal. Bioanal. Chem. 2002, 373, 691–695. J. GERSHONI, G. PALADE, Anal. Biochem. 1982, 124, 396–405. H. TOWBIN, T. STAEHELIN, J. GORDIN, Proc. Natl. Acad. Sci., USA 1979, 76, 4350–4354. B. JOHNSSON, S. LOFAS, G. LINDQUIST, Anal. Biochem. 1991, 198, 268–277. http://www.biacore.com R. BENTERS, C. M. NIEMEYER, D. W¨OHRLE, ChemBioChem 2001, 2, 686–694.

45 B. de BOER, H. K. SIMON, M. P. L. WERTS, E. W. van der VEGTE, G. HADZIIOANNOU, Macromolecules 2000, 33, 349–356. b) C. PREININGER, H. CLAUSEN-SCHAUMANN, A. AHLUWALIA, D. DEROSSI, Talanta 2000, 52, 921–930. 46 a) M. MENNIG, P. W. OLIVERA, H. SCHMIDT, Thin Solid Films 1999, 351, 99–102. b) W. E. VARGAS, J. Appl. Phys. 2000, 88, 4079–4084. c) W. QUE, Z. SUN, Y. L. CHAN, C. H. KAM, Thin Solid Films 2000, 359, 177–183. 47 P. ANGENENDT, J. GLO¨ KLER, D. MURPHY, H. LEHRACH, D. J. CAHILL, Anal. Biochem. 2002, 309, 253–260. 48 http://www.apbiotec.com 49 a) R. P. HUANG, R. HUANG, Y. FAN, Y. LIN, Anal. Biochem. 2001, 294, 55–62. b) R. WIESE, Y. BELOSLUDTSEV, T. POWDRILL, P. THOMPSON, M. HOGAN, Clin. Chem. 2001, 47, 1451–1457. c) S. W. TAM, R. WIESE, S. LEE, J. GILMORE, K. D. KUMBLE, J. Immunol. Methods 2002, 261, 157–165. 50 D. FIGEYS, Proteomics, 2002, 2, 373–382. 51 http://www.calipertech.com 52 http://www.nanogen.com 53 M. PAWLAK, E. SCHICK, M. A. BOPP, M. J. SCHNEIDER, P. OROSZLAN, M. EHRAT, Proteomics, 2002, 2, 383–393. 54 V. BENOIT, A. STEEL, M. TORRES, Y.-Y. YU, H. YANG, J. COOPER, Anal. Chem. 2001, 73, 2412–2420. 55 http://www.combimatrix.com 56 http://.www.zyomyx.com 57 S. A. SOPER, A. C. HENRY, B. VAIDYA, M. GALLOWAY, M. WABUYELE, R. L. MCCARLEY, Anal. Chim. Acta 2002, 470, 87–99. 58 a) D. FIGEYS, S. P. GYGI, G. MCKINNON, R. AEBERSOLD, Anal. Chem. 1998, 70, 3728–3734. b) C.-H. CHIOU, G.-B. LEE, H.-T. HSU, P.-W. CHEN, P.-C. LIAO, Sensors and Actuators B 2002, 86, 280–286. 59 a) H. BECKER, M. ARUNDELL, A. HARNISCH, D. H¨ULSENBERG, Sensors and Actuators B 2002, 86, 271–279. b) D. FIGEYS, Y. ZHANG, R. AEBERSOLD, Electrophoresis 1998, 19, 2338–2347. 60 P. SWANSON, R. GELBART, E. ATLAS, L. YANG, T. GROGAN, W. F. BUTLER, D. E. ACKLEY, E. SHELDON, Sensors Actuators B 2000, 64, 22–30. 61 C. J. YU, Y. WAN, H. YOWANTO, J. LI, C. TAO, M. D. JAMES, C. L. TAN, G. F.

References

62

63

64 65

66

67 68 69

BLACKBURN, T. J. MEADE J. Am. Chem. Soc. 2001, 123, 11155–11161. E. M. BOON, D. M. CERES, T. G. DRUMMOND, M. G. HILL, J. K. BARTON, Nature Biotech. 2000, 18, 1096– 1100. a) M. DIJKSMA, B. KAMP, J. C. HOOGLVLIET, W. P. BENNEKOM, Anal. Chem. 2001, 73, 901–907. b) M. RIEPL, V. M. MIRSKY, I. NOVOTNY, V. TVAROZEK, V. REHACEK, O. S. WOLFBEIS, Anal. Chim. Acta 1999, 392, 77–84. c) C. BERGGREN, B. BJARNASON, G. JOHANSSON, Biosens. Bioelectron. 1998, 13, 1061–1068. http://www.bionchip.com a) F. PATOLSKY, E. KATZ, A. BARDEA, I. WILLNER, Langmuir 1999, 15, 3703–3706. b) C. RUAN, L. YANG, Y. LI, Anal. Chem. 2002, 74, 4814–4820. C. N. CAMPBELL, D. GAL, N. CRISTLER, C. BANDITRAT, A. HELLER, Anal. Chem. 2002, 74, 158–162. http://www.infineon.com A. G. FRUTOS, J. M. BROCKMAN and R. M. CORN, Langmuir 2000, 16, 2192–2197. L. A. RUIZ-TAYLOR, T. L. MARTIN, F. G. ZAUGG, K. WITTE, P. INDERMUHLE, S. NOCK, and P. WAGNER Proc. Natl Acad. Sci. USA. 2001, 98, 852–857.

70 http://www.piercenet.com 71 a) I. WILLNER, A. RIKLIN, B. SHOHAM, D. RIVENZON, E. KATZ, Adv. Mater. 1993, 5, 912–915. b) K. LINDROOS, U. LILJEDAHL, M. RAITIO, A.-C. SYVa¨nen, Nucl. Acids Res. 2001, 29, e69. 72 a) J. RINKE, M. MEINKE, R. BRIMACOMBE, G. FINK, W. ROMMEL, H. FASOLD, J. Mol. Biol. 1980, 137, 301–304. b) J. CIESIOLKA, P. GORNICKI, J. OFENGAND, Biochemistry 1985, 24, 4931–4938. 73 http://www.exiqon.com 74 http://www.somalogic.com 75 http://www.prolinx.com 76 M. Sch¨aferling, S. SCHILLER, H. PAUL, M. KRUSCHINA, P. PAVLICKOVA, M. MEERKAMP, C. GIAMMASI, D. KAMBHAMPATI, Electrophoresis 2002, 23, 3097–3105. 77 http://www.avidity.com 78 http://www.iba-go.de 79 E. HOCHULI, H. D¨OBELI, A. SCHACHER, J. Chromat. 1987, 411, 177–184. 80 http://www.greinerbioone.com 81 http://qiagen.com 82 C. C. BAMDEN, G. B. SIGAL, J. L. STROMINGER, G. M. WHITESIDES, USP 6,197,515, March 2001. 83 G. MACBEATH, Nature Genet. 2002, 32, 526–532.

27

1

Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays Greta J. Wegner, Hye Jin Lee, and Robert M. Corn University of Wisconsin, Madison, USA

Originally published in: Protein Microarray Technology. Edited by Dev Kambhampati. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30597-1

1 Introduction

Multi-component arrays of immobilized biomolecules are rapidly becoming essential tools for the screening of bioaffinity interactions. Biomolecular array technology is especially amenable to proteomics research, since much of a protein’s function is accomplished through specific interactions with DNA, peptides, and other proteins. Array-based assay formats have been developed both for the detection of proteins in a sample [1], and for the identification of new protein–protein interactions [2, 3]. Arrays are also now regularly employed for fundamental studies of protein binding interactions. For example, peptide arrays have been used to study the binding interactions of antibodies [4, 5] and to characterize the sequence-specific activity of enzymes [6, 7]. Arrays of proteins have also been used for the high-throughput screening of protein–protein interactions and to identify novel biochemical activities [2, 3]. The majority of array-based assays currently employ fluorescent, enzymatic or radiolabeled biomolecules. Further advancement of analytical systems that do not require labeled biological molecules to detect affinity binding interactions is needed. Surface plasmon resonance (SPR) imaging is emerging as a label-free surface sensitive optical technique that measures the adsorption of solution phase molecules to arrays of biomolecules on thin gold films by changes in the local index of refraction. Label-free detection is particularly advantageous for the assay of small molecule-binding to proteins, since labels can interfere with the biological activity of the target molecule. To date, SPR imaging has been used to study reversible protein binding to DNA, peptide, protein, and carbohydrate arrays [8–10]. This review summarizes the recent advances that we have made in the implementation of SPR imaging to study DNA, RNA, and protein interactions with Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

DNA, peptide and protein probes immobilized in an array format. After a brief description of the SPR imaging technique in Section 2, two surface attachment chemistries used for the covalent immobilization of thiol-modified DNA, peptides, and carbohydrates through the modification of self-assembled monolayers of alkanethiols are discussed in Section 3. In Section 4, an array fabrication method based on chemical protection and photopatterning is described. Several examples of SPR imaging experiments using photopatterned arrays are presented including the study of DNA–DNA hybridization and protein–DNA binding interactions. We will conclude this article in Section 5 with a second array fabrication method based on microfluidic stenciling and address its application to peptide–protein and protein–protein binding interactions.

2 Surface Plasmon Resonance Imaging

Surface plasmon resonance (SPR) measurements are surface-sensitive optical methods used to characterize organic and biological layers on gold or noble metal thin films. These measurements utilize the optical field enhancement that occurs at a metal/dielectric interface when surface plasmons are created. Surface plasmons are electromagnetic waves, excited by p-polarized light, that propagate parallel to the gold surface. The optical fields decay exponentially from the surface of the metal and have a maximum decay length of about 200 nm. Within this region, the optical fields are sensitive to changes in the index of refraction caused by the adsorption or desorption of molecules onto the surface. As a consequence, SPR experiments have been amenable for the label-free study of reversible biomolecule binding interactions on gold films in real-time using two different methods: (1) scanning angle SPR and (2) SPR imaging techniques. Angle shift measurements are the most commonly undertaken SPR technique and have been popularized by the commercial Biacore instrument. In a scanning angle experiment, the percentage of light reflected off of a thin gold film optically coupled to a prism is measured as a function of incident angle. A prism or grating coupling arrangement is required because surface plasmons cannot be excited directly at planar air/metal or water/metal interfaces because momentum-matching conditions are not satisfied. A hemispherical prism is used to tightly focus light from a laser onto a single region of a gold thin film. Figure 1 shows three SPR curves and their theoretical fits where the percent reflectivity of light is measured as a function of the incident angle for light with excitation wavelengths of 1152 (), 814 (), and 633 nm (◦) [11]. For the SPR curve measured at an excitation wavelength of 814 nm, increasing the incident angle from 50 to 50.8◦ causes nearly 100% of the light to be reflected as the critical angle is reached and total internal reflection occurs. Further increase in the angle results in a steep drop in percent reflectivity as light is incorporated into surface plasmons until a minimum is reached at 53.5◦ , the surface plasmon angle. The position of the SPR angle is sensitive to changes in the index of refraction and/or thickness of the adsorbed film causing

2 Surface Plasmon Resonance Imaging

Fig. 1 SPR reflectivity curves showing the relationship between percentage reflectivity and angle of incidence for excitation wavelengths of 1152 nm (), 814 nm (), and 633 nm (◦) for an SF-10 glass/Au/water (in situ) assembly. Solid lines show the theoretical fit obtained by three-phase Fresnel calculations. Sharper SPR curves are obtained with increasing wavelength and can be applied to study thicker films with angle shift measurements and to enhance the contrast of SPR imaging experiments. The angle at which SPR imaging measurements are ob-

tained is marked by a vertical line on the SPR reflectivity curve obtained at an excitation wavelength of 814 nm. Inset shows the relationship between the change in percentage reflectivity (%R) and the change in index of refraction (nf ) at a fixed incident angle of 56.33◦ . A linear relationship is observed for %R less than 10%. Reprinted

with permission from Analytical Chemistry, 71 3928–3934, copyright 1999 American Chemical Society and Analytical Chemistry, 73 1–7, copyright 2001 American Chemical Society.

the minimum to shift. Shifts in angle position can be correlated to changes in thickness or index of refraction using N-phase Fresnel calculations as described by Hansen [12]. SPR measurements can be enhanced by using nearinfrared excitation wavelengths between 800 and 1100 nm. Mid-IR light results in sharper SPR curves than those obtained at an excitation wavelength of 633 nm from a HeNe source (refer to Figure 1). This allows a more precise determination of the SPR angle and can be used to measure thicker films. While scanning angle experiments are used to study a single region on a gold surface, SPR imaging is used to monitor the spatially resolved adsorption of biomolecules onto a multicomponent array. This technique is especially important for biological experiments where the simultaneous measurement of multiple interactions on a single chip is desired. SPR imaging measurements of the change in percent reflectivity are performed at a fixed angle near the SPR angle, as shown by a vertical line passing through the SPR reflectivity curve measured using an excitation wavelength of 814 nm as shown in Figure 1. Figure 2 shows a schematic representation of the SPR imaging instrument used to detect the adsorption of biopolymers in solution to surface-immobilized biomolecules such as DNA and peptides [11]. A collimated white light source is coupled with a narrow band pass

3

4 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

Fig. 2 Schematic diagram of the surface plasmon imager apparatus. P-polarized white light impinges on a prism sample apparatus at a fixed optimal angle, near the surface plasmon angle. The light is then passed through a narrow band pass filter and images are collected by a CCD camera. The analyte is delivered to the array fabricated on a gold thin film using a simple

500 :L flow cell. An SPR difference image of the sequence-specific hybridization of 18mer DNA to a two-component DNA array is shown in the bottom left corner of the figure. A maximum of 160 array elements can be studied on one SPR chip with a total surface area of 0.8 cm2 using square elements with 500 :m widths.

filter centered at 830 nm and used in place of a laser source. By using this configuration, laser fringes which can interfere with image quality are avoided and the enhanced contrast obtained using an NIR source is maintained. Images with lateral resolutions on the order of 50 µm can be obtained using an excitation wavelength of 830 nm. The light is passed through a polarizer in a direction incident to a high index glass prism/sample assembly. Here, light impinges on the back of a sample consisting of a gold thin film chemically modified with an array of biomolecules. Analytes are delivered to the array by either a 500-µL flow cell or through a set of parallel microfluidic channels from PDMS requiring 1 µL of sample per channel. Images are collected using a CCD camera after the light reflected off the sample is passed through a narrow band pass filter. The light interacts with the biomolecules immobilized on the thin gold film to create surface plasmons, inducing attenuation in the light reflected from the surface. As biomolecules from solution adsorb onto the array, the local index of refraction changes causing an increase in the percentage of incident light reflected off each array element of the sample, at a fixed optimal angle. The lower left corner of Figure 2 contains a typical SPR difference image showing sequence-specific adsorption of complementary DNA onto a two-component DNA array. Like the scanning angle SPR measurements, quantitative information can be obtained from SPR imaging measurements. The inset in Figure 1 shows the relationship between the percentage reflectivity and the index of refraction for changes in percentage reflectivity under 10% [13]. This relationship can also be applied to determine the surface coverage of biomolecules adsorbed onto a surface. SPR imaging measurements have been used for the quantitative evaluation of the interactions between biomolecules in solution and DNA or peptide probes immobilized

3 Surface Attachment Chemistries

on the array. In the case of biomolecules that have a high molecular weight, the number of immobilized surface target sites can be decreased in order to stay within this linear %R region [10]. The sensitivity of the SPR imaging approach is about 10 femtomoles (1 µL of a 10 nM solution) for 18-mer single-stranded DNA hybridized to a DNA array [14], and 1 femtomole (1 µL of a 1 nM solution) for specific antibody adsorption onto a peptide array [10]. Although SPR imaging measurements do not match the sensitivity of fluorescence or radioactive methods, it is more than adequate for many applications including the study of protein–protein interactions and has the advantage of employing direct detection of binding interactions.

3 Surface Attachment Chemistries

The development of well-characterized surface chemistries for the attachment of biological molecules onto gold thin films in an array format is necessary for SPR imaging measurements of biomolecular affinity interactions. Noble metal thin films are required for the propagation of surface plasmons. As a consequence, commercially available DNA arrays made on glass substrates, such as those produced by Affymetrix [15–18], or peptide arrays prepared by the SPOT synthesis technique on cellulose membranes [19–21], are not viable options for SPR imaging investigations. Instead, we have employed self-assembled monolayers of long chain alkanethiols that are ω-terminated with an amine functional group as the foundation of the array [8, 9, 11, 22]. Chemical modification of the self-assembled monolayers is used to tether biological molecules to the surface. In this section, two methods using the bifunctional linkers SSMCC (sulfosuccinimidyl 4-(N-maleimidomethyl) cyclohexane-1-carboxylate) and SATP (N-Succinimidyl S-acetyl-thiopropionate) are described for the covalent attachment of thiol-containing DNA, peptide, carbohydrate and capture probe molecules. 3.1 SSMCC Attachment Chemistry

In the first reaction scheme, the heterobifunctional linker SSMCC, which contains both an N-hydroxysulfosuccinimide (NHSS) ester and a maleimide functionality, is used to covalently link the amine-terminated self-assembled monolayer of 11mercaptoundecylamine (MUAM) to a thiol-modified probe molecule (Figure 3a) [8, 22]. First, the NHSS ester moiety of SSMCC is reacted with the free amines of the alkanethiol monolayer to form amide bonds terminating with maleimide groups. Next, thiol-modified DNA or cysteine-containing peptides are introduced to the thiol-reactive maleimide monolayer and attached by the creation of a thioether bond. DNA molecules attached by this method have a surface coverage of 1.0 × 1012 molecules cm−2 [13].

5

6 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

Fig. 3 Surface attachment chemistry for the immobilization of thiol-modified DNA and cysteine-containing peptides: (a) The linker SSMCC is reacted with a wellpacked self-assembled monolayer of 11mercaptoundecylamine (MUAM) to create a maleimide-modified surface. The maleimide surface is then used to covalently attach thiol-modified DNA or cysteine-containing

peptides. (b) In the second approach, SATP is reacted with MUAM to create a protected thiol surface. Upon deprotection with a basic solution containing DTT, the free sulfhydryl is reacted with dipyridyl disulfide to create a pyridyl disulfide surface. Thioldisulfide exchange reactions are used to couple thiol-containing biomolecules to the surface.

3.2 SATP Attachment Chemistry

An alternative reaction scheme, more amenable for generating peptide arrays, involves the attachment of thiol-containing biomolecules onto gold surfaces through a disulfide linkage [10, 23]. The reaction scheme for this surface modification process is shown in Figure 3b. In this approach, a self-assembled monolayer of MUAM is reacted with the molecule SATP, a bifunctional linker containing an NHS ester and a protected sulfhydryl. The NHS ester reacts with the amines present on the surface to form a stable amide bond resulting in a protected sulfhydryl surface. Exposure of the surface to an alkaline solution containing DTT removes the protecting acetyl group revealing an active sulfhydryl surface. Next, the sulfhydryl surface is reacted with 2,2 -dipyridyl disulfide to form a pyridyl disulfide surface. Thiol–disulfide exchange reactions are then performed in order to attach thiol-modified DNA or cysteine-containing peptides to the substrate, with a surface coverage of 1013 molecules cm−2 . This type of surface attachment chemistry has the advantage of being reversible, as the disulfide bond can be cleaved in the presence of DTT to regenerate the sulfhydryl-terminated surface.

4 SPR Imaging Experiments Using Photopatterned Arrays

4 SPR Imaging Experiments Using Photopatterned Arrays

Arrays used for SPR imaging experiments contain multiple individually addressable components and are prepared by a combination of self-assembly, surface attachment chemistry, and array patterning using either photopatterning or PDMS microfluidic channels. This section will explore the first array fabrication methodology developed within our laboratories for the creation of robust DNA and peptide arrays using chemical protection and deprotection and photopatterning. This multistep fabrication procedure is outlined in Figure 4 [8]. The first step of the fabrication method is the creation of a temporary hydrophobic background using the bulky amine protecting group, 9-fluorenylmethoxycarbonyl (Fmoc), frequently used in solid phase peptide synthesis. The N-hydroxysuccinimide ester of Fmoc (FmocNHS) is reacted with the terminal amines of a packed-self assembled alkanethiol

Fig. 4 Multi-step array fabrication process using protection/deprotection chemistry and photopatterning techniques to prepare DNA and peptide arrays. The bottom left corner of the figure shows individually addressable hydrophilic drops of DNA pinned to the surface by a hydrophobic background. First, a bare gold surface is modified with a self-assembled monolayer of 11-mercaptoundecylamine (MUAM). This amine-terminated surface is reacted with the hydrophobic protecting group Fmoc.

UV-light is used to break the gold–thiol bond and create bare gold pads on the surface. These pads are subsequently filled with MUAM. A bifunctional linker is used to attach DNA or peptide probes to create an array. Finally, the Fmoc is removed with base and replaced with a polyethylene derivative to prevent the non-specific adsorption of biopolymers to the background. Reprinted

with permission from Journal of the American Chemical Society, 121, 8044–8051. Copyright 1999 American Chemical Society.

7

8 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

monolayer to form a stable carbamate linkage. The surface is then exposed to UV light through a patterned quartz mask to create arrays composed of 500 by 500 µm square elements with 500 µm spacing between the elements. The gold–thiol bond is cleaved in the regions where the light shines through the mask resulting in a patterned gold surface that consists of bare gold pads surrounded by Fmoc. Next, the surface is immersed in an ethanolic solution of MUAM. The amine-terminated alkanethiols self-assemble in the bare gold pads producing a surface containing reactive hydrophilic MUAM pads surrounded by the hydrophobic Fmoc background. The lower left-hand corner of Figure 4 contains an image of an array showing the individually addressable hydrophilic spots surrounded by a hydrophobic background. Chemical protection allows droplets of hydrophilic DNA or peptides to be “pinned”, without contaminating other sequences contained on neighboring pads on the array, and covalently attached using the linkers SSMCC or SATP. These arrays can be easily used with a mechanical array spotter to introduce multiple peptide or DNA elements onto one chip for high-throughput studies. For example, arrays composed of more than 300 elements can be produced using square elements with 250-µm widths. The array fabrication is then completed by removing the hydrophobic Fmoc background and replacing it with polyethylene glycol (PEG) to reduce the nonspecific adsorption of biopolymers. Since Fmoc is a base-labile protecting group, the original amine-terminated surface can be easily regenerated by exposure to a secondary amine. An NHS derivative of polyethylene glycol (PEG) can react with the terminal amines of the SAM to form a stable amide linkage. The formation of the temporary Fmoc surface is necessary, since PEG is too hydrophilic to prevent contamination between DNA or peptide droplets. The DNA arrays can then be analyzed by SPR imaging using a 500 µL flow cell to introduce biopolymer analytes to the array surface. DNA arrays fabricated using SSMCC attachment chemistry can be used for more than 25 hybridization cycles [13] while the SATP chemistry can be used for more than 30 cycles without degradation of the array [23]. 4.1 DNA–DNA Hybridization

The hybridization of complementary DNA to a five-component DNA array fabricated by photopatterning and SSMCC chemistry was examined by SPR imaging. All surface-immobilized DNA probes were tested by hybridization to respective perfect-match DNA complements. The probes were immobilized in a distinct geometric pattern to easily identify interacting sequences. The DNA sequences used in this experiment are listed in Table 1, and denoted as A–E. The surface was first exposed to a 100 nM solution of the 16-mer DNA oligonucleotide complement to probe A for 15 min (Figure 5a). The SPR difference image shown in this figure was produced by subtracting the images taken before and after hybridization. Regeneration of the surface was achieved by rinsing with 8 M urea. Repeated cycles of hybridization and denaturation were used to obtain Figure 5b–e.

4 SPR Imaging Experiments Using Photopatterned Arrays Table 1 DNA sequences used in Figure 5

Surface-bound probe DNA

DNA sequences (5 −3 )

Probe A Probe B Probe C Probe D Probe E

TAC TCA CCC GTC CGC C CTT TTA TGT TTG AAC CAT GCG GTG GCC GAT CAC CCT CTC CGT GGC TTT CTG GTT A CTT TGA GTT TCA GTC

Probe DNA is modified with a 5 -thiol modifier and a 15-T spacer is appended to the 5 end of the probe DNA.

Excellent sequence-specific adsorption of the complementary DNA was observed for all probes and with little non-specific adsorption to the background. The approximately equal SPR signal under identical experimental conditions indicates that the probes are equally accessible for hybridization to target molecules in solution. 4.2 Mismatch Binding Protein, MutS

Mismatch binding proteins, such as MutS in E. coli are used by cells to identify and repair mismatches, short insertions, and deletions that are caused by replication

Fig. 5 SPR difference images showing the hybridization of perfect-match DNA complements to a DNA array containing five different probes. The sequences were immobilized on 500 by 500 :m array elements in the pattern shown in the figure. (a) First, the array was exposed to a 100 nM solution of the DNA complement to probe A for 15 min. Hybridization adsorption onto the ar-

ray is indicated by a change in the percentage reflectivity of incident light. (b) After rinsing with 8 M urea to regenerate the surface, the experiment was repeated with the DNA complement of probe B. Successive rounds of denaturation and hybridization of the remaining DNA complement probes resulted in images c–e. The same region of the array is shown for all images.

9

10 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

errors. The ability to identify mismatches in DNA is necessary for the technology used to detect mutations and single nucleotide polymorphism in double-stranded DNA. Such assays may be important in the future for correlating genomic information from an individual with the development of diseases [24–26]. As an example, SPR imaging was used to show the interaction of MutS with mismatch DNA sequences (A–D) immobilized onto an array [9, 27]. The sequences and additional experimental information are reported elsewhere [9]. This photopatterned array was exposed to a solution containing the sequence Z. Probe A did not hybridize with sequence Z and remained single stranded. Sequence Z formed a duplex with probe B containing a two-base insertion, a perfectly matched duplex with probe C, and a duplex containing a one-base G/T mismatch with probe D. Figure 6 shows an SPR difference image after MutS was allowed to bind to the DNA array. Adsorption of MutS was observed to both duplex B containing the insertion and duplex D containing the G/T mismatch, which has the strongest mismatch interaction with MutS [28]. Minimal binding occurred to the perfect match or the single-stranded DNA, showing that it was possible to study the sequence-specific interactions of MutS to DNA using SPR imaging measurements of DNA arrays. The utility of mismatch binding proteins to identify single-base mismatches, as well as short insertions and

Fig. 6 SPR imaging measurements of E. coli mismatch binding protein MutS adsorption onto a DNA array. This array was created with DNA probes A through D immobilized on 500 by 500 :m array elements in the pattern shown on the left of the figure. The array was then exposed to a solution containing the sequence Z. Z does not bind at all to probe A leaving it single strandedbut binds to probe B to create a duplex containing a two-base insertion, to probe C in a perfectly complementary manner, and to probe D to form a duplex containing a

single-base mismatch. An SPR difference image of the binding of MutS to the array is shown on the right of the figure. The image shown is the difference between two images collected before and after exposure of the surface to MutS. Only 12 array elements from the total patterned surface area of 0.8 cm2 are presented in these images. Reprinted with permission from An-

nual Review of Physical Chemistry 51, pp 41–63. Copyright 2000 by Annual Reviews www.annualreview.org.

5 SPR Imaging Experiments of Arrays Created by Microfluidic Stencils 11

deletions in double-stranded DNA will lead to the application of mismatch binding proteins in mutation and single nucleotide polymorphism detection assays. 4.3 Bacterial Response Regulators, OmpR and VanR

Response regulators are DNA binding proteins that are essential components of the two-component signal transduction phosphorylation relay system used by bacteria to respond to environmental stimuli and to adapt accordingly through transcriptional activation of certain genes [29]. Since few response regulators have been studied in depth, DNA arrays are a potential tool to determine the DNA sequences recognized by a particular response regulator. As an example, the sequence-specific binding of two response regulators, OmpR and VanR, to DNA probe molecules was studied [30]. OmpR is one of the most well-characterized response regulators and is important in bacterial response to environmental osmotic pressure [31], while VanR is involved in the development of antibiotic resistance [32]. A photopatterned array was prepared containing six different immobilized DNA probes known to interact with either OmpR or VanR. The DNA array was then exposed to solutions of corresponding complementary DNA to create surfaceimmobilized double-stranded DNA. Figure 7a shows an SPR difference image after the array was exposed to a 100 nM solution of OmpR. OmpR primarily absorbed to the DNA sequences OmpF1, F1F2, and C2 which were known to interact only with OmpR. The array was then regenerated with urea and exposed to complementary DNA before VanR (500 nM) was introduced to the array. VanR primarily bound to the DNA sequences VanH1, H2, and R1 (Figure 7b). Little nonspecific adsorption of either VanR or OmpR to the other sequences on the chip was observed. DNA probe sequences for VanR and OmpR response regulators can be found in the literature [30]. In addition to sequence specificity, SPR imaging measurements of DNA arrays can also be used to monitor inhibitor reactions that obstruct the binding of response regulators to DNA controlling gene expression in the bacteria. Figure 8 shows the effect of the inhibitor CpdA on VanR binding to the DNA sequences VanH1, H2, and R1 as measured by SPR imaging. The changes in percentage reflectivity were measured as a function of VanR concentration with and without 5 µM of CpdA. Integration of the line profile was performed to determine the average change in percentage reflectivity for the DNA sequences VanH1, H2, and R1. As can be seen in Figure 8, more adsorption is observed in the absence of the inhibitor than when CpdA is included with the VanR solution.

5 SPR Imaging Experiments of Arrays Created by Microfluidic Stencils

We have recently developed a second fabrication and detection approach based on the coupling of microfluidic networks to SPR imaging measurements with

12 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

Fig. 7 Surface plasmon imaging difference images of response regulator adsorption to double-stranded DNA immobilized on a photopatterned array composed of 500 by 500 :m array elements. (A) Specific adsorption of a 100 nM solution of OmpR to the DNA sequences OmpF1, F1F2, and C2. (B) VanR (500 nM) adsorbs to the DNA sequences VanH1, H2, and R1. OmpR is known to bind to the DNA sequences

OmpF1, F1F2, and C2 and VanR is known to bind VanH1, H2, R1. There is little nonspecific adsorption of the response regulators to the other sequences on the chip. DNA probe sequences for VanR and OmpR response regulators can be found in the literature. Reprinted with permission from Langmuir, Vol. 19, 1486–1492. Copyright 2003 American Chemical Society.

Fig. 8 Plot showing the change in percentage reflectivity observed when VanRphosphate (•) and VanR-phosphate plus 5 mM CpdA (◦) adsorb to VanR DNA promoter sequences as a function of protein concentration. Integration of the line profile of VanR binding to DNA sequences VanH1, H2, and R1 was used to determine the av-

erage percentage reflectivity at each protein concentration. Less VanR-phosphate adsorbs to the DNA probes in the presence of the inhibitor CpdA than in its absence.

Reprinted with permission from Langmuir, Vol. 19, 1486–1492. Copyright 2003 American Chemical Society.

5 SPR Imaging Experiments of Arrays Created by Microfluidic Stencils 13

the specific aim of lowering detection limits, reducing analysis time, and decreasing chemical consumption and sample volume [14]. This approach relies on the construction of microfluidic networks within the polymer polydimethylsiloxane (PDMS), which are physically sealed to chemically-modified gold surfaces. These PDMS microchannels were used in two ways: (i) to fabricate “1-D” arrays consisting of lines of immobilized peptides and DNA probes and (ii) to create “2-D” detection arrays in which a second set of PDMS microchannels were placed perpendicular to a 1-D line array in order to deliver target samples with small volume [14]. 5.1 1-D Peptide Array for Antibody Binding Measurements

The fabrication of 1-D line arrays on gold surfaces is as follows. First, a set of parallel microchannels from PDMS were created by replication from 3-D silicon wafer masters that were created photolithographically from a 2-D chrome mask pattern (Figure 9) [33, 34]. The microchannels were physically attached to a gold substrate modified with MUAM. A simple differential pumping system was used to introduce the chemical linkers SATP or SSMCC into the microchannels. Next, the DNA or peptide probe molecules were reacted within the microchannels. Once the immobilization was complete, the channels were removed and the alkanethiolterminated monolayer was treated with PEG-NHS to prevent non-specific adsorption to the surface. Since the PDMS microchannels physically define where the probe molecules are immobilized, protection and deprotection chemistry is not required.

Fig. 9 Schematic representation of the microfabrication strategy used to create DNA, peptide and protein linear arrays. A self-assembled monolayer of MUAM was formed on a clean gold surface. Polydimethyl siloxane (PDMS) microchannels were used to deliver bifunctional linkers and

probe biomolecules to the surface. The microchannels were removed and the background was protected with a polyethylene glycol derivative. Adapted with permission from Analytical Chemistry, 73 5525–5531. Copyright 2001 American Chemical Society.

14 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

Fig. 10 SPR difference images showing the sequence-specific binding interactions of antibodies to a peptide linear array containing two probes A and B corresponding to the peptide sequences Myc and HA. The peptides were immobilized in the pattern shown in the figure. (a) First, the array was

exposed to a 25 nM solution of anti-Myc. Specific adsorption of anti-Myc to probe A is observed. (b) After rinsing with pH 11.5 buffer to regenerate the surface, the experiment was repeated 100 nM anti-HA. AntiHA primarily bound to probe B, with little non-specific interaction with probe A.

By changing the spacing of the channels, up to 100 different peptide or DNA sequences can be immobilized on one chip. The array can then be coupled with a large volume flow cell (500 µL) to introduce analytes to the line array. SPR imaging measurements were used to monitor the sequence-specific adsorption of antibodies to a two-component peptide array. The linear array contained the peptide sequences, Myc and HA denoted A and B, respectively (Figure 10). The introduction of a 25 nM solution of anti-Myc though a 500 µL flow cell resulted in specific adsorption to probe A. The anti-Myc was removed from the surface using pH 11.5 buffer prior to exposing the array to a solution of 100 nM anti-HA. Anti-HA primarily bound to probe B, although some non-specific adsorption probe A was observed. 5.2 2-D DNA Array for RNA Hybridization

In a second approach, microfluidic channels were used as a small volume flow cell during SPR imaging measurements to reduce the required analyte and to simultaneously introduce multiple analytes. First, a 1-D array composed of DNA elements was prepared and the background protected with PEG. Then a second set of PDMS channels was placed perpendicular to the immobilized DNA lines, forming a 2-D array, and used to introduce analytes to the array surface during SPR imaging

5 SPR Imaging Experiments of Arrays Created by Microfluidic Stencils 15

Fig. 11 (a) Schematic diagram of parallel polydimethyl siloxane (PDMS) microchannels used to deliver small volumes of analyte (1 :L) to peptide and DNA linear arrays. (b) Schematic representation of the

SPR imaging experimental set-up incorporating PDMS microfluidics to deliver a small volume of target sample. This configuration allows different analytes to be delivered through each channel.

experiments (Figure 11a). Figure 11b shows the schematic diagram of an SPR imager set-up where the large flow cell (500 µL) has been replaced with microfluidic channels constructed in PDMS reducing the required sample volume to 1 µL. An example of an SPR difference image of a three-component 2-D hybridization DNA array exposed to GUS gene ssRNA through PDMS microchannels is shown in Figure 12. About 20 femtomoles of RNA in a total solution volume of 1 µL was delivered through each channel. The DNA sequences and more information are reported elsewhere [14]. Varying SPR signal intensities were observed for the three different probes in Figure 12 and can be attributed to differences in the binding efficiency of the GUS gene ssRNA to the different surface-bound ssDNA probes. The ability to detect 20 femtomoles of ssRNA is sufficient for a number of biological applications, including the direct detection of messenger RNA (mRNA) from

16 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

Fig. 12 An SPR difference image showing adsorption of GUS gene ssRNA onto DNA probes. Each channel was 300 :m wide, 35 :m deep, and 1.2 :m long with 900 :m spacing between channels. Reprinted with permission from Analytical Chemistry, 73 5525–5531. Copyright 2001 American Chemical Society.

highly expressed genes and the detection of ribsomal RNA (rRNA) from complex biological samples. 5.3 2-D Peptide Array for Antibody Binding Measurements

SPR imaging technology integrating microfluidics was applied to monitor the sequence-specific interactions of antibodies to peptide motifs for epitope mapping applications. For example, peptide arrays were used to study the residues essential to the binding motif of anti-FLAG M2, an antibody commonly used for the purification of fusion proteins [35]. SPR imaging is currently used to study cell adhesion processes and to investigate the enzymatic modifications of peptides by sequence-specific phophatases, kinases, and proteases. Ultimately, these techniques could be used to identify the important residues in previously uncharacterized protein–protein interactions based on peptide recognition motifs. Sequence-specific adsorption of anti-FLAG M2 to four different peptide epitopes based on the FLAG peptide tag was studied with SPR imaging [10]. Peptide line arrays were created using a microfluidic fabrication process and coupled to a second microchannel with a wraparound design (Figure 13a). The “worm” microchannel has a total sample volume of 5 µL and is used to deliver antibody to the peptide array. Figure 13b shows a difference image obtained by subtracting images of a fourcomponent peptide array before and after the introduction of a solution of 100 nM anti-FLAG M2. A line profile taken across the channels indicated that the greatest amount of antibody adsorption is observed at elements where the original sequence, F1, was immobilized. The least amount of binding was observed to the peptide sequences F3 and F4. These peptides differed from the original sequence by the substitution of an amino acid important to the binding motif by

5 SPR Imaging Experiments of Arrays Created by Microfluidic Stencils 17

Fig. 13 (a) A worm microchannel is used to deliver 5 :l aliquots of antibody solution to the peptide linear array. This microfluidic configuration is suitable for the delivery of a small volume of a single analyte to the array with a detection area equivalent to a configuration using a large volume flow cell. The worm channel was 500 :m wide, 35 :m deep, and 12 mm long with 500 :m spacing between channels. (b) SPR difference image showing the adsorption of 100 nM anti-FLAG to a peptide array composed

of four epitopes, differing by a single amino acid, based on the FLAG peptide sequence. The line profile reveals that the greatest adsorption occurs at elements containing the original sequence, F1. Diminished adsorption is observed at sequences, F3 and F4, containing alanine substitutions for essential residues of the binding motif. Peptide probe sequences can be found in the literature. Reprinted with permission from Analytical Chemistry, 74, 5161–5168. Copyright 2002 American Chemical Society.

an alanine residue. In contrast to the F3 and F4 peptides, the F2 sequence had a substantially higher signal, since this peptide contained an alanine substitution for a non-essential residue in the binding motif. These data demonstrate that SPR imaging can be used to compare the amount of protein binding to peptides differing by a single amino acid and to determine the importance of each residue for a peptide–protein interaction.

18 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

Fig. 14 Langmuir isotherms fitted to the adsorption of anti-FLAG M2 onto an array containing the peptides F1 (•) and F2 (◦). The change in percentage reflectivity was determined by integrating the line profiles for the F1 and F2 peptides at varying anti-FLAG M2 concentrations. The adsorp-

tion constants were calculated by the fit of the Langmuir isotherms, to be 1.5 × 108 M−1 and 2.8 × 107 M−1 for F1 and F4 respectively. Reprinted with permission

from Analytical Chemistry, 74, 5161–5168. Copyright 2002 American Chemical Society.

Quantitative SPR imaging measurements can also be used to simultaneously determine adsorption constants for the interactions of antibodies with multiple peptide motifs on a single chip [10]. The change in percentage reflectivity was measured for the peptide sequences F1 and F2 at increasing antibody concentrations. This data is shown in Figure 14 where the SPR signal resulting from antibody adsorption is plotted as a function of anti-FLAG M2 concentration, ranging from 1 to 150 nM. These data points were fitted with a Langmuir isotherm. The adsorption constants were found to be 1.5 × 108 M−1 for F1 and 2.8 × 107 M−1 for F2, showing that the original sequence had a stronger antibody peptide interaction. The detection limit for the F1 peptide with anti-FLAG M2 was 0.5 nM. 5.4 2-D Protein Array

With the sequencing of many genomes complete, it has become clear that highthroughput methods for studying gene products is necessary in order to elucidate the vast amount of information encoding biological function in an organism. Such studies would be enhanced by the addition of label-free analytical tools such as SPR imaging. The utility of SPR imaging for the study of protein–peptide interactions demonstrated the feasibility of this technique for measuring the adsorption of biomolecules to immobilized proteins. The first step for adapting SPR imaging measurements to the study of protein–protein interactions is the creation of a robust reproducible, biologically-active protein array. Most protein arrays are constructed

5 SPR Imaging Experiments of Arrays Created by Microfluidic Stencils 19

by random attachment strategies, using naturally occurring lysines and terminal amines for surface attachment [36]. However, such approaches have the potential to interfere with the binding site of the protein, negatively affecting biological activity. To target such problems, we have established a general methodology to create oriented arrays of fusion proteins attached to gold thin films. In an oriented fusion protein array, the fusion protein or tag will interact with a capture agent immobilized on the gold thin film leaving the protein of interest free to interact with proteins in solution. We are currently investigating several fusion protein/capture agent pairs including the histidine peptide tag with NTA [37–39], glutathione Stransferase (GST) with glutathione or anti-GST [40, 41], and maltose binding protein (MBP) with maltose [42, 43]. Since these tags and capture agents are commonly used for the purification of fusion proteins, they are well-characterized and the proteins can be released to follow up SPR imaging measurements with further off-chip solution phase experiments. Protein arrays fabricated using NTA-modified gold films to capture histagged proteins are a promising tool for the study of protein–protein interactions using SPR imaging (see Figure 15a). A worm microchannel was used to pattern a selfassembled alkanethiol monolayer on a gold thin film with NTA capture agents. This microchannel was then removed and the whole surface was reacted with a polyethylene glycol derivative to prevent the non-specific adsorption of proteins

Fig. 15 (a) Schematic representation of a general approach for the creation of an oriented fusion protein array on gold surfaces. (b) The NTA capture agent was immobilized through a worm microchannel. Next, a parallel set of PDMS channels was used to simultaneously deliver multiple histidinetagged analytes to the NTA elements. (c) An example of an SPR difference image

showing the immobilization of histidinetagged ubiquitin B and polyhistidine D to NTA-modified monolayers in the presence of nickel ions. In the absence of nickel ions there is little non-specific adsorption of histagged ubiquitin A or polyhistidine C to the NTA surface, suggesting that a specific chelation interaction is occurring.

20 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

to the background around the NTA capture agents. Next a second set of parallel microchannels oriented perpendicular to the immobilized NTA lines was used to introduce proteins to the surface (Figure 15b). This fabrication method provides a convenient and fast way to create arrays composed of one protein per channel. As shown in the SPR difference image in Figure 15c, both polyhistidine and histidinetagged ubiquitin interact with the NTA surfaces in the presence of nickel ions, while little adsorption of these biomolecules is observed when nickel ions are not present, showing the specificity of the interaction. This protein fabrication strategy has the advantage that SPR imaging measurements can be used to evaluate protein immobilization onto the array prior to using the chip to study protein–protein interactions. Our current efforts are focused on the application of SPR imaging measurements to study the interactions of protein, DNA, and peptides to protein chips. Initial results indicate the SPR imaging techniques will become an innovative tool for proteomics research. 6 Conclusions

SPR imaging is emerging as a promising tool for the study of bioaffinity interactions in an array format. This review presented various recent applications of SPR imaging measurements for the study of DNA, RNA, and protein interactions with biomolecular arrays. The key components for these experiments are a wellcharacterized chemical modification of gold thin films to immobilize biomolecular probe molecules coupled with a robust array fabrication method based on photopatterning or microfluidics. The development of two different methods of array fabrication has increased the versatility of the SPR imaging technique; the linear arrays created from microfluidics are particularly efficient in terms of time and materials for arrays requiring only a small number (< 100) of different surfacebound biomolecules, whereas the generalized photopatterned array fabrication methodology is more applicable to systems where the number of surface-attached biomolecules is greater than 100. The successful application of DNA, peptide, and carbohydrate arrays to study protein binding interactions demonstrates the wide variety of bioaffinity interactions that can be observed with SPR imaging. The study of protein–protein interactions by SPR imaging is still in its infancy, but we predict that this technology will have a significant role to play in the high-throughput screening of protein–protein interactions, the study of synthetic chemical libraries, the binding and inhibition of cell receptors, and the elucidation of regulatory pathways in biological systems. 7 Acknowledgments

This research is funded by the National Institute of Health (3R01 GM59622-03S 1 and 8 ROI EB00269-02) and the National Science Foundation (CHE-0133151).

References

References 1 MILLER, J. C., BUTLER, E. B., TEH, B. S., HAAB, B. B. The application of protein microarrays to serum diagnostics: Prostate cancer as a test case. Disease Markers 2001, 17, 225–234. 2 ZHU, H., KLEMIC, J. F., CHANG, S., BERTONE, P., CASAMAYOR, A., KLEMIC, K. G., SMITH, D., GERSTEIN, M., REED, M. A., SNYDER, M. Analysis of yeast protein kinases using protein chips. Nature Genet. 2000, 26, 283–289. 3 ZHU, H., BILGIN, M., BANGHAM, R., HALL, D., CASAMAYOR, A., BERTONE, P., LAN, N., JANSEN, R., BIDLINGMAIER, S., HOUFEK, T., MITCHELL, T., MILLER, P., DEAN, R. A., GERSTEIN, M., SNYDER, M. Global analysis of protein activities using proteome chips. Science 2001, 293, 2101–2105. 4 LAUNE, D., MOLINA, F., MANI, J.-C., DEL RIO, M., BOUANANI, M., PAU, B., GRANIER, C. Dissection of an antibody paratope into peptides discloses the idiotope recognized by the cognate anti-idiotypic antibody. J. Immun. Methods 2000, 239, 63–73. 5 UTHAIPIBULL, C., AUFIERO, B., SYED, S. E. H., HANSE, B., GUEVARA PATINO, J. A., ANGOV, E., LING, I. T., FEGERDING, K., MORGAN, W. D., OCKENHOUSE, C., BIRDSALL, B., FEENEY, J., LYON, J. A., HOLDER, A. A. Inhibitory and blocking monoclonal antibody epitopes on merozite surface protein 1 of the malaria parasiste plasmodium falciparum. J. Mol. Biol. 2001, 307, 1381–1394. 6 HOUSEMAN, B. T., HUH, J. H., KRON, S., J. MRKSICH, M. Peptide chips for the quantitative evaluation of protein kinase activity. Nature Biotech. 2001, 20, 270–274. 7 KILPERT, K., HANSEN, G., WESSNER, H., SCHNEIDER-MERGENER, J., HOHNE, W. Characterizing and optimizing protease/peptide inhibitor interactions, a new application for spot synthesis. J. Biochem. 2000, 128, 1051–1057. 8 BROCKMAN, J. M., FRUTOS, A. G., CORN, R. M. A multi-step chemical modification procedure to create DNA arrays on gold surfaces for the study of protein–DNA interactions with surface plasmon resonance imaging. J. Am. Chem. Soc. 1999, 121, 8044–8051.

9 FRUTOS, A. G., BROCKMAN, J. M., CORN, R. M. Reversible protection and reactive patterning of amine- and hydroxyl-terminated self-assembled monolayers on gold surfaces for the fabrication of biopolymer arrays. Langmuir 2000, 16, 2192–2197. 10 WEGNER, G. J., LEE, H. J., CORN, R. M. Characterization and optimization of peptide arrays for the study of epitope–antibody interactions using surface plasmon resonance imaging. Anal. Chem. 2002, 74, 5161–5168. 11 NELSON, B. P., FRUTOS, A. G., BROCKMAN, J. M., CORN, R. M. Near-infrared surface plasmon resonance measurements of ultrathin films. 1. Angle shift and SPR imaging experiments. Anal. Chem. 1999, 71, 3928–3934. 12 Hansen, W. N. ELECTRIC fields produced by the propagation of plane coherent electromagnetic radiation in a stratified medium. J. Opt. Soc. Am. 1968, 58, 380–390. 13 NELSON, B. P., GRIMSRUD, T. E., LILES, M. R., GOODMAN, R. M., CORN, R. M. Surface plasmon resonance imaging measurements of DNA and RNA hybridization adsorption onto DNA microarrays. Anal. Chem. 2001, 73, 1–7. 14 LEE, H. J., GOODRICH, T. T., CORN, R. M. SPR imaging measurements of 1-D and 2-D DNA microarrays created from microfluidic channels on gold thin films. Anal. Chem. 2001, 73, 5525–5531. 15 PEASE, A. C., SOLAS, D., SULLIVAN, E. J., CRONIN, M. T., HOLMES, C. P., FODOR, S. P. A. Light-generated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl Acad. Sci. USA 1994, 91, 5022–5026. 16 SCHENA, M., SHALON, D., DAVIS, R. W., BROWN, P. O. Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science 1995, 270, 467–470. 17 BRENNER, S., JOHNSON, M., BRIDGHAM, J., GOLDA, G., LLOYD, D. H., JOHNSON, D., LUO, S., MCCURDY, S., FOY, M., EWAN, M., ROTH, R., GEORGE, D., ELETR, S., ALBRECHT, G., VERMAAS, E., WILLIAMS, S. R., MOON, K., BURCHAM, T., PALLAS, M.,

21

22 Surface Plasmon Resonance Imaging Measurements of Biomolecular Arrays

18

19

20

21

22

23

24

25

DUBRIDGE, R. B., KIRCHNER, J., FEARON, K., MAO, J., CORCORAN, K. Gene expression analysis by massively parallel signature sequencing (MPS) on microbead arrays. Nature Biotech. 2000, 18, 630–634. WINZELER, E. A., RICHARDS, D. R., CONWAY, A. R., GOLDSTEIN, A. L., KALMAN, S., MCCULLOUGH, M. J., MCCUSKER, J. H., STEVENS, D. A., WODICKA, L., LOCKHART, D. J., DAVIS, R. W. Direct allelic variation scanning of the yeast genome. Science 1998, 281, 1194–1197. WENSCHUH, H., VOLKMER-ENGERT, R., SCHMIDT, M., SCHULZ, M., SCHNEIDER-MERGENER, J., REINEKE, U. Coherent membrane supports for parallel microsynthesis and screening of bioactive peptides. Biopolymers 2000, 55, 188–206. FRANK, R. Spot-synthesis: An easy technique for the positionally addressable, parallel chemical synthesis on a membrane support. Tetrahedron 1992, 48, 9217–9232. REINEKE, U., VOLKMER-ENGERT, R., SCHNEIDER-MERGENER, J. Applications of peptide arrays prepared by the SPOT-technology. Curr. Opin. Biotech. 2001, 12, 59–64. FREY, B., CORN, R. M. Covalent attachment and derivitization of poly(L-lysine) monolayers on gold surfaces as characterized by polarization-modulation FT-IR spectroscopy. Anal. Chem. 1996, 68, 3187–3193. SMITH, E., WANAT, M. J., CHENG, Y., BARREIRA, S. V. P., FRUTOS, A. G., CORN, R. M. Formation, spectroscopic characterization, and applications of sulfhydryl-terminated alkanethiol monolayers for the chemical attachment of DNA onto gold surfaces. Langmuir 2001, 17, 2502–2507. HIRSCHHORN, J. N., SKLAR, P., LINDBLAD-TOH, K., LIM, Y. M., RUIZ-GUTIERREZ, M., BOLK, S., LANGHORST, B., SCHAFFER, S. E., WINCHESTER, E., LANDLER, E. S. SBE-TAGS: An array-based method for efficient single-nucleotide polymorphism genotyping. Proc. Natl Acad. Sci. USA 2000, 97, 12164–12169. REICH, D. E., CARGILL, M., BOLK, S., IRELAND, J., SABETI, P. C., RICHTER, D. J., LAVERY, T., KOUOUMJIAN, R., FARHADIAN,

26 27

28

29

30

31

32

33

34

35

S. F., WARD, R., LANDER, E. S. Linkage disequilibrium in the human genome. Nature 2001, 411, 199–204. Brookes, A. J. THE essence of SNPs. Gene 1999, 234, 177–186. BROCKMAN, J. M., NELSON, B. P., CORN, R. M. Surface plasmon resonance imaging measurements of ultrathin organic films. Annu. Rev. Phys. Chem. 2000, 51, 41–63. SU, S. S., LAHUE, R. S., AU, K. G., MODRICH, P. Mispair Specifity of Methy-directed DNA Mismateh Correction in vitro. J. Biol. Chem. 1987, 263, 6829–6835. Hoch, J. A. TWO component and phosphorelay signal transduction. Curr. Opin. Microbiol. 2000, 3, 165–170. SMITH, E. A., ERICKSON, M. G., ULIJASZ, A. T., WEISBLUM, B., CORN, R. M. Surface plasmon resonance imaging of transcription factor proteins: interactions of bacterial response regulators with DNA arrays on gold films. Langmuir 2003, 19, 1486–1492. MARTINEZ-HACKERT, E., STOCK, A. M. Structural relationships in the OmpR family of winged-helix transcription factors. J. Mol. Biol. 1997, 269, 301–312. HALDIMANN, A., FISHER, S. L., DANIELS, L. L., WALSH, C. T., WANNER, B. L. Transcriptional regulation of the Enterococcus faecium BM4147 vancomycin resistance gene cluster by the VanS-VanR two-component regulatory system in Escerichia coli K-12. J. Bacteriol. 1997, 179, 5903–5913. JACKMAN, R. J., DUFFY, D. C., OSTUNI, E., WILLMORE, N. D., WHITESIDES, G. M. Fabricating large arrays of microwells with arbitrary dimensions and filling them using discontinuous de-wetting. Anal. Chem. 1998, 70, 2280–2287. DUFFY, D. C., MCDONALD, J. C., SCHUELLER, O. J. A., WHITESIDES, G. M. Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 1998, 70, 4974–4984. SLOOTSTRA, J. W., KUPERUS, D., PLUCKTHUN, A., MELOEN, R. H. Identification of new tag sequences with differential and selective recognization properties for the anti-FLAG monoclonal

References

36

37

38

39

40

antibodies M1, M2, and M5. Mol. Divers. 1996, 2, 156–164. SCHREIBER, S. L., MACBEATH, G. Printing proteins as microarrays for high-throughput function determination. Science 2000, 289, 1760–1763. BORNHORST, J. A., FALKE, J. J. Purification of proteins using polyhistidine affinity tags. Methods Enzymol. 2000, 326, 245–254. SCHMITT, J., HESS, J., STUNNENBERG, H. G. Affinity purification of histidine-tagged proteins. Mol. Biol. Rep. 1993, 18, 223–230. HO, C. H., LIMBERIS, L., CALDWELL, K. D., STEWART, R. J. A meta-chelating pluronic for immobilization of histidine-tagged proteins at interfaces: Immoblilization of firefly luciferase on polystyrene beads. Langmuir 1998, 14, 3889–3894. SIMONS, P. C., VANDER JAGT, D. L. Purification of glutathione S-transferase from human liver by glutathione-affinity

chromatography. Anal. Biochem. 1977, 82, 334–341. 41 WANG, C., CASTRO, A. F., WILKES, D. M., ALTENBERG, G. A. Expression and purification of the first nucleotide-binding domain and linker region of human multidrug resistance gene product: Comparison of fusions to glutathione S-transferase, thioredoxin and maltose-binding protein. Biochem. J. 1999, 338, 77–81. 42 CATTOLI, F., SARTI, G. C. Separation of MBP fusion proteins through affinity membranes. Biotechnol. Prog. 2002, 18, 94–100. 43 BACH, H., MAZOR, Y., SHAKY, S., SHOHAM-LEV, A., BERDICHEVSKY, Y., GUTNICK, D. L., BENHAR, I. Escherichia coli maltose-binding protein as a molecular chaperone for recombinant intracellular cytoplasmic single-chain antibodies. J. Mol. Biol. 2001, 312, 79–93.

23

1

Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors Gary Franklin, and Alan McWhirter Biacore AB, Uppsala, Sweden

Originally published in: Protein Microarray Technology. Edited by Dev Kambhampati. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30597-1

1 Introduction

The phenomenon of SPR (surface plasmon resonance) was ﬁrst shown to be applicable to biological molecules as long ago as 1983 [1] and has since played an increasingly important role in the study of biomolecular interactions. Biacore, the pioneering company in the optical biosensor ﬁeld, combined SPR detection with an advanced system of microﬂuidics and specially developed sensor surface coupling chemistries to generate a unique technology for generating high quality, information-rich data on molecular binding events. The technology is label-free, highly ﬂexible in terms of the range of biomolecules that it can accommodate and can deliver a wide range of data types, including analyses of concentration, speciﬁcity, afﬁnity and kinetics. The technology has been used very successfully in a number of areas, of which the three major areas have been academic life science research, commercial pharmaceutical and biotechnology applications and within the food industry. The instruments available cover a wide assortment of user requirements, ranging from manual systems designed for simple, ﬂexible basic research applications, to automated, high-throughput systems aimed at the drug development industry, as well as systems designed to function within the GxP (Good Laboratory Practice, Good Manufacturing Practice, Good Clinical Practice etc.) environment now required by many regulatory authorities. Biacore users have taken advantage of the inherent ﬂexibility of the technology to study many diverse types of molecules, although proteins have consistently represented the principle focus. As the genomics era begins to give way to the age of proteomics, the need for high quality data on the molecular interactions of proteins has never been greater. While Biacore, along with many other technologies, Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

was making an important contribution towards the understanding of protein interactions long before the term “proteomics” ever existed, the widely anticipated proteomics explosion will certainly present new challenges and opportunities to all working within this ﬁeld. 1.1 The Proteomics Revolution

During the last decade of the 20th century, biomedical science was characterized by the technological advances that powered the genomics revolution. The race to complete the sequencing of the human genome was probably the most absorbing scientiﬁc development of its time. We are now in possession of a staggering amount of information regarding the nucleic acid sequence of our own, and many other species. The total number of human genes (which has not yet been totally covered by the “completion” of the human genome project) is currently estimated at around 27 000 [2]. When compared to genomic sequencing projects carried out in other species however, it is clear that there is no real correlation between gene number and biological complexity among different organisms and that higher levels of genetic organization and control of expression of the genetic material must be involved. The primary challenge now is to relate this vast store of sequence information to its biological functions. Since the vast majority of genes are currently thought to function at the protein level, the next logical step is to re-examine the genome in terms of the proteins it produces. This will require an even larger-scale research effort however, since it is becoming increasingly clear that the number of distinctly functional human proteins produced is much higher than the number of genes that exist. This provides a likely explanation for the “gene number versus complexity” paradox referred to above. The genes of higher eukaryotes are complex in structure, often containing multiple promoters that direct alternative transcripts, and recent evidence suggests that around 50 % of all genes may be subject to alternative splicing [3]. In addition to multiple transcripts and splice variants, it has also been estimated that on average, each human gene may produce three different post-translationally modiﬁed protein variants, all with distinct functions [4]. Taken together, these variables mean that there are multiple proteins produced from each gene, with current estimates ranging anywhere between 3 and 20 proteins per gene. The term “proteome” was ﬁrst coined in the mid-1990s to introduce the concept of a protein-level equivalent of “genome”, and can be loosely deﬁned as “the entire protein complement that is encoded by a genome”. The proteome concept is considerably more complex than its genomic predecessor however, since the protein complement varies enormously between different cells within the organism and can change from minute to minute within an individual cell, depending on the internal and external signals received. Another factor is that the functional roles of proteins in the proteome involve interaction on a much greater scale of complexity than for nucleic acids. In addition, identiﬁcation of a protein on the basis of its amino acid composition alone may not provide the crucial information in all cases, since the post-translational modiﬁcations alluded to previously can play a critical role in physiological and pathological functions (reviewed in [4]).

2 Biacore Technology

Consequently, there are multiple challenges for the much-vaunted “new era” of proteomics. Very high throughput discovery programs will be required to identify all the proteins produced by the genome and to determine where and when these proteins are actually expressed in very complex, dynamic combinations. In addition, entirely different, lower throughput approaches will be required to provide the detailed characterization of interactions between functionally connected biomolecules. This last aspect in particular (embodied in the branch known as “interaction proteomics” or “functional proteomics”) illustrates the magnitude of the problems ahead. It should be clear from the application examples presented here (see Section 3) that understanding biomolecular interactions in the context of biological function requires a far higher level of detail than “do they bind?” and that the ability to describe interactions in terms of afﬁnity and (particularly) rate constants will be a prerequisite for real progress in the future. Biomolecular interactions are transient events and their kinetic characteristics are vitally important for biological function. A full understanding of how proteins function on a mechanistic level cannot, therefore, rely on steady-state measurements alone [5–8]. This temporal aspect of functional proteomic studies adds yet another level of complexity. Many different technologies will doubtless play important roles in the development of proteomics, and Biacore is ideally placed to make a signiﬁcant contribution. In fact, even before the term “proteomics” existed, Biacore has always been used in approaches that fundamentally belong to the proteomics ﬁeld. The ability of Biacore to measure protein concentrations, assess binding speciﬁcity, to characterize binding interactions in terms of afﬁnity and rate constants and (as described in Section 4) provide identiﬁcation by linking SPR to MS, make this a key proteomics technology. Moving from the study of isolated proteins or individual binding partners into more industrialized proteomics applications does not, therefore, represent a fundamental paradigm shift but is merely an issue of scale, as will be discussed later. This article will now describe how Biacore’s approach to SPR technology functions on a technical level and how these features contribute to the study of biomolecular interactions. This will be followed by selected examples of how the technology has been successfully applied to a number of important questions, with relevance to both academic biomedical research and the pharmaceutical and biotechnology industries. Finally, current and future developments will be discussed in the context of the evolving areas of proteomics and protein microarrays.

2 Biacore Technology

Biacore’s optical biosensors are built around three principal components: Ĺ an optical system responsible for the generation and detection of the SPR signal that is triggered by biomolecular interaction events Ĺ a sensor chip with a biospeciﬁc surface where biomolecular interactions take place Ĺ a liquid handling system with precision pumps and an integrated microﬂuidic cartridge (IFC) for controlled ﬂow of sample over the sensor surface

3

4 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

Fig. 1 Ligand, analyte and capturing molecule in relation to the sensor surface, as deﬁned in a Biacore assay.

These three cornerstones of the technology will be described in detail, along with a discussion of their practical and theoretical consequences. In the interests of clarity, some specialized terminology relating to the interactants used in a Biacore assay should also be pointed out. The interactant immobilized on the sensor surface is referred to as the ligand and should not be confused with the use of this term in the biological context of ligands/receptors. The ligand can be attached either directly to the sensor surface, or indirectly via an immobilized capturing molecule, and the interactant in free solution is referred to as the analyte. This is shown schematically in Figure 1. 2.1 Surface Plasmon Resonance

The detection principle of the technology depends upon the phenomenon of surface plasmon resonance (SPR), which occurs in thin conducting ﬁlms at an interface between media of different refractive index [9]. In Biacore systems, the media are the glass of the sensor chip and the sample solution, and the conducting ﬁlm is a gold layer on the sensor chip surface. Under conditions of total internal reﬂection, the light leaks an electric ﬁeld intensity called an evanescent wave ﬁeld across the interface into the medium of lower refractive index, without actually losing net energy. The amplitude of the evanescent ﬁeld wave decreases exponentially with distance from the surface, and the effective penetration depth is about half the wavelength of the incident light. In Biacore instruments, the light source is a near-infrared LED of peak intensity wavelength 760 nm and the penetration distance of the evanescent wave is about 300 nm. At a certain combination of angle of incidence and energy (wavelength), the incident light excites plasmons (electron charge density waves) in the gold ﬁlm. As a result, a characteristic absorption of energy via the evanescent wave ﬁeld occurs and SPR is seen as a drop in the intensity of the reﬂected light at a speciﬁc incident angle (the SPR angle). These principles are illustrated in Figure 2. The reduced intensity of reﬂected light is not caused by light absorption in the sample in the conventional (transmission spectroscopy) sense. The light is totally internally reﬂected inside the optical unit, and it is the evanescent wave that

2 Biacore Technology

Fig. 2 The principle of SPR detection in Biacore instruments. Schematic drawing showing the optical interface, sensor chip and microﬂuidic components that comprise the main technology cornerstones in Biacore’s approach to SPR biosensors. As drawn here, the ligand is immobilized on the lower (dark grey) surface of the sensor chip and the analyte is passed over the surface under continuous ﬂow conditions, via the integrated microﬂuidics system. An evanescent wave is created at the chip surface by the action of total internal reﬂection of the incident light, and this penetrates the sample side of the interface. As indicated in the enlarged view, the amplitude of

this wave decays exponentially with distance from the surface. Polarized components of the evanescent wave ﬁeld excite electromagnentic surface plasmon waves within the gold ﬁlm, enhancing the evanescent wave and causing a characteristic drop in the reﬂected light intensity. Changes in refractive index at the ﬂow cell side of the interface caused by changes in mass concentration at the sensor surface, result in a shift in the angle of incidence required to create surface plasmon resonance. This SPR angle is measured as a change in the detector position for the reﬂected intensity dip that results from the SPR phenomenon.

penetrates the sample. The signiﬁcance of this feature is that measurements can be made on turbid or even opaque solutions, without interference from conventional light absorption or scattering by the sample. When the detecting molecule is attached to the sensor chip or when analyte binds to the detecting molecule, the concentration at the sensor chip surface increases leading to an SPR response, observed as a change in the SPR angle. The speciﬁc refractive index contribution (i.e. the change in refractive index produced by a unit change in concentration) is very similar for different proteins regardless of composition, and values for other macromolecules are of the same order of magnitude. Consequently, the response measured is related to the mass of analyte bound and is largely independent of the nature of the analyte. Refractive index contributions for different molecules are additive, so that the amount of detecting molecule attached and the amount of analyte bound can both be measured with the same detection principle.

5

6 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

2.2 The Sensor Surface and Immobilization Chemistry

As outlined in the previous section, SPR measurements with Biacore are based on the interaction of an analyte in solution with a ligand attached to the sensor surface. At the heart of this process is the sensor chip, which provides both the physical conditions necessary to generate the SPR signal, and the surface on which the interaction being studied occurs. The speciﬁcity of an SPR analysis is determined through the nature and properties of the molecule attached to the sensor surface. Broadly speaking, biomolecules may be attached to the surface of the sensor chip using three different approaches: 1. Covalent immobilization, where the molecule is attached to the surface through a covalent chemical link. 2. High afﬁnity capture, where the molecule of interest is attached by non-covalent interaction with another molecule (which in turn is usually attached using covalent immobilization). 3. Hydrophobic adsorption, which exploits non-speciﬁc hydrophobic interactions between the molecule of interest (or a hydrophobic carrier such as a lipid monolayer or bilayer) and the sensor chip surface. 2.2.1 Sensor chips The sensor chip has two essential features: Ĺ A glass surface covered with a thin gold layer, creating the conditions required for generating an SPR signal. Ĺ A coating of some sort on the gold layer, providing a means for attaching the ligand and an environment where the interaction being studied will occur. The coating does not contribute directly to the SPR effect, and varies between different types of chips.

Although SPR can be generated in thin ﬁlms made from different materials, all Biacore chips are coated with a uniform thin layer of gold. Gold gives a well-deﬁned reﬂectance minimum with easily handled visible light wavelengths, and is also amenable to covalent bonding of surface matrix layers, and at the same time the metal is largely inert in physiological buffer conditions. Unmodiﬁed gold is not generally a suitable surface environment for biomolecular interactions however, and is covered with a covalently bonded monolayer of alkanethiol molecules. This serves to protect the biological samples from contact with the gold and at the same time provides a means of attachment of a surface matrix, which further enhances the usefulness of the surface in biomolecular contexts. On most sensor chips, the surface is covered with a “hydrogel” matrix of carboxymethylated dextran, a ﬂexible unbranched carbohydrate polymer forming a surface layer of approximately 25–100 nm thick. The dextran is covalently attached to an epoxide-modiﬁed alkanethiol monolayer. Surfaces for various applications have been tailored by the size of the dextran, differing in the range of 10 kDa to

2 Biacore Technology

1 MDa [10]. The dextran matrix imparts several important properties to the sensor surface: Ĺ It provides a hydrophilic environment favorable to most solution-based biomolecular interactions. Ĺ It provides a deﬁned chemical basis for covalent attachment of biomolecules to the surface using a wide range of well-deﬁned chemistries (as will be described shortly). Ĺ It increases the surface capacity many-fold in comparison with a ﬂat surface. Ĺ It extends the surface detection principle to encompass a thin layer with an extension of the same order of magnitude as the penetration depth of the evanescent wave responsible for the detection principle. The sensitivity of the detection is thereby increased considerably in comparison with a ﬂat surface.

Dextran is also an inert molecule in the context of biomolecular interactions, and the ﬂexibility of the unbranched polymer chains allows the “surface-attached” biomolecules to move with relative freedom within the surface layer. Dextranbased surface matrices therefore provide an excellent environment for biomolecular interactions [10]. Furthermore, the most frequently utilized carboxymethylated derivative of dextran improves the hydrophilicity and serves as an excellent starting point for various immobilization alternatives. The negatively charged polymer may give rise to unwanted electrostatic interactions with the immobilized ligands and thus to a risk of denaturation. These potential effects are normally eliminated by working under physiological salt conditions, where the electrostatic effects are effectively shielded. The basic structure of a sensor chip and its surface is shown in Figure 3.

Fig. 3 Schematic illustration of the structure of the sensor chip surface.

7

8 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

While this is not the forum for a detailed description of speciﬁc sensor chips manufactured by Biacore, a number of variations exist on the basic structure described which are designed for a range of speciﬁc application areas. Some of these speciﬁc application-related formats will be discussed later, where appropriate. 2.2.2 Immobilization Chemistry As mentioned previously, ligands can be attached to the sensor chip using a variety of chemical methods, allowing a broad range of different types of biomolecules to be analyzed by SPR. The gold layer and dextran matrix on the sensor surface are stable under a wide range of conditions, including extremes of pH and many organic solvents. Once the ligand has been immobilized, the stability of the sensor surface is determined primarily by the stability of the attached ligand itself. The carboxymethylated dextran matrix on the sensor surface is amenable to a range of chemistries for ligand immobilization which exploit different groups on the ligand molecule. Immobilization approaches may be directed towards amine, carboxyl, thiol or hydroxyl groups on the ligand, or may use speciﬁc tags introduced into the ligand by either chemical modiﬁcation or recombinant techniques. Amine coupling chemistry is the most widely applicable approach for attaching biomolecules covalently to the sensor surface. With this method, the dextran matrix on the sensor chip surface is ﬁrst activated with a mixture of 1-ethyl-3(3-dimethylaminopropyl) carbodiimide (EDC and N-hydroxysuccinimide (NHS) to give reactive succinimide esters [11]. The ligand is then passed over the surface and the esters react spontaneously with amino groups or other nucleophilic groups to link the ligand covalently to the dextran (see Figure 4). Most proteins contain several amine groups so that efﬁcient attachment can be achieved without seriously affecting the biological activity of the ligand. In some instances, however, amine coupling may involve groups at or near the active site or binding site of the ligand, with the result that attachment is accompanied by loss of activity. In such cases, the ligand can be attached using alternative coupling chemistry or a capturing approach. Thiol coupling [12] utilizes exchange reactions between thiol and active disulﬁde groups. The active disulﬁde moiety may be introduced either on the dextran matrix (to exchange with a thiol group on the ligand, referred to as the ligand thiol approach) or on the ligand molecule (to exchange with a thiol group introduced on the dextran

Fig. 4 Amine coupling of ligand to the sensor chip surface.

2 Biacore Technology

Fig. 5 Ligand thiol (A) and surface thiol (B) coupling of the ligand to the sensor surface.

matrix, referred to as the surface thiol approach). The reagent 2-(2-pyridinyldithio) ethaneamine (PDEA) can be used for introducing active disulﬁde groups. The amine group in PDEA can be used to attach the molecule to activated carboxyl groups on either the surface or the ligand. The thiol approach can help to immobilize ligands in a deﬁned orientation, since the number of potential attachment sites is often less than with amine coupling, and in many cases is reduced to one single site. Surface thiol coupling is also valuable for acidic proteins, since the substitution with PDEA raises the isoelectric point of the protein, improving the electrostatic interaction of ligand and surface matrix prior to coupling. The two different thiol coupling approaches are illustrated in Figure 5. Aldehyde coupling is a third chemical immobilization method [12], and is useful for ligands containing aldehyde groups, which can be either native or introduced by oxidation of cis-diols. In this case, immobilization requires activation of the surface with hydrazine or carbohydrazide. Aldehyde coupling provides an alternative approach for immobilizing glycoproteins and other glycoconjugates. The method is particularly suitable for ligands containing sialic acid, since these residues are very easily oxidized to aldehydes. The chemistry of aldehyde coupling is summarized in Figure 6.

2.2.3 General Capture Methods Immobilization of ligand by capturing involves high afﬁnity binding of the ligand to an immobilized capturing molecule. In general, the regeneration step in the assay procedure removes ligand along with any remaining bound analyte, so that new ligand needs to be captured for each assay cycle. The capturing molecule is

9

10 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

Fig. 6 Aldehyde coupling of ligand to the sensor surface.

attached to the surface using one of the covalent chemical approaches described previously. Capturing approaches can provide an alternative to covalent immobilization in situations where it is difﬁcult to ﬁnd chemical methods that give satisfactory results, or where the assay requires that the ligand on the surface can be changed. Capturing can also provide a ligand puriﬁcation step, for example, when tagged recombinant ligands are captured from partially puriﬁed material through binding of the tag to a speciﬁc capturing molecule. The basic requirement for successful capturing is a robust high afﬁnity interaction between the capturing molecule and the ligand. Monoclonal antibodies are frequently used as capturing agents. 2.2.4 Specialized Capture Methods Although one of the major advantages of SPR technology is label-free detection of binding interactions, commonly used labeling techniques employed for puriﬁcation or other purposes can also be taken advantage of for the purposes of ligand immobilization. Biotin-labeled molecules for example, can be captured directly onto sensor chips pre-immobilized with streptavidin [12]: while this is technically a form of capture, the afﬁnity of the streptavidin–biotin interaction (10−15 M) makes this a permanent immobilization of the ligand in practice. This option is particularly useful when using nucleic acid ligands, but is also used for other applications, such as biotinylated liposomes [13] and biotinylated glycosaminoglycans such as heparin [14]. Histidine (his)-tagged recombinant proteins may also be conveniently captured onto a specialized sensor chip surface on which nitrilotriacetic acid (NTA) has been covalently linked to the carboxymethylated dextran matrix. This surface is activated by a simple pulse of NiCl2 , which forms a chelating complex with NTA that binds polyhistidine peptides [15]. This is a useful generic method for working with his-tagged proteins and the captured ligands are easily removed by a pulse of EDTA. 2.2.5 Lipids and Membrane Proteins Hydrophobic molecules such as lipids present specialized requirements for SPR analysis, speciﬁcally in terms of sensor surface attachment, since they cannot be directly coupled to a standard carboxymethylated dextran surface for interaction studies with other molecules. This is an important issue, particularly since membrane-bound receptors are also a very important class of molecules for

2 Biacore Technology

biomedical research and these are most appropriately studied in the context of a lipid environment. To facilitate these types of applications, speciﬁc sensor chips are available. These either have a ﬂat hydrophobic surface, suitable for adsorbed lipid monolayers, or a dextran surface modiﬁed with lipophilic compounds, which enables liposomes to maintain a membrane-like lipid bilayer structure on the sensor surface. In addition to these speciﬁc sensor chips, several other inventive approaches to lipid-related applications have also been demonstrated [13, 16–19]. Taken together, the range of available surface chemistry options enables the study of almost any conceivable ligand, from low molecular weight substances to whole cells. In the event of immobilization difﬁculties with a particular ligand, the option of using a capture molecule is also available (see Figure 1). 2.3 The Microﬂuidics System

The method of delivery of samples to the sensor surface is also a critical function and can vary considerably among the different commercial variants of SPR technology. The solution employed by Biacore involves a ﬂexible microﬂuidics system based around an integrated microﬂuidic cartridge, or IFC [20]. This consists of a series of micro-channels and membrane valves encased in a plastic housing, and serves to control delivery of liquid to the sensor chip surface. Samples are transferred through a needle and into the IFC, which connects with the detector ﬂow cells. These are formed by the sensor chip pressing against molded channels in the IFC. The precise construction of the IFC and the number of ﬂow cells available varies amongst the different instrument series, but the common purpose is to allow analyte to pass over the sensor surface in a continuous, pulse-free and controlled ﬂow that maintains a constant analyte concentration at the sensor surface during sample injection. An example of a microﬂuidics system is presented in Figure 7.

Fig. 7 The microﬂuidics system and ﬂow cells

11

12 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

In the ﬁrst Biacore instruments, samples were addressed in separate ﬂow cells formed by the IFC. More recently, a technique of hydrodynamic addressing has been introduced whereby solutions are directed over different detection spots in a single ﬂow cell, allowing more precise comparisons between spots. The construction is also designed to provide an almost instantaneous switch from sample to buffer, with minimum dispersion between the injections. These are important features of the technology, as they permit the SPR measurement of rapid events and enable, for example, very high-resolution kinetic studies to be undertaken. A more detailed discussion of the microﬂuidics system from the assay perspective will be taken up in Section 2.2.5. 2.4 Biacore Assay Basics

The deﬁnitions of basic assay formats and interaction partners are given at the beginning of Section 2, but it may also be useful to deﬁne some additional fundamental Biacore terminology at this point: Ĺ The resonance unit (RU) is the unit of measurement of the SPR response and is directly proportional to the concentration of biomolecules on the sensor surface. Ĺ The real-time SPR response is presented as a sensorgram, which plots response against time. Sensorgrams may be analyzed to provide information on the rates of interaction between molecules. Ĺ Report points are sensorgram responses taken at speciﬁc times, averaged over a short time window, or taken as the slope of the sensorgram over a deﬁned time window. Ĺ Regeneration is the process of removing bound analyte from the surface after an analysis cycle, in preparation for a new cycle.

SPR studies, as with most advanced, high technology molecular techniques, require care and planning from the user in order to provide reliable, high quality results. Proper instrument maintenance, intelligent planning of assays and careful experimental technique all are important factors. As mentioned in several places within this chapter, Biacore systems are designed to enable all appropriate controls to be incorporated into assays. While care should always be exercised when carrying out assays, particular attention needs to be paid when carrying out afﬁnity and kinetic assays. These approaches involve the simultaneous evaluation of multiple sensorgrams derived from concentration series of the analyte. It is very important here that the concentration dilutions are accurate and that the different solutions are carefully matched in order to calculate correct binding constants. 2.4.1 Correlation of SPR Response with Surface Binding While the SPR response is proportional to the concentration of bound analyte at the sensor surface, changes in the refractive index caused by the bulk solution

2 Biacore Technology

Fig. 8 Schematic sensorgram, illustrating the contribution of the bulk refractive index to the SPR signal.

will also contribute to the measured response. In practice, the bulk response is usually irrelevant in terms of data evaluation, since the binding response can be processed to take account of this. The absolute response measures the total SPR signal (including bulk contribution) and is normally recorded for both ligand and reference spots. The relative response subtracts a speciﬁed baseline value from the total SPR signal obtained. Depending upon the particular system and assay set-up used, a reference-subtracted response corresponding to the difference between ligand and reference spots can also be presented. To a ﬁrst approximation, this eliminates the contribution from the bulk solution and corresponds to the amount of ligand and analyte on the surface. A schematic sensorgram which indicates the bulk contribution effect is shown in Figure 8. The precise relationship between SPR signal and the amount of surface-bound analyte depends to some degree on the nature of the molecule involved, but is linear for any given molecule over experimentally relevant ranges. Direct studies using various proteins, for example, have shown a linear response and established an approximate conversion factor of 1000 RU ng−1 protein mm2 , using a standard carboxymethylated dextran sensor chip [21]. Taking a matrix depth of 100 nm, a 1000-RU response approximates to an equivalent solution protein concentration of 10 mg mL1 . This relationship varies for other types of molecules and it is known for example that nucleic acids have a slightly higher speciﬁc refractive index increment than proteins. In terms of detection limits, these vary depending on the molecules involved and the assay format used (see below), but the speciﬁed limits quoted, based on antibody detection of myoglobin, vary between 50 pM to less than 1 pM depending on the speciﬁc type of Biacore instrument. The sensorgram represents one of the key strengths of this technology, since it provides real-time, continuous monitoring throughout all phases of a biomolecular interaction. SPR responses are taken at a frequency of 1 to 10 s−1 , producing

13

14 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

an information-rich data format than enables evaluation of complex interaction properties, but which also gives immediate visual feedback on each interaction. SPR responses at deﬁned points during analysis cycles are taken from the sensorgram in the form of report points that are automatically assigned by the control software and/or assigned by the user. 2.4.2 Assay Design: Assay Formats In addition to direct binding assays (whether directly to an immobilized or captured ligand), a number of other assay formats are possible. Sandwich assays (direct binding with enhancement) are an extension to the direct assay in which a secondary “enhancement molecule” is allowed to interact with the bound analyte in order to increase the level of the SPR response. This approach can also be used to conﬁrm the identity of the analyte when working with complex mixtures for example, since two speciﬁc binding interactions must take place in order to see an enhanced response. Inhibition assays (or solution competition assays) exploit the ability of the analyte to inhibit the binding of a high MW detecting molecule to the surface. Typically, the analyte or a derivative thereof is attached to the surface as the ligand, while the detecting molecule (added to the samples at a ﬁxed, known concentration) is a macromolecule that binds speciﬁcally to the analyte. The mixture is incubated for a deﬁned time, and then injected over the sensor surface to measure the remaining free detecting molecules. The amount of free detecting molecule is then inversely related to the concentration of analyte in the sample. Inhibition assays are most commonly used for low MW analytes. They may however, also be used as an alternative to direct binding level assays for macromolecular analytes, in situations where immobilization or regeneration of the detecting molecule on the sensor surface is not satisfactory. In surface competition assays, a binding partner to the analyte is used as the ligand, and a high molecular weight analog to the analyte (typically analyte conjugated to a carrier protein) is added in constant amount to the samples to be measured. The basis of the assay is competition between analyte and the high MW analog for binding to the ligand. The measured response is the sum of the contributions from the analyte and the high MW molecule: as with the solution competition approach, the response is inversely related to the amount of analyte in the sample. The surface competition approach can have advantages over the more common inhibition assay format in situations where immobilization of the analyte on the sensor chip surface presents problems. The four basic assay strategies are shown in Figure 9. The sandwich and two competition-format assays are used widely for making concentration measurements, whereas afﬁnity and kinetic measurements principally employ direct binding assays. 2.4.3 Assay Design: Surface Preparation The range of surface coupling chemistry approaches supported by Biacore gives the user great freedom in designing an assay and choosing which interaction partner to immobilize on the sensor surface. In many cases, this choice can be made purely on the basis of experimental efﬁciency and convenience. In an ideal

2 Biacore Technology

Fig. 9 Schematic illustration of the four principle Biacore assay formats.

situation, the ligand and analyte identities of the biomolecules can be assayed in reciprocal experiments, conﬁrming that surface coupling does not affect the observed binding behavior. As described in Section 2.2, certain types of ligand may necessitate using speciﬁc sensor chips, although capture-type approaches can always be considered. However, it may be preferable to use certain molecules as ligand or analyte speciﬁcally, such as in the case of RNA molecules, which are generally more likely to be used as ligands. Their strong negative charge can produce electrostatic repulsion effects with the surface matrix at physiological pH if used as analyte which might of course affect apparent binding and kinetic properties [22]. It is often convenient in any case to capture nucleic acid ligands onto a streptavidin surface, given the frequent and convenient use of 5 biotin labeling of oligonucleotides. It is also worth remembering that the SPR signal is related to the mass concentration of bound molecule on the surface, so that the molecular weight (MW) of the interaction partners may also be an important factor. For a given number of interaction events, the SPR response is proportional to the MW of the analyte. Conversely, for a given level of ligand immobilization, a higher MW ligand will provide proportionately fewer binding sites. These issues are normally expressed in terms of Rmax , which deﬁnes the theoretical maximum binding capacity of the surface for a deﬁned pair of ligand and analyte molecules: Rmax =

MWanalyte ×Rligand ×n MWligand

(1)

15

16 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

where: Rmax is the maximum analyte binding capacity in RU; MWanalyte and MWligand are the molecular weights of analyte and ligand respectively; Rligand is the amount of immobilized ligand in RU and n is the stoichiometry of analyte binding to ligand. For example, with ligand molecular weight = 50,000 Da, analyte molecular weight = 10,000 Da and ligand immobilized level = 10,000 RU, the theoretical maximum binding capacity assuming 1:1 binding of analyte to ligand is given by Rmax =

10, 000 ×10, 000=2, 000RU 50, 000

(2)

In addition to aiming for immobilization levels in relation to the expected concentration and MW of the analyte, the optimal Rmax can vary considerably depending on the particular assay. Concentration assays, for example often use a relatively high Rmax of several hundred to several thousand RU, whereas for kinetic rate constant determinations, much lower Rmax values of the order of 10–50 RU are recommended (reviewed in [23]). 2.4.4 Regeneration The process of removing bound analyte from the sensor chip surface after analysis of a sample in preparation for the next analysis cycle (regeneration) is another important feature of Biacore’s approach to SPR. This enables sensor surfaces to be prepared with immobilized ligand and re-used for multiple rounds of analysis. The number of times a sensor surface can be regenerated depends on the nature of the attached ligand, but normally ranges from ﬁfty to several hundred. When the ligand is attached directly to the surface, the aim of regeneration is to remove the analyte from the ligand without destroying the ligand activity. When a capturing approach is used, regeneration generally removes both ligand and analyte from the capturing molecule, and the stability of the ligand under regeneration conditions is irrelevant. The optimal regeneration solution varies depending on the nature of the ligand–analyte interaction and the stability of the two molecules, but can often be achieved by simple exposure to acidic (dilute HCl or glycine-HCl) or basic (NaOH) solutions. Many other alternatives can be used, however, and it is possible to use sequential exposure to two different regeneration solutions and to vary the contact time for the solutions on the sensor surface. Efﬁcient regeneration is important for successful assays. Incomplete regeneration or loss of the binding activity from the surface impairs the performance of the assay and the useful lifetime of the sensor chip can be shortened. Suboptimal regeneration can be tolerated in some simple types of yes/no binding assay, since a certain loss of ligand activity can be adjusted for using suitable controls. The demands on efﬁcient regeneration are much stricter in the case of afﬁnity or kinetic assays however, since it is important to maintain a constant surface activity between cycles within the concentration series for each sample.

2 Biacore Technology

2.5 Theoretical and Practical Implications of Technology Design for Biacore Assays

The technological approaches taken by Biacore are designed to optimize SPR analysis of biomolecular interactions. Some of the consequences of these technology implementations have been referred to in the previous sections, but attention will now be focused on the more detailed theoretical and practical implications for designing and running SPR assays with Biacore. David Myszka has also written a review that covers many of these issues [23]. 2.5.1 Label-free Detection The label-free detection principle that forms the basis of SPR technology represents one of its strongest advantages over more traditional methods. In addition to being inconvenient and time consuming, labeling procedures may also have adverse effects on the behavior of the labeled molecule, by changing the properties of, or occluding the binding site for example. The label itself can also introduce complications: ﬂuorophores are usually hydrophobic and are frequently associated with background problems for example, and radiolabeling suffers from drawbacks connected with safety, isotopic decay and radiolysis-induced instability of labeled compounds. Labeling techniques are of course more sensitive in certain instances, but this must be weighed against the disadvantages of time-consuming and laborious procedures and high sample consumption. One important aspect of label-free SPR technology however, is the generic nature of the detection principle. The SPR response from any given analysis is derived from changes in the total mass concentration at the sensor surface and does not distinguish between potentially different molecular identities. This raises an important issue with regard to speciﬁcity and experimental design. Non-speciﬁc binding of analytes to the native sensor surface can be easily controlled for by incorporating a “blank” ﬂow cell in the analysis. This is rarely a problem however, and the most likely source of non-speciﬁc binding problems lies with the ligand chosen. For this reason, it is important to use very pure preparations of the intended ligand and to ensure that it is a sufﬁciently speciﬁc molecule for the intended assay, since this will largely determine the chances of success. Biacore’s ongoing development of combining SPR with mass spectrometry (MS, see Section 4.1) represent an interesting approach to provide molecular identiﬁcation along with binding information. This promises to be particularly useful in the case of ligand ﬁshing-type experiments, where potentially unknown binding partners may be involved. 2.5.2 The Optical Detection System and Bulk Effects As mentioned previously, the optical detection system in Biacore will detect all surface changes in refractive index, including those contributed by different bulk solutions. In most situations, using relative-responses and careful reference-subtraction will take care of these effects. Particular care is needed however, when working in conditions of low analyte responses, high surface ligand density and organic solvents with large bulk refractive index effects. These conditions most commonly arise

17

18 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

when working with low molecular weight drug compounds, which often require 1–10 % dimethylsulphoxide (DMSO) in order to maintain solubility. Under these conditions, the excluded volume effect between ligand-bound and blank reference surfaces (allowing more of the solvent to access to the blank surface) can produce bulk subtraction errors that are of the same order of magnitude as the expected analyte responses. This requires great care when preparing samples, since small differences in DMSO concentration will produce a signiﬁcant bulk response difference. Fortunately, this complication can be overcome by normalizing responses against a solvent correction curve prepared by running a concentration series of DMSO-containing buffer solutions over the ligand and reference surfaces [24]. 2.5.3 Surface-based Versus Solution-based Analysis SPR, as the name implies, is an intrinsically surface-based technique. Despite the many advantages that this provides, potential effects on molecular behavior caused by coupling one of the interaction partners to the sensor surface per se must also be considered. The possibility always exists for example, that chemical coupling of a ligand to the sensor surface may interfere with a critical analytebinding region (labeling procedures also carry an analogous risk, as discussed previously). This possibility can usually be tested for by carrying out an activity test on the immobilized sensor surface, using a positive control sample. In the unlikely event of such problems occurring, the range of coupling chemistry options available provides alternative immobilization strategies, as well as the possibility of reversing the experimental design and using the molecule in question as the analyte. Another question that has been raised is whether meaningful binding data can really be addressed by any surface-based method. The argument is that in contrast to solution-based assays, the restrictions imposed upon the interaction partner which is immobilized onto a surface will produce experimental artifacts. This is an over-simpliﬁcation of course, and interactions in simple solution assays are unlikely to precisely reﬂect many biomolecular interactions that occur in vivo. Furthermore, the carboxymethyl dextran hydrogel on Biacore sensor chips presents a ﬂexible matrix for ligand binding and probably allows a considerable degree of molecular movement [10]. Formal proof that surface-based methods are in fact highly appropriate for biomolecular interaction studies has come from studies designed to directly compare solution-based techniques with SPR. These have demonstrated that afﬁnity, kinetic and thermodynamic constants obtained using Biacore are extremely consistent with those obtained using a range of solutionbased techniques [25]. 2.5.4 The Microﬂuidic Design and Mass Transport Effects The liquid ﬂow system in Biacore’s approach to SPR lies at the heart of the design philosophy and contributes greatly to its powerful kinetics capabilities. As with all surface-based analysis methods however, there is one complicating factor that needs to be considered for both system and assay design, namely the issue of mass transport [26]. For the analyte to be able to bind to the sensor surface, the molecules must be transported from the bulk solution to the surface. This is a

2 Biacore Technology

diffusion-controlled process and under the laminar ﬂow conditions that exist in the IFC, the transport rate is proportional to the analyte concentration in the bulk solution, with a proportionality constant called the mass transport coefﬁcient (km ). This constant is proportional to the cube root of the liquid ﬂow rate and depends on ﬂow cell dimensions and the diffusion properties of the analyte in the sample solution [27]. If mass transport is much faster than the association of the analyte with the ligand, the observed binding will be determined by the interaction constants. Conversely, if mass transport is much slower than association, the binding will be wholly limited by the mass transport process and kinetic data for the interaction will not be obtained. In practice, many experimental systems will actually lie between these extremes, and the binding will be determined by contributions from both mass transport and interaction kinetics. This may be represented in a reaction scheme as: Abulk km Asurface + B ka AB km

kd

(3)

where km is the rate constant for mass transport to and from the surface and ka and kd are the rate constants for the formation and dissociation of the complex. It is important to understand the experimental parameters that determine whether the association rate is limited by mass transport or interaction. For a given analyte and ﬂow cell dimensions, only two factors are signiﬁcant: 1. The ﬂow rate, which affects the mass transport rate constant km but has no effect on the interaction kinetics. Increasing the ﬂow rate increases km , so that mass transport is less likely to be limiting. This effect is however relatively small, since km is proportional to the cube root of the ﬂow rate. 2. The concentration of surface binding sites, which affects the absolute interaction rate but has no effect on mass transport. This is the most important parameter. Increasing the surface binding capacity increases the initial interaction rate, so that mass transport is more likely to be limiting.

As an interaction proceeds, the concentration of available surface binding sites falls and interactions which are limited by mass transport in the initial phase can therefore, become interaction-limited as the amount of free ligand sites decreases. The mass transport rate is also inﬂuenced by the diffusion coefﬁcient of the analyte, which in turn depends on the molecular weight. A large analyte has a lower diffusion coefﬁcient, and will therefore be more subject to mass transport limitations than a small analyte with the same kinetic parameters. The very same parameters that govern the inﬂuence of mass transport during the association phase also apply during dissociation: released analyte can either diffuse into the bulk buffer ﬂow and be removed, or may in theory re-bind to the ligand. In this context too, a high ﬂow rate and low surface binding capacity will minimize any such effects. It is important to realize that mass transport is not a “problem” in SPR analysis, but a physical property of the system which can be readily accounted for in data

19

20 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

interpretation [28] and which can also be manipulated to some advantage during assay design. When carrying out concentration measurements, it can be desirable to perform assays under conditions where the interaction is entirely controlled by mass transport [29], since under such conditions, the binding rate is entirely dependent on the analyte concentration. This makes the assay independent of afﬁnity effects and therefore more robust for analysis of multiple analytes. These conditions are best achieved by using low ﬂow rates and immobilizing the ligand at a high surface-binding capacity. For assays designed to measure kinetic rate constants on the other hand, the requirements are reversed. If the rate of mass transport is much faster than the association of analyte with ligand, the observed binding will be determined entirely by the interaction constants. The optimal conditions to achieve this are a combination of high ﬂow rate and a low surface binding capacity [23]. The height of the ﬂow cell also affects the phenomenon [20] and some Biacore systems are designed with a lower ﬂow cell that helps to improve mass transport. Given that in practice assays can be governed by a combination of interaction constants and mass transport rate, the ﬁtting algorithms used to determine rate constants from sensorgram data are designed to incorporate a mass transport component that ensures correct analysis [30]. In the extreme case of a completely mass transport-limited interaction however, the sensorgrams will not contain any kinetic information and the calculated rate constants will be meaningless. Fortunately, software tools are now designed to deal with this potential problem and can identify binding interactions that are entirely controlled by mass transport. 3 Biacore Applications in Basic and Clinical Research 3.1 Introduction

The publication of around 400 papers each year covering a wide range of biological and clinical research ﬁelds is testament to the versatility of Biacore’s SPR technology, with examples of quantitative analyses of biomolecular interactions to be found in virtually all disciplines within the life sciences. New innovative applications are continually appearing in the literature and demonstrate that the technology has the potential to replace and often improve on the speed and data quality of established techniques. This section will review a selection of recently published applications with particular emphasis on protein:protein interactions and protein:nucleic acid interactions in cancer research and neuroscience, two areas within which Biacore has become ﬁrmly established in recent years. 3.2 Applications of Biacore in Cancer Research

The design of targeted therapeutic reagents to cellular signaling events that initiate oncogenic transformation or maintain a tumorigenic phenotype in a cell requires

3 Biacore Applications in Basic and Clinical Research

that the speciﬁcity and kinetics of the molecular interactions in the signaling cascades are accurately determined. Possibly the most accessible interactions against which intervention may be directed are those that occur at the cell surface between growth factors and constitutively active receptors. In one approach, peptide mimetics were designed based on a functionally inhibitory antibody to HER2, an oncogenic member of the EGF family of tyrosine kinase receptors which is commonly expressed in human breast, ovarian and colon cancer and which is associated with a poor prognosis [31]. A Biacore assay was used to screen and select peptide mimetics from a range of candidates that bound with high afﬁnity and with favorable kinetics, most importantly those with slow dissociation rates (Figure 10). Those with the slowest dissociation rates were also most effective at inducing functions such as reduced tyrosine phosphorylation of downstream signaling molecules in cell-based assays. Most encouragingly, cells treated with these mimetics were hypersensitive to radiation-induced apoptosis suggesting that the mimetics may be used clinically in conjunction with radiotherapy. Simultaneous competition studies using Biacore showed that the mimetics and the parent antibody were directed against the same epitopes on HER2 [32]. These studies also suggested that the receptor was disabled because the mimetics inhibited receptor dimerization [33], a prerequisite for the propagation of signals into the cytoplasm following engagement with a cognate ligand. A strategy of structure-based targeting of the dimerization interfaces of tyrosine kinase receptors may therefore be generally applicable to the design of antagonists of oncogenically transformed tyrosine kinase receptors. It is important to note that in this study, although the afﬁnity of the mimetic for HER2 was lower than the parent antibody, the dissociation rate constant, kd , was similar, indicating that the complex was stable. Potential therapeutic reagents may therefore be missed during a screening process that relies on afﬁnity data alone and both association and dissociation rate constants (ka and kd , respectively) may reveal effective binders (see [8] and [7] for detailed studies on the treatment of ka and kd data in Biacore assays to select effective binders from a large base of candidate drugs). Although growth factors generally bind with high afﬁnity to one favored cellular receptor, they may nevertheless also bind alternative receptors. Platelet-derived growth factor (PDGF), for example, exists both as a homodimer (AA or BB) and a heterodimer (AB). While PDGF-AA binds only the PDGF-α receptor, PDGF-BB and PDGF-AB can bind both the PDGF-α and -β receptors. In addition, it was recently shown in a Biacore study that the pleiotropic scope of PDGF-BB is wider still and binds low-density lipoprotein receptor-related protein [34]. It does not always follow, however, that the functional consequences of binding alternative receptors are easily differentiated. The functions of two different cell membrane receptors, lymphotoxin-β receptor (LTβR) and herpes virus entry mediator (HveA), both of which bind a common ligand, have now been deﬁned using Biacore. Peptides bearing selected point mutations were generated from LIGHT, a pro-apoptotic TNF ligand superfamily member. Mutants were selected that bound one, but not the other binding partner [35]. One selected mutant that bound HveA, but not LTβR in the Biacore assay, failed to induce apoptosis in target cells. The cell-based assays showed that a dominant negative mutant of TRAF3, an intracellular protein

21

22 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

Fig. 10 Screening a series of peptide mimetics using an SPR-based kinetic assay. (A) Dose dependence curves for binding of AHNP to the immobilized HER2 receptor. AHNP-Y was injected at concentrations of 0.5 (a), 1 (b), 2 (c), 4 (d) and 8 mM (e) at a ﬂow rate of 20 :L min. The sensorgrams show binding of AHNP-Y to the immobilized HER2 (the ﬁrst 300 s) followed by peptide dissociation from the receptor surface

(the ﬁnal 240 s). (B) Biacore analysis of the inhibitory effect of AHNP-Y on mAb 4D5 binding to immobilized HER2. Sensorgrams show binding of 1 nM mAb 4D5 to HER2 after pre-injection of 0 (a), 0.3 (b), 1.5 (c), 6 (d), 20 (e), and 60 mM (f) AHNP-Y. Inset: the correlation between the initial rate of 4D5 binding to HER2 and concentration of pre-injected AHNP-Y.

3 Biacore Applications in Basic and Clinical Research

that binds the cytosolic domain of LTβR and transmits signals leading to cell death, blocked LIGHT-mediated apoptosis. These functional cell-based assays conﬁrmed the Biacore binding data, indicating that LTβR but not HveA propagates LIGHTmediated apoptosis. Although mouse monoclonal antibodies have been raised to many human tumorspeciﬁc markers [36–39] their use in cancer treatment is greatly limited by the fact that the patient inevitably generates a potentially harmful immune response. Attempts to address these limitations have been made by the generation of singlechain Fv (scFv) molecules that retain speciﬁcity without presenting the highly antigenic constant regions of the whole antibody molecule to the host. In one variation of this strategy, a fully human, gene-fused bi-speciﬁc antibody (bs-Ab) was created with speciﬁcity both to IgG Fc receptors (CD16) on the surface of cytotoxic immune cells and to HER2 on tumor cells [40]. The hypothesis was that this bsAb might function as a bridge and actively recruit host cytotoxic immune cells to tumor cells. In separate Biacore experiments in which either CD16 or HER2 were immobilized on a sensor chip it was demonstrated that the gene fusion process did not greatly alter the binding kinetics of either of the component scFvs in the bs-Ab. Conﬁrmation of “conservation-of-function” after conjugation is an important stage in the development of any carrier system in which an active molecule is conjugated to a carrier. The function of the bs-Ab was also tested in vitro, where it increased lysis of human ovarian cancer cells which over-expressed HER2 when presented together with peripheral blood lymphocytes. While whole monoclonal antibodies are potentially useful reagents for diagnostic applications, human antibody fragments obtained via phage antibody technology are a more attractive alternative for human cancer therapy since they might be expected to penetrate tumors more efﬁciently than whole antibodies [41–43]. Fibronectin is a ubiquitous extracellular matrix protein, one variant of which contains a novel splice-generated domain (ED-B). This isoform is found in the stroma of fetal and neoplastic tissues and in the walls of developing neoplastic blood vessels, but is absent from normal adult tissues. Reasoning that this protein is a suitably generic tumor-speciﬁc target, a number of single chain antibody fragments and dimeric variants to ED-B were generated and an optimal binding fragment was selected for clinical trials [44]. Biacore’s SPR technology was used to screen antibody fragment libraries and to compare detailed binding properties to immobilized ED-B. The afﬁnity dissociation constants (KD ) of the antibody fragments for ED-B varied widely but those that bound ED-B with higher afﬁnity than the parent fragment were shown in most cases to do so as a result of slower dissociation rates. When fragments were assessed for tumor targeting in vivo using real-time photodetection of ﬂuorescently-labeled antibodies, the efﬁciency of tumor homing correlated closely with the slower off-rates of ED-B binding detected in the Biacore assays. Biacore’s SPR technology was therefore used to select optimal antibody fragments for diagnostic imaging and may therefore be useful in the preclinical stage of selecting reagents for therapeutic strategies targeted at the tumor vascula-ture. Cell surface receptors bind to their cognate ligands and to recruited intracellular second messengers at a hydrophobic interface: the external and internal faces of the

23

24 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

Fig. 11 Signal transduction occurs in the plasma membrane. Ras is anchored in the plasma membrane via one farnesyl and one or two palmitoyl modiﬁcations at its C-terminus, where activation by GEFproteins (here Sos) and recruitment of downstream effectors such as Raf-kinase occurs.

plasma membrane, respectively. Proteins assume markedly different structures in hydrophobic and aqueous microenvironments. Even if the transmembrane domain of the receptor is the only part of the molecule that sits directly within a lipid bilayer, structural disturbances in this region of the protein may alter the protruding intraand extracellular domains. It is thus reasonable to assume that interactions between proteins may yield a very different picture depending on the microenvironment in which the interaction is studied. A lipid microenvironment may be vital to allow immobilized receptors to assume a conﬁguration that reasonably mimics that found in situ and thereby present a functional binding interface for interacting molecules. For example, in order to interact with the serine/threonine kinase, Raf , the proto-oncogene, Ras, must insert into a biological membrane to transduce signals from activated receptor tyrosine kinases to the intracellular MAP-kinase pathway (Figure 11). In order to be able to make kinetic measurements of the interaction between Raf and Ras, the plasma membrane was thus simulated in vitro on a hydrophobic sensor chip by loading the chip with liposomes prior to molecular interaction analysis [45]. Biacore experiments showed that GTP-bound Ras bound speciﬁcally to Raf only if Ras was immobilized on the liposome-loaded chip by prior fusion with a synthetic lipid tail (Figure 12). In functional assays, cells that were microinjected with nominally oncogenic truncated Ras without a lipid tail did not transform into tumor cells. These cells, in contrast and in full agreement with the Biacore data, only transformed if truncated Ras was co-injected along with a lipid anchor. To be able to grow beyond a volume of about 1 mm3 , a tumor, once initiated, requires its own microvasculature. Inhibition of angiogenesis, or the process of neovascularization, is an attractive strategy in the intervention of tumor spreading. Members of the vascular endothelial cell growth factor (VEGF) family are principal effectors of angiogenesis as they inﬂuence the division and migration of endothe-

3 Biacore Applications in Basic and Clinical Research

Fig. 12 Insertion of lipid-modiﬁed Ras protein into an artiﬁcial membrane on Sensor Chip HPA. Sensor Chip HPA was treated with POPC (palmitoyl oleoyl phosphatidyl choline)-containing lipid vesicles and conditioned by treatment with NaOH and bovine

serum albumin at high ﬂow rates. Farnesylated Ras is inserted into the artiﬁcial membrane, while all non-covalently attached molecules are easily displaced from the lipid surface by the detergent octylglycoside.

lial cells by activating cognate transmembrane receptors, among them VEGFR-1 and VEGFR-2. Recombinant, soluble variants of these receptors that lack transmembrane domains can be used to sequester VEGF in solution and block binding to native cell-bound receptors. However, while cell-based approaches indicate that heparin is necessary to facilitate the functional presentation of VEGF to the receptors, cell-free assays suggest that this is not the case for the isolated soluble receptors. Kinetic analysis using Biacore showed that heparin actually inhibited the interaction with soluble receptors [46] and that this was due to a reduction in association rate. In this way, Biacore analysis therefore enabled baseline parameters to be established for the binding of soluble VEGFR-2 to VEGF and to eliminate the potential problem of a heparin requirement in therapeutic applications. Intervention in the deregulated cell cycle is another potential therapeutic anticancer strategy. Progression from the G1 (gap) phase to the S phase (DNA synthesis) of the cell division cycle is regulated by a family of cyclin-dependent protein kinases (CDKs) which are regulated in their turn by many other proteins including members of the cyclin family. In one study, binding between PCNA (proliferating cell nuclear antigen) and mutants of p21, a CDK inhibitor, was measured using both a yeast two-hybrid assay and Biacore [47]. Although these two approaches gave similar qualitative data, only Biacore provided information on binding afﬁnity and further showed that trimerization of PCNA is a prerequisite for p21 binding. The technology was also used to screen for short peptides that inhibited the p21–PCNA interaction in competition assays. One p21-like peptide was found that abolished this interaction beyond the level of detection.

25

26 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

Cyclin G is regulated by p53 in response to growth factor stimulation and its overexpression has been shown to inhibit the proliferation of human sarcoma cells. Although this implies a role for cyclins in cell cycle control, the physiological functions of cyclin G have not been characterized. A cDNA clone encoding a potential cyclin G binding partner has been isolated [48]. This novel serine/threonine protein kinase was named cyclin G-associated kinase (GAK). Immunoprecipitation/Western blotting experiments corroborated GAK binding to cyclin G and indicated that both proteins interact weakly with CDK2. Since such cell-based experiments do not rule out indirect association of proteins within a complex, Biacore’s SPR technology was used to conﬁrm that GAK binds directly to the N-terminus of cyclin G. The kinetic data provided by the Biacore analysis also showed that this association was stable. When the weak interaction between cyclin G and CDK2 suggested by immunoprecipitation was also investigated by SPR using recombinant proteins, no binding was detected, suggesting that any in vivo association between these two proteins either requires post-translational modiﬁcations, or that it is indirect. Biacore’s SPR technology therefore identiﬁed the primary molecular targets of cyclin G, providing the basis for future investigations into this system and its role in cancer. Cell cycle arrest and apoptosis are often demonstrably co-regulated. For example, survivin, a factor expressed during the G2/M phase of the cell cycle, co-localizes in centrosomes with the apoptotic protein, caspase-3. Although available data suggest that survivin allows cells to progress from G2 to M by binding caspase-3 and thus diverting cells from apoptosis, this has been difﬁcult to prove. Biacore was applied to ﬁnd out if survivin binds directly to caspase proteins and if this inhibits caspase function [49]. Survivin was immobilized on the sensor chip and was shown to bind caspase-3 and caspase-7 with high afﬁnity. In order to try to explain the discrepancies between Shin et al.’s caspase inhibition results and previous studies that failed to demonstrate binding, the authors noted that whereas their own experiments were carried out with all components pre-warmed to 25 ◦ C, the survivin used in other studies was kept on ice until immediately before the assay. Using the temperature control available with the instrument, it was demonstrated that compared to experiments carried out at 25 ◦ C, the afﬁnity of survivin for caspase-3 was reduced 100-fold at 4 ◦ C and may explain why the caspase:survivin interaction remained undetected in earlier assays. 3.3 Applications of Biacore in Clinical Research

Possibly the most attractive advantage of Biacore as a means of assessing binding partners in clinical research is the lack of a need for sample preparation or cleanup prior to running the experiment. The following examples are from reports in the literature on the analysis of clinical samples such as tissue homogenates, -plasma and urine. During the preclinical stage of the development of potentially therapeutic antibodies, promising alternatives are tested for their afﬁnity to the target [50, 51]. In one study, chimeric and humanized murine antibodies to CD44, a marker for head

3 Biacore Applications in Basic and Clinical Research

and neck cancer, were afﬁnity-ranked using Biacore and then assessed for biodistribution and efﬁcacy in mice bearing head-and-neck squamous cell carcinoma xenografts [52]. Based on both these sets of data, the important ﬁnding was made that lower afﬁnity antibodies to CD44 were superior to higher-afﬁnity versions in their ability to target tumors. While tumor uptake of low afﬁnity binders was up to 50 % higher than that of their high afﬁnity counterparts, blood levels and uptake into non-cancerous tissue was unaffected. Biacore has recently made its debut in the analysis of adverse events arising from the use of humanized antibodies in phase I clinical trials [53]. The monoclonal antibody, A33 recognizes a 43-kDa cell surface glycoprotein that is expressed in cancerous human colonic epithelium and which is absent from most normal tissues. A rapid and reliable means is needed to detect and quantify any host immune response to the humanized antibody which could possibly be harmful to the patient. A Biacore assay was used in which human immunoglobulin was immobilized on a sensor chip and whole patient serum was used as the analyte to detect any potentially harmful anti-human antibody response. This study showed that Biacore is sufﬁciently robust and consistent between sample runs to be used as an on-site method in clinical trials. The detection and measurement of antibody markers in human plasma is a common prognostic and diagnostic means of patient evaluation, as well as in monitoring clinical changes in response to therapy. A Biacore assay has been used to detect antibodies to gangliosides found in patients with neuropathies suspected of having an autoimmune etiology [54]. By immobilizing gangliosides on a sensor chip, crude diluted patient sera could then be used to detect the presence of antiganglioside autoantibodies. The assay was faster than ELISA partly due to the lack of sample preparation and further, had a superior limit of detection that could be even further improved through signal ampliﬁcation by using isotype-speciﬁc antibodies after serum injection (Figure 13). Several further examples exist of Biacore’s SPR technology in clinical evaluation including the detection of fucosylation of α1-Acid Glycoprotein (AGP), an acute phase event in inﬂammation. Antibodies in this case were ﬁrstly immobilized on a sensor chip to sequester AGP directly from human serum and the level of fucosylation was determined by the injection of a fucose-binding lectin [55]. The technique was used to follow the inﬂammatory status of severe burns patients. Other examples of the measurement of autoantibodies directly from patient plasma with little sample preparation include the detection of anti-DNA antibodies [56] and anti-protein S antibodies [57] in systemic lupus erythematosis and antifactor XII antibodies in patients with anti-phospholipid syndrome [58]. Biacore in combination with mass spectrometry (SPR-MS) has been shown in a proof-of-principle study to be applicable to the clinical situation [59]. Human urine or plasma were spiked with physiological levels of cystatin C, β2-microglobulin, urinary protein-1 and retinol-binding protein and injected over speciﬁc antibodies immobilized on a sensor chip. The captured analytes were then digested and identiﬁed by mass spectrometry. SPR-MS was used to detect simultaneously β2microglobulin and cystatin C in a single spiked plasma sample by injecting over a

27

28 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

Fig. 13 Use of secondary antibodies for identiﬁcation of the anti-GM1 autoantibody isotype. (A) Analysis of a serum sample from a patient with multifocal motor neuropathy with anti-IgM and anti-IgG antibodies, indicates that the antibodies in the

serum are IgM. (B) A similar analysis of a sample from a patient with Guillain–Barr´e syndrome indicates the presence of IgG antibodies. (Reprinted with the permission

of Dr Armin Alaedini, Department of Neurology & Neuroscience, Cornell University).

chip with different antibodies immobilized in two different ﬂow cells. Additionally the method was able to detect protein complexes such as that between retinolbinding protein (RBP) and transthyretin using an anti-RBP antibody. A mass spectrum from the chip surface showed traces of both species enabling detection of in vivo assembled protein complexes using SPR-MS. 3.4 Applications of Biacore in Neuroscience

Alzheimer’s disease is associated with the deposition of polymerized amyloid βpeptide (Aβ) protein in lesions in the brain parenchyma. However, Aβ protein is expressed by almost all nucleated cells and is not secreted at elevated levels in patients compared with healthy subjects. Why the protein should be deposited to form lesions exclusively in tissues of the central nervous system is unknown. The most common plasma proteins are also present in cerebrospinal ﬂuid (CSF), but

3 Biacore Applications in Basic and Clinical Research

at much lower concentrations. Biacore’s SPR technology was used in a screening strategy to ﬁnd out whether any proteins common to both plasma and CSF could bind Aβ protein and thus potentially inhibit deposition [60]. Two forms of Aβ protein are known; one with 40 amino acids and a longer variant that contains two extra C-terminal residues (Aβ1–42). The latter polymerizes more readily into plaque-forming ﬁbrils called β-amyloid. Two parallel immobilization strategies were followed to monitor the binding of plasma/CSF proteins to both forms of Aβ protein. Plasma and CSF proteins were tested for their capacity to inhibit Aβ protein polymerization in a microtitre assay in which Aβ protein or Aβ1–42 were immobilized as templates to which further Aβ protein was added in the presence of the test protein. This revealed that the most inhibitory substances were albumin, α1-antitrypsin, IgA and IgG, proteins also found at a lower concentration in CSF. In Biacore experiments, albumin was run over ﬂow cells prepared as indicated above and was shown to bind only to immobilized, polymerized Aβ protein. The authors suggest that albumin, at concentrations found in plasma, may bind and prevent the deposition of polymerized Aβ protein. In inﬂammation, levels of albumin in CSF may be sufﬁciently reduced to abrogate this protective function, enabling the formation of β-amyloid. In addition to Aβ protein, serum amyloid P component (SAP) is also found in amyloid deposits in Alzheimer’s disease and, like Aβ protein, tends to selfassociate in the presence of calcium. Calumenin is one of a family of endoplasmic reticulum proteins that coordinate calcium in maintaining the structural integrity of helix–loop–helix motifs in certain proteins. One study elegantly demonstrated how Biacore could be used to deﬁne the interaction and characterize the conditions necessary for it to occur [61]. An unidentiﬁed binding partner for calumenin was ﬁrst detected when a crude placental extract was run over recombinant calumenin immobilized on an afﬁnity column in the presence of calcium. After elution and PAGE, one 30-kDa band was isolated and the protein was internally sequenced by tryptic digestion and identiﬁed as SAP. Calumenin was then immobilized as a ligand on a sensor chip. The results conﬁrm that SAP is a ligand for calumenin and that the formation and stability of the complex requires calcium. It is therefore possible that given the tendency of both SAP and calumenin to form insoluble complexes, calumenin may participate in amyloidosis, the pathological process by which amyloid is deposited in the CNS of patients with Alzheimer’s disease. Spinal muscular atrophy (SMA) is a disease that is unequivocally linked to a gene called survival of motor neurons (SMN) [62, 63]. The disease is severely disabling and is characterized by the loss of motor neurons from the lower spinal cord. There is a strong correlation between the severity of SMA and levels of SMN protein. The function of SMN protein is unclear although it is thought to be involved in mRNA splicing. To determine a possible mechanism to explain why mis-sense mutations in SMN should compromise protein function and hence cause disease, Biacore was used to discover how native peptide fragments of SMN, deﬁned by whole, discrete exons, interact with each other [64]. To identify and select the tools necessary to map these protein interactions, a panel of monoclonal antibodies was raised to full-length SMN protein. The antibodies were then tested using Biacore’s SPR

29

30 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

technology for reactivity against the peptides encoded by each exon in the cloned gene. The antibodies were immobilized on a sensor chip and screened with all the fragments. Antibodies were thereby identiﬁed which speciﬁcally recognized certain peptides within the protein and which could then be used to characterize intra-chain interactions in SMN that may determine conformational settings of functional importance in the full-length protein. By then immobilizing each peptide fragment and screening with exon-deﬁned peptides, the authors deﬁned the fragments of SMN which possessed the potential to interact with each other. This information, combined with epitope mapping, enabled an experiment to be constructed which gave a wealth of information about the structural intramolecular events behind the function of a protein. One monoclonal antibody that was shown to recognize the peptide product of exon “A” alone was immobilized on a sensor chip and subsequent injections of different peptides showed that the capture of peptide A enabled peptides “B1” and “B2” to bind sequentially. These data show not only that peptide A appears to form a scaffold to allow the assembly of B1 and B2, but that the functional protein product is a result of the creation of a novel epitope formed by the combination of B1 and B2. This study is recommended as a model of how Biacore’s SPR technology can be used to test the molecular mechanisms underlying the expression of disease. Alzheimer’s disease is a neurodegenerative disease in which Aβ protein participates in neurotoxic aggregates which localize to sites of cell death and neuronal apoptosis. Toxicity may also be caused by the localized release of activated inﬂammatory cytokines in the formation of so-called senile plaques. Aβ protein has been shown to induce the release of pro-inﬂammatory cytokines by cultured cells and this function was shown to be inhibited by ganglioside GM1 [65]. Biacore’s SPR technology was used to demonstrate direct binding of Aβ protein and liposomeimmobilized GM1, suggesting the mechanism behind this neuropathologic function. Interaction was shown to depend on the presence of sialic acid. As Aβ protein is known to be neurotoxic in a β-pleated conformation, the authors suggest that one function of GM1 may be to maintain Aβ protein in an inactive state and thus prevent the initiation of the inﬂammatory response of neurons in Alzheimer’s disease. In a modiﬁed approach to the liposome capture strategy of Ariga and Yu [65] described above, liposomes were immobilized on a sensor chip via an antibody [16]. In addition to ganglioside GM1, lipopolysaccharide (LPA) was incorporated into the liposome construction in this study, enabling capture by an anti-LPA antibody and mild regeneration conditions. A rigorous investigation into the immobilization of gangliosides has been carried out in an effort to develop a simple assay for anti-ganglioside antibodies [66]. A requirement of the technique is to present the glycolipid in an appropriate orientation for antibody characterization i.e. in a manner in which the sugar moiety presented in tumor cells in vivo is available to the analyte and which is thus a potential target for therapeutic monoclonal antibodies. The gangliosides were immobilized via the lipid tail, as using the more common aldehyde coupling procedure via the carbohydrate domain masks the epitope. The authors present a simple and rapid protocol, yielding a stable surface that can be readily reconditioned and reused. Gangliosides

3 Biacore Applications in Basic and Clinical Research

were injected over the unmodiﬁed surface of a sensor chip, with ganglioside GD3 covalently immobilized by photoactivation for comparison. Although overall immobilization levels were low compared with those achieved by covalent linkage, the molar level of binding (speciﬁc binding) was much higher. Phase I clinical trials, in which chimeric anti-ganglioside antibodies were tested in children with neuroblastomas [67], have shown that a host immune response is still induced to the murine component of the antibody. For this reason, engineered humanized antibodies wele synthesizd that retain the murine speciﬁcity-determining CDR regions, but from which the ﬂanking V-region sequences have been replaced with human counterparts. This produces a theoretically less immunogenic protein in which the murine component is reduced from 30 to 10 %. Nakamura et al. [67] based the characterization of the interaction between these humanized antibodies and immobilized gangliosides GD2 and GD3 on the protocol of Catimel et al. [66] above. Their Biacore experiments revealed that humanization changed the kinetic characteristics of the antibodies in ways that are otherwise difﬁcult if not impossible to predict. Biacore’s SPR technology thus provides information about the molecular mechanisms underlying functional changes in humanized antibodies, a critical part of the development of novel antibodies for human immunotherapy. Nerve growth factor (NGF), a member of the neurotrophin family, is implicated in the differentiation and survival functions of neurons [68, 69]. Biacore’s SPR technology has been used to characterize many types of receptor:ligand interactions, including that between NGF and the Trk family of receptors containing intracellular tyrosine kinase domains. Shc is a so-called adaptor protein comprising a C-terminal SH2 domain and an N-terminal phosphotyrosine-binding (PTB) domain. One of its known functions is to link ligand-activated growth factor receptors, including the human NGF receptor TrkA, to the Ras signaling pathway. This functions through its participation in the multi-molecular signaling complex, Shc-Grb2-Sos. The SH2 domain recognizes phosphotyrosine indirectly via adjacent C-terminal amino acids, while the PTB domain directly binds phosphotyrosine located within a speciﬁc amino acid motif. Biacore’s SPR technology was used to study the speciﬁcity and kinetics of the interaction between the PTB domain of Drosophila Shc and phosphopeptides derived from an EGF-receptor homolog, containing structural features shared by other receptors including TrkA, such as conserved hydrophobic amino acid residues [70]. This system is therefore likely to be a suitable model for other receptor:ligand interactions. Initial experiments in which a peptide containing phosphotyrosine ﬂanked by 11 N-terminal and ﬁve C-terminal residues was immobilized on a sensor chip, showed that the association and dissociation rates of the Shc PTB domain were somewhat slower than typical, previously determined SH2 interactions. This may imply that in the interaction between Shc and an activated receptor, Shc is likely to bind rapidly to its target via its SH2 domain while its PTB domain limits the rate of dissociation. By measuring the ability of truncated or point-mutated phosphopeptides to inhibit the binding of the Shc PTB domain to native phosphopeptide, the speciﬁcity of the target sequence surrounding phosphotyrosine in the growth factor receptor was determined in a Biacore competition assay. Phosphopeptides were

31

32 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

truncated at various positions N-terminal to phosphotyrosine, incubated in solution with Shc and passed over native phosphopeptide immobilized on a sensor chip. C-terminal deletions had little effect, but deleting N-terminal amino acids revealed that the presence of hydrophobic residues at positions −5 and −7 were critical for Shc to effectively bind the receptor. As expected, nanomolar concentrations of unmodiﬁed peptide efﬁciently inhibited binding of Shc while an interaction could still be detected even in the presence of high concentrations of deletion-peptides. In a study of some relevance to Biacore assays in the receptor:ligand interaction ﬁeld in general, the receptor for mammalian TrkA was characterized [71] in which the extracellular ligand-binding domain of TrkA was over-expressed in insect cells, puriﬁed and immobilized on a sensor chip. The association rate of NGF-binding to recombinant TrkA extracellular domain was close to previously reported values using the full-length receptor. These results suggest that recombinant, puriﬁed receptors may be used as a model in the analysis of NGF:TrkA interactions using Biacore’s SPR technology and that the strategy may be generally applicable to a wider range of receptor:ligand interactions. The physico-chemical composition of the microenvironment of the binding site on a receptor for its cognate ligand is critical. Ionic strength and pH, for example, affect non-covalent bonds and so by varying the buffer composition, judgments can be made on the nature of the interaction. Biacore’s SPR technology was used to discover whether local variations in pH changed the capacity of Zn2+ and Cu2+ to modulate NGF binding to its receptor [72]. To do this, the cations were included in running buffers with pH varying from 5.5 to 7.4, mimicking local conditions in cerebral acidosis, a condition frequently arising after stroke or traumatic insult. This pH range also covers the pKa value of histidine, the position and electrostatic status of which are known to inﬂuence cation binding. The authors used the strategy described by Woo et al. [71] to prepare and immobilize recombinant TrkA. The dissociation equilibrium constant, KD, for the NGF:TrkA interaction was calculated over a range of pH values covering the pKa value of histidine. This revealed that Zn2+ , but not Cu2+ , lost its ability to bind NGF and inhibit its interaction with TrkA under acidic conditions. The physiological consequences of this depend on the cell type and context; TrkA-expressing cells in an acidic environment and needing NGF for survival may well beneﬁt from the inactivation of Zn2+ , whereas the effects might be detrimental if NGF, in contrast, initiates signals leading to cell death. NGF functions both within and outside the nervous system, for example it is released from mast cells during allergic inﬂammation and is implicated in the pathophysiology of asthma. Using in vitro studies it was shown that a novel protein called FIZZ1, isolated from the lavage ﬂuid of inﬂamed murine bronchoalveoli, dampened NGF-mediated survival and gene expression in neurons [73]. Biacore was used to show that an antagonistic mechanism, which is not based on competition for a single receptor, operates to inhibit NGF function in TrkA-bearing cells. The data suggested that rather than inhibiting the interaction of NGF with its receptor, FIZZ1 functions indirectly by binding to a different and as yet unknown receptor, possibly regulating TrkA activity by “inside-to-out” signaling.

3 Biacore Applications in Basic and Clinical Research

The rationale for one Biacore study on Alzheimer’s disease [74] was to identify small peptides that can inhibit aggregation and therefore the toxic neurodegenerative effects of β-amyloid (Aβ) in the diseased brain. Although Aβ aggregates are present in neurotoxic lesions, it is not known if they are causative. Short peptides directed to the self-recognition motifs in Aβ were synthesized and were used to screen the afﬁnities of the interactions. Solution-based methods for screening Aβ inhibitors consume large amounts of the target and they are limited by the tendency of Aβ to aggregate in solution and thus present multiple target forms. This problem is avoided in Biacore assays because the target is immobilized on a planar sensor chip surface at a density and in an orientation that can be carefully controlled by the user. A 26-amino acid fragment of Aβ comprising amino acids 10 to 35 with a Cterminal cysteine was immobilized at low density on a sensor chip thus presenting a uniform surface. Further, inter-assay variability was low as a single chip surface could be regenerated several times and re-used for all test interactions. A range of concentrations of each peptide, based on variations of a ﬁve-amino acid sequence corresponding to the central hydrophobic domain of Aβ known to be responsible for self-association, was injected over Aβ on the chip surface for 500 s and the response at equilibrium (Req ) was plotted against time. As the binding was weak, only relative afﬁnities were measured, with the equilibrium response determined after 90 % of the contact time at each concentration. Of several peptides based on multiple length C-terminal extensions of a pentapeptide in the central hydrophobic domain of Aβ (which generally bound more strongly than the original pentapeptide to Aβ), a pattern emerged in which the afﬁnity for Aβ was sensitive to site-speciﬁc positioning of positively-charged lysine residues. For example, it was previously determined that the addition of six lysine residues directly adjacent to the basic KLVFF sequence generated a more effective inhibitor of cytotoxicity in cell-based assays. In the Biacore assay, peptides in which three of these residues were separated from the basic sequence by three negativelycharged residues had a 14-fold lower afﬁnity for Aβ, while the afﬁnity of a peptide of similar size bearing three directly adjacent lysine residues was almost unchanged. The peptides were then tested in cytotoxicity microtitre assays by incubating Aβ with neuroblastoma cells in the presence of peptides and compared with the Biacore afﬁnity data (Table 1). High afﬁnity binders protected against toxicity by greater than 80 % in viability assays while low afﬁnity binders were less effective. It is hoped that these peptides will be useful as probes of the mechanisms underlying amyloid plaque formation and in the design of drugs. Biacore’s SPR technology, therefore, is proposed as a reliable means to measure the afﬁnity of small peptides for Aβ, a protein which is notoriously difﬁcult to handle in solution-based assays. Such data may help predict the suitability of candidate peptides as inhibitors of Aβ toxicity in Alzheimer’s disease. The glutamate receptor δ2 (GluRδ2) [75, 76] is exclusively expressed on the dendritic synapses of Purkinje cells [77], structurally unique cells that are among the most organizationally complex neurons in the mammalian nervous system. Although they receive an enormous amount of data from neurons of the spinal cord and elsewhere, they are nevertheless able to convert this input into interpretable

33

34 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors Table 1 Peptides based on C-terminal extensions to a central KLVFF fragment derived from

the central hydrophobic domain of Aβ were tested for their afﬁnity to Aβ immobilized on Pioneer Chip B1. Afﬁnity (KD ) is related to the ability of the same peptide to inhibit Aβmediated cell cytotoxicity in a cell-based assay Peptide sequence*

KD (l M)

Viability (%)+

KLVFFRRRRRR KLVFFKKKKKK KLVWWKKKKKK KLVFFKKKK KLVFWKKKKKK KLVFFKK KLVFFKKKEEE KKKKLVFF KLVFFEKEKEK KKKKKK KLVFFEEEKKK

40 40 40 37 65 80 90 180 300 400 1300

> 90 80–90 80–90 70–80 70–80 70–80 60–70† 60–70† 60–70† 60–70† 60–70†

+ Cellular viability of human neuroblastomas cells was assessed in a microtiter assay using an MTT assay in which cells were exposed to Aβ in the presence or absence of peptides. ∗ K, lysine; L, leucine; V, valine; F, phenylalanine; R, arginine; W, tryptophan; E, glutamic acid. † This is a similar level of viability as seen after treatment with negative controls.

information and are, in fact, the sole means of output in the cerebellar cortex. That GluRδ2 is limited to these cells possibly implies a function in this extraordinary feat of information processing. Biacore analysis showed that an intracellular protein called delphilin co-localizes with GluRδ2 at parallel ﬁber (input)–Purkinje cell (output) synapses [78] and that recombinant delphilin PDZ domain binds to the C terminus of GluRδ2 with moderate afﬁnity, mostly accounted for by rapid dissociation. The authors also showed that delphilin bound proﬁlin, an actin-binding protein, and dissociated very slowly. It is possible therefore that delphilin acts as a molecular bridge linking GluRδ2 to the actin cytoskeleton via proﬁlin with rapid turnover of the GluRδ2:delphilin interaction contributing to the capacity of the Purkinje cell to process a very large amount of input data. Spinal muscular atrophy (SMA) is predominantly a childhood disease of motor neurons in the spinal cord, affecting the voluntary muscles and causing the impediment of functions such as locomotion and head and neck control. The disease is strongly linked to mutations in the SMN1 (survival motor neuron 1) gene. It has recently been suggested that the clinical symptoms of the disease may be due to increased motor neuron death through the inability of mutated SMN protein to dimerize, and thereby sequester and inhibit the pro-apoptotic function of the tumor suppressor protein, p53 [79]. An elegant surface plasmon resonance strategy was followed in which Biacore was used initially to conﬁrm conventional immunoprecipitation (i.p.) data using an antibody to p53 showing that SMN associated with p53. This ﬁnding was substantiated using Biacore in sequential binding experiments that demonstrated how the technology might be used as an alternative to immunoprecipitations and may represent a more convenient tool for detecting molecular interactions in whole cell

3 Biacore Applications in Basic and Clinical Research

Fig. 14 Comparison between immunoprecipitation/blotting and Biacore in the detection of molecular complexes. Both assays reveal that SMN protein derived from whole cell extracts is associated with p53 in the cell. Results achieved by immunoprecipitation are available within 1 day whereas the Biacore assay takes minutes.

extracts (Figure 14). Brieﬂy, anti-p53 antibody was coupled to a sensor chip. Next, whole cell extracts from SMN-transfected cells (treated with a protease inhibitor that protects the labile p53) were passed over the chip, followed by exposure to anti-SMN antibody. Cell extracts that did not contain detectable levels of p53 were used in parallel and demonstrated the speciﬁcity of the interaction. SMA patient-derived mutations were also tested for p53 binding. Two SMN proteins carrying C-terminal mutations found in patients with severe type I SMA and a peptide from a patient with less severe type II SMA were immobilized on a sensor chip and exposed to p53. The magnitude in reduction of SMN peptide binding strongly correlated with the clinical subset. This defect is likely to be attributable to the inability of the SMN mutant proteins to efﬁciently self-associate. Young et al. [79] then applied the sequential Biacore approach to show that this very inability to self-associate may prevent SMN from sequestering and inhibiting p53 function. In these experiments, p53 failed to bind to immobilized monomeric SMN and was only induced to bind when the same chip was supplemented with dimeric SMN.

35

36 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

The signiﬁcance of these ﬁndings is supported in the same body of work by results from indirect confocal immunoﬂuorescence experiments showing that SMN and p53 co-localize in the Cajal bodies (sites of small nuclear ribonucleoprotein particles involved in RNA splicing) of normal ﬁbroblasts, but not in those derived from SMA patients. 3.5 Drug: Plasma Protein Interactions

The clinical potential of drug compounds is greatly affected by the nature of their interactions with circulating plasma proteins, such as human serum albumin (HSA) and α1-acid glycoprotein (AGP). The effects of plasma protein binding by drugs are varied and can inﬂuence factors such as the free concentration of drug compound in the circulation, transport and distribution around the body and the duration of drug action. Biacore offers the user the ability to run rapid, automated assays with high sensitivity and relatively low sample consumption. The sensitivity of the technology varies from system to system but association rate constants (ka ) of up to 5 × 106 M−1 s−1 and dissociation rate constants (kd ) of between 10−4 and 1 s−1 are accurately and reproducibly attainable with the latest available systems. Lead compounds can be analyzed from sub-nanomolar to millimolar concentrations. R Biacore’s latest model, Biacore S51, is speciﬁcally designed for drug discovery applications from simple ranking, to detailed characterization of the interactions. Hit selection assays may be employed for ranking of lead compounds for binding to one or two plasma proteins, with a throughput of 96 compounds in 12 h. The data produced from single concentration analyses permit the ranking of compounds in terms of binding response to single or multiple plasma proteins. Based on this ranking assay, compounds for which a high-resolution analysis is required can then be selected. These are then taken for detailed compound characterization assays which use concentration series of compounds to generate data sets from which KD values can be calculated using appropriate ﬁtting models. The KD values may then be transformed to give percentage-bound ﬁgures, which can be calculated for single or combined plasma protein binding, as appropriate. It may also be possible to maximize the level of the compound characterization to include kinetic analyses of drug–plasma protein interactions in some cases. 3.6 Drug:DNA Interactions

A number of drugs bind cellular DNA and inhibit targeted protein–DNA interactions. A prerequisite for the design of new types of compounds is rapid penetration of cells and interaction with a speciﬁc sequence of DNA. Such compounds, structurally predicted from the gene sequence and functionally tested using Biacore’s SPR technology, could have many therapeutic and biotechnological uses. Relatively small molecules can be remarkably speciﬁc in their sequence recognition. A “prodrug” of furamidine, for example, binds strongly to the minor groove of DNA and

3 Biacore Applications in Basic and Clinical Research

is selective for AT-rich sequences [80]. Biacore has been used in combination with DNA footprinting methods [81], which are employed to quickly characterize compound interactions with deﬁned sequences of various sizes to discover new DNAbinding drugs. Biacore is then used to resolve questions of stoichiometry, relative afﬁnities and binding kinetics. The characterization of DNA sequence recognition and DNA binding of indolocarbazole antitumor drugs is described elegantly in two studies [81, 82]. Thermodynamic characteristics of binding of organic cations to AT and mixed sequence DNA sites are shown by isothermal calorimetry and Biacore to be very dependent on structure, solvation and sequence of the DNA binding site (see also [83] and [84]). Recently Biacore was shown to be a rapid and reliable means to detect DNA base pair mismatches by using a synthetic low molecular weight ligand that speciﬁcally binds G:G mismatches. This work has far-reaching implications for SPR in genome analysis [85]. The technique depends on the development of reagents that speciﬁcally recognize certain types of aberrations in native DNA. Today, single nucleotide polymorphisms (SNPs) are routinely detected using methods that rely largely on the separation of DNA fragments by gel electrophoresis or HPLC, neither of which are readily suited to high-throughput analysis. The sheer volume of sequence information from the human genome project demands the development of a tool for rapidly scanning DNA for SNPs and Sando et al. [85] have used Biacore’s SPR technology as an alternative means to this end. They designed and synthesized a low molecular weight ligand that speciﬁcally binds double-stranded DNA containing G:G mismatches arising from native C:G pairings and which has minimal reactivity with any of the other seven possible mismatch combinations (A:A, A:C, A:G, T:T, T:C, T:G or C:C). The ligand, naphthyridine, is modeled to recognize the characteristic open bulge that is created between DNA strands at the site of a G:G mismatch, at which no complementary base (cytosine) is present with which to form a hydrogen-bonded base pair. Naphthyridine is synthesized as a dimer and is therefore able to form simultaneous hydrogen bonds with both guanines on each DNA strand. Experiments have shown that the melting temperature, i.e. that at which half of the double-stranded DNA in a sample dissociates into single chains and which is a measure of DNA stability, increased in the presence of dimeric, but not monomeric naphthyridine, to a much higher threshold than that of native DNA or DNA of an otherwise identical composition containing other mismatches. The synthetic naphthyridine ligand contains an aminoalkyl chain and can therefore be covalently linked by amine coupling to carboxyl groups on a sensor chip. 27-mer doublestranded DNA was passed over the prepared chips and binding was seen only if the DNA contained G:G mismatches. DNA containing other SNPs or native DNA did not bind to the ligand. The sensitivity of the assay was shown by a linear increase in response with concentration of analyte from 125 to 1000 nM, a range that was optimized by changes in salt concentration. The intensity of response was a function of both DNA concentration and length. Other non-gel based mismatch screening methods such as melting curve analysis or MutS capture of mismatched fragments rely on detection systems such as

37

38 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

labeling with a ﬂuorescent dye or the use of labeled primers. Sando et al. [85] showed that Biacore’s SPR technology is sensitive, speciﬁc, requires no primers or probes and can be adapted for high throughput analysis. This is the ﬁrst report in which Biacore has been used for DNA mutation analysis and while the design and synthesis of ligands that are speciﬁc for other SNPs will be difﬁcult, the principle of Sando et al.’s work is a promising development in accelerating DNA mutation analysis (see also [86] for a commentary on this exciting work). One further study shows that a second class of small molecule, pyrrole-imidazole-containing polyamides, can also function as DNA mismatch recognition agents [87]. 3.7 Nucleic Acid Structure and Analysis

In addition to identifying prospective binders to DNA in the search for novel drugs, Biacore has also been used to investigate the effects of deliberate alterations in the structure of nucleic acids on their function. For example, it has been shown that the substitution of the sugar-phosphodiester backbone of DNA to create so-called pyrrolidine-amide oligonucleotide mimics (POM) [88] enabled the synthetic singlestranded DNA analogs to bind to complementary native strands with higher afﬁnity than did other native strands. Additionally it was shown that POMs have a much faster association rate for binding to RNA than for binding to DNA. Simple cations and polyamines are also shown to affect the binding afﬁnity of ssDNA in duplex formation and the rate of DNA duplex formation and dissociation. Afﬁnity and kinetic analyses have been performed on several different types of DNA structures including perfect linear matches, bulged loops, mismatched duplexes and hairpin duplexes. Of possibly critical importance in cancer, Biacore studies have shown that polyamines (commonly found in tumor cells) may stabilize mismatched duplexes via increasing the thermal stability of the duplex and causing a reduction in kd [89]. Protein:DNA interactions have also been tested by immobilizing a range of different structural forms of DNA on a sensor chip surface and testing afﬁnity and kinetic parameters with interacting proteins [90]. This conceptually interesting work shows that DNA polymerase binds quantitatively and kinetically more efﬁciently to primed DNA than to bluntend dimers. “Morpholinos” (MORFs) are a recently developed class of synthetic nucleotide oligomers that carry nucleotides in the same deﬁned sequences as their parent nucleic acid but in which the sugar backbone is replaced with a phosphorodiamidate backbone [91]. MORFs are reported to be more nuclease-resistant, more soluble and more rigid than native DNA and thus potentially may be highly efﬁcacious as carriers of sequence-targeted radiopharmaceuticals in cancer treatment [92]. Anti-sense DNA is a frequently investigated strategy aimed at the targeted inhibition of the translation of oncogenic protein genes in vivo. The efﬁcacy of such treatment, however, is limited by the difﬁculty of ensuring delivery of antisense oligonucleotides into cells. Anti-sense oligonucleotides were conjugated with cell-penetrating peptides and Biacore was used to conﬁrm that the binding afﬁnities of these oligonucleotides for their targets after conjugation were similar to unconjugated oligonucleotides [93]. An extension of the anti-sense approach is a strategy

3 Biacore Applications in Basic and Clinical Research

in which double-stranded DNA is targeted, the so-called anti-gene approach. Here, a DNA triple helix is transiently formed between a speciﬁc dsDNA and a singlestranded oligonucleotide and which may potentially inhibit transcription of speciﬁc genes. These transient structures, however, are unstable in vivo and Biacore was used in one study to monitor the stability of the complexes formed between dsDNA and chemically-stabilized oligonucleotides [94]. Interactions between small deoxyribozymes and RNA are also open to Biacore on-chip analysis. In one interesting study, the role of higher order RNA structure in self-association has been investigated [95]. The authors propose that mRNA transcribed from mutated genes can form structural anomalies that are regulatory parameters in themselves i.e. that the shape of the RNA in addition to its sequence is important. The implication is that even if a mutation in a codon does not result in an amino acid change in the translated protein, the structural alteration of the transcript may nevertheless have functional consequences. Accurate mRNA splicing for example, is sensitive to the maintenance of RNA structure in order to allow interactions with other cellular components. The authors immobilized short sequences of RNA which recognized and cleaved RNA sequences containing a loop resulting from one such mutation. The cleavage function was shown to be both loop size- and sequence-dependent and further, the reaction could be monitored in real time on the chip surface. Most real-time “on-chip” reactions have been concerned with monitoring realtime changes in molecular weight of an immobilized substrate during passage of an enzyme analyte although DNA synthesis during the passage of DNA polymerase has also been studied [96]. For selected reading on monitoring enzyme activity on Biacore chips through real-time alterations of substrate molecular weight, see also [97] (Design of helical proteins for real-time endoprotease assays); [98] (Proteolysis of the exodomain of recombinant protease-activated receptors: prediction of receptor activation or inactivation by MALDI mass spectrometry); [99] (Monitoring of the refolding process for immobilized ﬁreﬂy luciferase with a biosensor based on surface plasmon resonance); and [100] (Peptide chips for the quantitative evaluation of protein kinase activity). 3.8 Protein:RNA Interactions

The association of speciﬁc proteins with RNA molecules begins immediately as the nascent RNA chain is produced during transcription. From this point onwards, a series of different proteins are bound to and released from the RNA and are involved in co- and post-transcriptional RNA processing events such as splicing, export to the cytoplasm, regulation of RNA stability and translation. These interactions are highly dynamic and are critical in the regulation of the various steps involved in RNA processing. In addition, these individual RNA:protein interactions occur in the environment of extremely large and complex nucleic acid–protein complexes such as the transcriptosome and spliceosome. Until recently, however, studies on RNA-binding proteins have relied heavily on techniques which examine binding under equilibrium conditions (such as electrophoretic mobility shift assays). Although this type of

39

40 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

afﬁnity-based data is valuable in many instances, it does not address the key issue of the dynamics of the RNA:protein interactions. This requires a true kinetic analysis that can deliver individual association and dissociation rate constants (ka and kd ). Biacore’s SPR technology can be used to provide detailed kinetic data on the interaction of RNA-binding proteins with their speciﬁc RNA targets and this information can provide invaluable insights into their functions. In one study, detailed kinetic analysis was used to elucidate a novel two-stage binding interaction between the spliceosomal protein, U1A and its RNA target [6]. Although the range of surface chemistry approaches that are compatible with Biacore’s technology enables great ﬂexibility in selecting which interaction partner should be immobilized on the sensor surface, the nucleic acid is most often immobilized in RNA:protein studies. One reason for this is the relative ease with which RNA oligonucleotides can be chemically synthesized with a terminal biotin modiﬁcation which enables them to be immobilized in an oriented fashion on a commercially-available streptavidin surface (Sensor Chip SA). In one particular study on binding of the neuronal protein, HuD to RNA recognition motifs (RRMs), individual RRMs that had remained “hidden” during previous equilibrium analysis-based studies were shown to make important kinetic contributions to the overall binding function (see [22] and [101]). Many RNA-binding proteins contain multiple RRM regions and the results from the HuD study may therefore represent a general model for how investigation into this type of protein should be approached. In the same study, the analysis of spliceosomal protein U1A binding to its target RNA revealed a hitherto unknown two-stage mechanism of binding. This data was also able to identify key amino acid and nucleotide residues involved in the individual interaction stages and provided evidence for the types of forces involved. This model may also prove to have a much wider relevance, since a number of other RNA-binding proteins are known to contain similar distributions of basic amino acids around their RNA binding sites. In both cases, crucial new information regarding the mechanism of the protein:RNA interaction was obtained. These studies relied heavily on the ability of Biacore’s SPR technology to provide high-quality kinetic data. In addition to furthering our understanding of the many vital biological interactions between proteins and RNAs, kinetic approaches may also be invaluable in designing therapeutic small molecule inhibitors.

4 Current Developments and Future Perspectives 4.1 SPR-MS

The advent of the proteomics era has placed a new emphasis on techniques that enable the identiﬁcation of proteins. Of these, mass spectrometry has been the most signiﬁcant, with MALDI-TOF (matrix-assisted laser desorption/ionizationtime of ﬂight) and ESI (electrospray ionization) approaches dominating the ﬁeld. MS analysis is commonly used in combination with traditional separation

4 Current Developments and Future Perspectives 41

techniques, predominantly two-dimensional gel electrophoresis, in order to combine expression proﬁling with protein identiﬁcation. In contrast to the array technology approaches used in expression proﬁling at the mRNA level, analogous developments within proteomics will be greatly complicated by the extensive and complex interactions among the proteins expressed. In practice, this means that simple expression data will be of extremely limited value without a detailed understanding of the expressed proteins’ interaction partners. Combining SPR with MS would therefore provide a potentially powerful marriage of technologies within the proteomics ﬁeld, with the possibility of providing protein identiﬁcation which is selected on the basis of functional binding criteria. The painful lesson learned from previous genomic studies is that the expression of a gene in a particular biological context can often prove to be a meaningless observation in itself in the absence of direct functional data. A number of approaches to combining SPR and MS have been taken thus far. It is possible, for example, to physically remove the sensor chip from a Biacore instrument following binding of the analyte to the ligand and use it directly in a MALDI-TOF analysis [102, 103]. This approach produces minimal analyte loss, since all of the analyte molecules are bound within the area of the ﬂow cell which is used as the MALDI target. An alternative method is to elute captured analyte from the sensor surface, using a procedure designed to recover the sample in a small volume after SPR analysis [59, 104]. In addition to being able to use the same sensor surface for repeated recoveries, another advantage of this approach is that it is also possible to enzymatically digest the bound analyte on the sensor surface prior to or following recovery [105], which greatly facilitates MS identiﬁcation by allowing alignment of MS proﬁles against various databases. A disadvantage to this approach is the risk of contamination, which must always be guarded against. The SPR route into MS analysis holds a great deal of promise, especially in the ﬁeld of interaction proteomics, and as testiﬁed by the recently published work just described, much has already been achieved. Future developments will doubtless further improve the efﬁciency and automation of the technique. The small sample volume requirement and high sensitivity features of SPR technology, which are major advantages in terms of interaction analysis, mean that the amount of material that it is possible to obtain from a single SPR experiment for MS analysis remains somewhat restricted. This reﬂects the fact that the design of SPR instruments has hitherto been optimized very much from the viewpoint of the interaction analysis itself, rather than placing any emphasis on preparative purposes. As SPRMS continues to be developed as a combined technology concept however, it can be anticipated that the two constituent technologies will adapt to each other to provide an optimal solution. 4.2 The Challenges of High-throughput Protein Analysis and Array Technologies

The range of currently available technologies for studying proteins all appear to be restricted in the throughput that can be easily achieved. This suggests that the main

42 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

challenge for the next phase of development is principally one of scale. As interest in proteins shifts gradually away from considering isolated or restricted groups of molecules and more towards understanding how multiple proteins interact in complex, functionally interrelated networks, it is inevitable that increasingly higher throughput systems will be desirable. While protein microarray systems will certainly be developed over the next few years, it is worth considering some of the realities relating to the challenges involved. It would be optimistic to imagine that in the near future proteomics will be conducted on a genomics-scale, since the complexity of working with proteins as compared to nucleic acids presents more of a challenge for developments in automation. There are no current proteomics technologies analogous in performance to the PCR, high-throughput sequencing and SNP analysis technologies which have enabled such astonishingly rapid progress in genomics-related disciplines. Protein array chips will certainly be important tools in proteomics research, but the scale with which they will be used will depend on the information they provide. The word “array” may itself have very different meanings and interpretations. To the genomics ﬁeld, familiar with chip-based cDNA arrays for expression proﬁling for example, “array” conjures up an expectation of anything from several hundred to many tens of thousands of individual DNA sequences on a single analysis surface. Despite the highly speciﬁc nucleic acid sequence hybridization principle on which such systems are based and the simple, semi-quantitative nature of the data output, this type of array technology experienced difﬁculties in terms of reliability, reproducibility and not least, data analysis. The challenges for large-scale protein arrays are likely to be considerably more difﬁcult than for their DNA counterparts, with complications such as the difﬁculties in expressing and purifying large numbers of correctly folded proteins with the appropriate post-translational modiﬁcations, protein instability, much greater potential for cross-reactivity, greater difﬁculty in standardizing binding conditions and much more complex data handling. These fundamental challenges must be dealt with in some way by all the technologies that hope to contribute to the development of protein array approaches. In practice, the development of protein array chips will almost certainly involve a trade-off between relatively low-information screening and more detailed analysis approaches, in terms of the total number of simultaneously analyzable protein “spots” that are practical to use. Protein arrays will doubtless be applied in both the areas of “discovery proteomics” and “interaction proteomics” (see Section 1.1), but are likely to be adapted in quite different ways to meet these distinct challenges. While it may be possible for example to spot many thousands of proteins for semiquantitative expression proﬁling applications, it is highly unlikely that high-quality kinetic analysis of protein interactions will be carried out on the same scale in the foreseeable future (or even that there would be a demand for such an application). The nature and size of protein microarrays may vary greatly therefore, depending on the intended application. Currently, it is possible to simultaneously analyze a maximum of four immobilized proteins using Biacore’s SPR systems and this can be viewed as a very small-scale microarray system. The strengths of this technology are largely targeted towards producing high-quality, information-rich data (such

References

as high resolution kinetics) and this may position it more towards the high-data content, rather than ultra-high throughput end of the protein array scale. It is important to bear in mind that each data “spot” in a Biacore assay produces a real-time analysis of its interaction with the analyte over the entire course of binding and dissociation. In high-resolution kinetics analysis, this may involve SPR measurements taken at a rate of up to 10 s−1 over the course of many minutes. The amount of data produced by each interaction monitored is therefore very high and while this is one of the cornerstones of the success of Biacore’s technology in a broad range of different applications, it is something that must be considered in the context of larger array situations. As was experienced by the pioneers of large-scale DNA arrays, it may be that data analysis will provide a more important practical limitation to the scope of protein arrays than the developments in technological hardware which are involved. The technical feasibility of using SPR technology to detect interactions in a larger scale array format was announced to the media by Biacore several years ago however, and it is likely that future developments will lead to SPR array systems that are tailored in a format that is appropriate for useful applications within proteomics research, as well as in other applications in life science research and drug development. Precisely where the ﬁnal balance between array size and data/information content will fall remains to be seen for all the technologies that may contribute to the development of protein microarrays, but this may well vary considerably among them and may in the end be determined more forcefully by application requirements rather than by what may be technically possible to achieve.

References 1 LIEDBERG, B., NYLANDER, C., and LUNDSTRO¨ M, I. Surface plasmon resonance for gas detection and biosensing. Sensors Actuators. 1983, 4, 299–304. 2 VENTER, J. C., et al. The sequence of the human genome. Science 2001, 291, 1304–1351. 3 BRETT, D., POSPISIL, H., VALCARCEL, J., REICH, J., and BORK, P. Alternative splicing and genome complexity. Nature Genet. 2002, 30, 29–30. 4 BANKS, R. E., DUNN, M. J., HOCHSTRASSER, D. F., SANCHEZ, J. C., BLACKSTOCK, W., PAPPIN, D. J., and SELBY, P. J. Proteomics: new perspectives, new biomedical opportunities. Lancet 2000, 356, 1749–1756. 5 PARK, S., MYSZKA, D. G., YU, M., LITTLER, S. J., and LAIRD-OFFRINGA, I. A. HuD RNA recognition motifs play distinct

roles in the formation of a stable complex with AU-rich RNA. Mol. Cell Biol. 2000, 20, 4765–4772. 6 KATSAMBA, P. S., MYSZKA, D. G., and LAIRD-OFFRINGA, I. A. Two functionally distinct steps mediate high afﬁnity binding of U1A protein to U1 hairpin II RNA. J. Biol. Chem. 2001, 276, 21476–21481. 7 De GENST, E., ARESKOUG, D., DECANNIERE, K., MUYLDERMANS, S., and ANDERSSON, K. Kinetic and afﬁnity predictions of a protein–protein interaction using multivariate experimental design. J. Biol. Chem. 2002, 277, 29897–29907. 8 CHOULIER, L., ANDERSSON, K., HAMALAINEN, M. D., Van REGENMORTEL, M. H., MALMQVIST, M., and ALTSCHUH, D. QSAR studies applied to the prediction of antigen–antibody interaction kinetics as measured by

43

44 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

9

10

11

12

13

14

15

16

BIACORE. Prot. Eng. 2002, 15, 373– 382. KRETSCHMANN, E. Die Bestimmung optischer Konstanten von Mtallen durch Anregung von Oberﬂ¨achen plasmonenschwingungen. Zeitschrift Physik. 1971, 241, 313–324. L¨ofas, ˚ S., and JOHNSSON, B. A novel hydrogel matrix on gold surfaces in surface plasmon resonance sensors for fast and efﬁcient covalent immobilization of ligands. J. Chem. Soc. Chem. Comm. 1990, 21, 1526–1528. JOHNSSON, B., and L¨OFAS ˚ , S. Immobilization of proteins to a carboxymethyldextran modiﬁed gold surface for biospeciﬁc interaction analysis in surface plasmon resonance. Anal. Biochem. 1991, 198, 268–277. JOHNSSON, B., L¨OFAS ˚ , S., LINDQUIST, G., EDSTRO¨ M, Å., M¨ULLER HILLGREN, R.-M., and HANSSON, A. Comparison of methods for immobilization to carboxymethyl dextran sensor surfaces by analysis of the speciﬁc activity of monoclonal antibodies. J. Mol. Recog. 1995, 8, 125–131. TOKARSKA-SCHLATTNER, M., WALLIMANN, T., and SCHLATTNER, U. Multiple interference of anthracyclines with mitochondrial creatine kinases: preferential damage of the cardiac isoenzyme and its implications for drug cardiotoxicity. Mol. Pharmacol. 2002, 61, 516–523. CALDWELL, E. E., ANDREASEN, A. M., BLIETZ, M. A., SERRAHN, J. N., VANDERNOOT, V., PARK, Y., YU, G., LINHARDT, R. J., and WEILER, J. M. Heparin binding and augmentation of C1 inhibitor activity. Arch. Biochem. Biophys. 1999, 361, 215–222. NIEBA, L., NIEBA-AXMANN, S. E., PERSSON, A., H¨AMA¨ LA¨ INEN, M., EDEBRATT, F., HANSSON, A., LIDHOLM, J., MAGNUSSON, K., KARLSSON, Å.F., and PLU¨ CKTHUN, A. BIACORE analysis of histidine-tagged proteins using a chelating NTA sensor chip. Anal. Biochem. 1997, 252, 217–228. MACKENZIE, C. R., HIRAMA, T., LEE, K. K., ALTMAN, E., and YOUNG, N. M. Quantitative analysis of bacterial toxin afﬁnity and speciﬁcity for glycolipid

17

18

19

20

21

22

23 24

25

26

receptors by surface plasmon resonance. J. Biol. Chem. 1997, 272, 5533–5538. SEVIN-LANDAIS, A., RIGLER, P., TZARTOS, S., HUCHO, F., HOVIUS, R., and VOGEL, H. Functional immobilisation of the nicotinic acetylcholine receptor in tethered lipid membranes. Biophys. Chem. 2000, 85, 141–152. KARLSSON, O. P., and LOFAS, S. Flow-mediated on-surface reconstitution of g-protein coupled receptors for applications in surface plasmon resonance biosensors. Anal. Biochem. 2002, 300, 132–138. Cooper, M. A. OPTICAL biosensors in drug discovery. Nature Rev. Drug Discov. 2002, 1, 515–528. SJO¨ LANDER, S., and URBANICZKY, C. Integrated ﬂuid handling system for biomolecular interaction analysis. Anal. Chem. 1991, 63, 2338–2345. STENBERG, E., PERSSON, B., ROOS, H., and URBANICZKY, C. Quantitative determination of surface concentration of protein with surface plasmon resonance by using radiolabeled proteins. Coll. Interface Sci. 1991, 143, 513–526. KATSAMBA, P. S., PARK, S., and LAIRD-OFFRINGA, I. A. Kinetic studies of RNA–protein interactions using surface plasmon resonance. Methods 2002, 26, 95–104. Myszka, D. G. IMPROVING biosensor analysis. J. Mol. Recognit. 1999, 12, 1–6. KARLSSON, R., KULLMAN-MAGNUSSON, M., HAMALAINEN, M. D., REMAEUS, A., ANDERSSON, K., BORG, P., GYZANDER, E., and DEINUM, J. Biosensor analysis of drug–target interactions: direct and competitive binding assays for investigation of interactions between thrombin and thrombin inhibitors. Anal. Biochem. 2000, 278, 1–13. DAY, Y. S., BAIRD, C. L., RICH, R. L., and MYSZKA, D. G. Direct comparison of binding equilibrium, thermodynamic, and rate constants determined by surface- and solution-based biophysical methods. Prot. Sci. 2002, 11, 1017–1025. MYSZKA, D. G., MORTON, T. A., DOYLE, M. L., and CHAIKEN, I. M. Kinetic analysis of a protein antigen antibody

References

27

28

29

30

31

32

33

34

interaction limited by mass transport on an optical biosensor. Biophys. Chem. 1997, 64, 127–137. MATSUDA, H. Theory of the steady-state current potential curves of redox electrode reactions in hydrodynamic voltammetry. II Laminar pipe- and channel ﬂow. J. Electroanal. Chem. 1967, 15, 325–336. MYSZKA, D. G., HE, X., DEMBO, M., MORTON, T. A., and GOLDSTEIN, B. Extending the range of rate constants available from BIACORE: Interpreting mass transport-inﬂuenced binding data. Biophys. J. 1998, 75, 583–594. SIKAVITSAS, V., NITSCHE, J. M., and MOUNTZIARIS, T. J. Transport and kinetic processes underlying biomolecular interactions in the BIACORE optical biosensor. Biotechnol Prog. 2002, 18, 885–897. RODEN, L. D., and MYSZKA, D. G. Global analysis of a macromolecular interaction measured on BIAcore. Biochem. Biophys. Res. Comm. 1996, 225, 1073–1077. PARK, B. W., ZHANG, H. T., WU, C., BEREZOV, A., ZHANG, X., DUA, R., WANG, Q., KAO, G., O’ROURKE, D. M., GREENE, M. I., and MURALI, R. Rationally designed anti-HER2/neu peptide mimetic disables P185HER2/neu tyrosine kinases in vitro and in vivo. Nature Biotechnol. 2000, 18, 194–198. BEREZOV, A., ZHANG, H. T., GREENE, M. I., and MURALI, R. Disabling erbB receptors with rationally designed exocyclic mimetics of antibodies: structure–function analysis. J. Med. Chem. 2001, 44, 2565–2574. BEREZOV, A., CHEN, J., LIU, Q., ZHANG, H. T., GREENE, M. I., and MURALI, R. Disabling receptor ensembles with rationally designed interface peptidomimetics. J Biol Chem. 2002, 277, 28330–28339. LOUKINOVA, E., RANGANATHAN, S., KUZNETSOV, S., GORLATOVA, N., MIGLIORINI, M. M., LOUKINOV, D., ULERY, P. G., MIKHAILENKO, I., LAWRENCE, D. A., and STRICKLAND, D. K. Platelet-derived growth factor (PDGF)-induced tyrosine phosphorylation of the low density lipoprotein receptor-related protein

35

36

37

38

39

40

41

(LRP). Evidence for integrated co-receptor function betwenn LRP and the PDGF. J. Biol. Chem. 2002, 277, 15499–15506. ROONEY, I. A., BUTROVICH, K. D., GLASS, A. A., BORBOROGLU, S., BENEDICT, C. A., WHITBECK, J. C., COHEN, G. H., EISENBERG, R. J., and WARE, C. F. The lymphotoxin-beta receptor is necessary and sufﬁcient for LIGHT-mediated apoptosis of tumor cells. J. Biol. Chem. 2000, 275, 14307–14315. STEPLEWSKI, Z., LUBECK, M. D., and KOPROWSKI, H. Human macrophages armed with murine immunoglobulin G2a antibodies to tumors destroy human cancer cells. Science 1983, 221, 865–867. VALONE, F. H., KAUFMAN, P. A., GUYRE, P. M., LEWIS, L. D., MEMOLI, V., DEO, Y., GRAZIANO, R., FISHER, J. L., MEYER, L., MROZEK-ORLOWSKI, M. et al. Phase Ia/Ib trial of bispeciﬁc antibody MDX-210 in patients with advanced breast or ovarian cancer that overexpresses the proto-oncogene HER-2/neu. J. Clin. Oncol. 1995, 13, 2281–2292. WEINER, L. M., CLARK, J. I., RING, D. B., and ALPAUGH, R. K. Clinical development of 2B1, a bispeciﬁc murine monoclonal antibody targeting c-erbB-2 and Fc gamma RIII. J. Hematother. 1995, 4, 453–456. WEINER, L. M., ALPAUGH, R. K., AMOROSO, A. R., ADAMS, G. P., RING, D. B., and BARTH, M. W. Human neutrophil interactions of a bispeciﬁc monoclonal antibody targeting tumor and human Fc gamma RIII. Cancer Immunol. Immunother. 1996, 42, 141–150. MCCALL, A. M., ADAMS, G. P., AMOROSO, A. R., NIELSEN, U. B., ZHANG, L., HORAK, E., SIMMONS, H., SCHIER, R., MARKS, J. D., and WEINER, L. M. Isolation and characterization of an anti-CD16 single-chain Fv fragment and construction of an anti-HER2/neu/anti-CD16 bispeciﬁc scFv that triggers CD16-dependent tumor cytolysis. Mol. Immunol. 1999, 36, 433–445. YOKOTA, T., MILENIC, D. E., WHITLOW, M., and SCHLOM, J. Rapid tumor penetration of a single-chain Fv and

45

46 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

42

43

44

45

46

47

48

49

comparison with other immunoglobulin forms. Cancer Res. 1992, 52, 3402–3408. CASEY, J. L., KEEP, P. A., CHESTER, K. A., ROBSON, L., HAWKINS, R. E., and BEGENT, R. H. Puriﬁcation of bacterially expressed single chain Fv antibodies for clinical applications using metal chelate chromatography. J. Immunol. Methods 1995, 179, 105–116. BEGENT, R. H., VERHAAR, M. J., CHESTER, K. A., CASEY, J. L., GREEN, A. J., NAPIER, M. P., HOPE-STONE, L. D., CUSHEN, N., KEEP, P. A., JOHNSON, C. J., HAWKINS, R. E., HILSON, A. J., and ROBSON, L. Clinical evidence of efﬁcient tumor targeting based on single-chain Fv antibody selected from a combinatorial library. Nature Med. 1996, 2, 979–984. NERI, D., CARNEMOLLA, B., NISSIM, A., LEPRINI, A., QUERZE, G., BALZA, E., PINI, A., TARLI, L., HALIN, C., NERI, P., ZARDI, L., and WINTER, G. Targeting by afﬁnity-matured recombinant antibody fragments of an angiogenesis associated ﬁbronectin isoform. Nature Biotechnol. 1997, 15, 1271–1275. BADER, B., KUHN, K., OWEN, D. J., WALDMANN, H., WITTINGHOFER, A., and KUHLMANN, J. Bioorganic synthesis of lipid-modiﬁed proteins for the study of signal transduction. Nature 2000, 403, 223–226. HUANG, X., GOTTSTEIN, C., BREKKEN, R. A., and THORPE, P. E. Expression of soluble VEGF receptor 2 and characterization of its binding by surface plasmon resonance. Biochem. Biophys. Res. Commun. 1998, 252, 643–648. KNIBIEHLER, M., GOUBIN, F., ESCALAS, N., , Z. O., MAZARGUIL, H., JONSSON ´ H¨UBSCHER, U., and DUCOMMUN, B. Interaction studies between the p21Cip1/Waf1 cyclin-dependent kinase inhibitor and proliferating cell nuclear antigen (PCNA) by surface plasmon resonance. FEBS Lett. 1996, 391, 66–70. KANAOKA, Y., KIMURA, S. H., OKAZAKI, I., IKEDA, M., and NOJIMA, H. GAK: a cyclin G associated kinase contains a tensin/auxilin-like domain. FEBS Lett. 1997, 402, 73–80. SHIN, S., SUNG, B. J., CHO, Y. S., KIM, H. J., HA, N. C., HWANG, J. I., CHUNG, C.

50

51

52

53

54

55

56

W., JUNG, Y. K., and OH, B. H. An anti-apoptotic protein human survivin is a direct inhibitor of cas-pase-3 and -7. Biochemistry 2001, 40, 1117–1123. HEIDER, K. H., SPROLL, M., SUSANI, S., PATZELT, E., BEAUMIER, P., OSTERMANN, E., AHORN, H., and ADOLF, G. R. Characterization of a high-afﬁnity monoclonal antibody speciﬁc for CD44v6 as candidate for immunotherapy of squamous cell carcinomas. Cancer Immunol. Immunother. 1996, 43, 245–253. Van HAL, N. L., Van DONGEN, G. A., TEN BRINK, C. B., HEIDER, K. H., RECH-WEICHSELBRAUN, I., SNOW, G. B., and BRAKENHOFF, R. H. Evaluation of soluble CD44v6 as a potential serum marker for head and neck squamous cell carcinoma. Clin. Cancer Res. 1999, 5, 3534–3541. VEREL, I., HEIDER, K. H., SIEGMUND, M., OSTERMANN, E., PATZELT, E., SPROLL, M., SNOW, G. B., ADOLF, G. R., and Van DONGEN, G. A. Tumor targeting properties of monoclonal antibodies with different afﬁnity for target antigen CD44V6 in nude mice bearing head-and-neck cancer xenografts. Int. J. Cancer 2002, 99, 396–402. RITTER, G., COHEN, L. S., WILLIAMS, C., Jr., RICHARDS, E. C., OLD, L. J., and WELT, S. Serological analysis of human anti-human antibody responses in colon cancer patients treated with repeated doses of humanized monoclonal antibody a33. Cancer Res. 2001, 61, 6851–6859. ALAEDINI, A., and LATOV, N. A surface plasmon resonance biosensor assay for measurement of anti-GM(1) antibodies in neuropathy. Neurology 2001, 56, 855–860. LILJEBLAD, M., RYDEN, I., OHLSON, S., LUNDBLAD, A., and PAHLSSON, P. A lectin immunosensor technique for determination of alpha(1)-acid glycoprotein fucosylation. Anal. Biochem. 2001, 288, 216–224. EIVAZOVA, E. R., MCDONNELL, J. M., SUTTON, B. J., and STAINES, N. A. Speciﬁcity and binding kinetics of murine lupus anti-DNA monoclonal

References

57

58

59

60

61

62

63

64

antibodies implicate different stimuli for their production (In Process Citation). Immunology 2000, 101, 371–377. GUERMAZI, S., REGNAULT, V., GORGI, Y., AYED, K., LECOMPTE, T., and DELLAGI, K. Further evidence for the presence of anti-protein S autoantibodies in patients with systemic lupus erythematosus (In Process Citation). Blood Coagul. Fibrinolysis. 2000, 11, 491–498. JONES, D. W., NICHOLLS, P. J., DONOHOE, S., GALLIMORE, M. J., and WINTER, M. Antibodies to factor XII are distinct from antibodies to prothrombin in patients with the anti-phospholipid syndrome. Thromb. Haemost. 2002, 87, 426–430. NEDELKOV, D., and NELSON, R. W. Analysis of human urine protein biomarkers via biomolecular interaction analysis mass spectrometry. Am. J Kidney Dis. 2001, 38, 481–487. BOHRMANN, B., TJERNBERG, L., KUNER, P., POLI, S., LEVET-TRAFIT, B., NASLUND, J., RICHARDS, G., HUBER, W., DOBELI, H., and NORDSTEDT, C. Endogenous proteins controlling amyloid beta-peptide polymerization. Possible implications for beta-amyloid formation in the central nervous system and in peripheral tissues. J. Biol. Chem. 1999, 274, 15990–15995. VORUM, H., JACOBSEN, C., and HONORE, B. Calumenin interacts with serum amyloid P component. FEBS Lett. 2000, 465, 129–134. LEFEBVRE, S., BURGLEN, L., REBOULLET, S., CLERMONT, O., BURLET, P., VIOLLET, L., BENICHOU, B., CRUAUD, C., MILLASSEAU, P., ZEVIANI, M. et al. Identiﬁcation and characterization of a spinal muscular atrophy-determining gene. Cell 1995, 80, 155–165. MONANI, U. R., LORSON, C. L., PARSONS, D. W., PRIOR, T. W., ANDROPHY, E. J., BURGHES, A. H., and MCPHERSON, J. D. A single nucleotide difference that alters splicing patterns distinguishes the SMA gene SMN1 from the copy gene SMN2. Hum. Mol. Genet. 1999, 8, 1177–1183. YOUNG, P. J., MAN, N., LORSON, C. L., LE, T. T., ANDROPHY, E. J., BURGHES, A. H., and MORRIS, G. E. The exon 2b region of the spinal muscular atrophy protein,

65

66

67

68

69

70

71

72

73

SMN, is involved in self-association and SIP1 binding. Hum. Mol. Genet. 2000, 9, 2869–2877. ARIGA, T., and YU, R. K. GM1 inhibits amyloid beta-protein-induced cytokine release. Neurochem Res. 1999, 24, 219–226. CATIMEL, B., SCOTT, A. M., LEE, F. T., HANAI, N., RITTER, G., WELT, S., OLD, L. J., BURGESS, A. W., and NICE, E. C. Direct immobilization of gangliosides onto gold-carboxymethyldextran sensor surfaces by hydrophobic interaction: applications to antibody characterization. Glycobiology 1998, 8, 927–938. NAKAMURA, K., TANAKA, Y., SHITARA, K., and HANAI, N. Construction of humanized anti-ganglioside monoclonal antibodies with potent immune effector functions. Cancer Immunol. Immunother. 2001, 50, 275–284. Snider, W. D. FUNCTIONS of the neurotrophins during nervous system development: what the knockouts are teaching us. Cell 1994, 77, 627–638. IP, N. Y., and YANCOPOULOS, G. D. The neurotrophins and CNTF: two families of collaborative neurotrophic factors. Annu. Rev. Neurosci. 1996, 19, 491–515. LI, S.-C., LAI, K.-M. V., GISH, G. D., PARRIS, W. E., Van der GEER, P., FORMAN-KAY, J., and PAWSON, T. Characterization of the phospho-tyrosine-binding domain of the Drosophila Shc protein. J. Biol. Chem. 1996, 271, 31855–31862. WOO, S. B., WHALEN, C., and NEET, K. E. Characterization of the recombinant extracellular domain of the neurotrophin receptor TrkA and its interaction with nerve growth factor (NGF). Prot. Sci. 1998, 7, 1006–1016. ROSS, G. M., SHAMOVSKY, I. L., WOO, S. B., POST, J. I., VRKLJAN, P. N., LAWRANCE, G., SOLC, M., DOSTALER, S. M., NEET, K. E., and RIOPELLE, R. J. The binding of zinc and copper ions to nerve growth factor is differentially affected by pH: implications for cerebral acidosis. J. Neurochem. 2001, 78, 515–523. HOLCOMB, I. N., et al. FIZZ1, a novel cysteine-rich secreted protein associated with pulmonary inﬂammation, deﬁnes a

47

48 Real-time Characterization of Biomolecular Interactions using Biacore’s Optical Biosensors

74

75

76

77

78

79

80

81

new gene family. EMBO J. 2000, 19, 4046–4055. CAIRO, C. W., STRZELEC, A., MURPHY, R. M., and KIESSLING, L. L. Afﬁnity-based inhibition of beta-amyloid toxicity. Biochemistry 2002, 41, 8620–8629. YAMAZAKI, M., MORI, H., ARAKI, K., MORI, K. J., and MISHINA, M. Cloning, expression and modulation of a mouse NMDA receptor sub-unit. FEBS Lett. 1992, 300, 39–45. LOMELI, H., SPRENGEL, R., LAURIE, D. J., KOHR, G., HERB, A., SEEBURG, P. H., and WISDEN, W. The rat delta-1 and delta-2 subunits extend the excitatory amino acid receptor family. FEBS Lett. 1993, 315, 318–322. ARAKI, K., MEGURO, H., KUSHIYA, E., TAKAYAMA, C., INOUE, Y., and MISHINA, M. Selective expression of the glutamate receptor channel delta 2 subunit in cerebellar Purkinje cells. Biochem. Biophys. Res. Commun. 1993, 197, 1267–1276. MIYAGI, Y., YAMASHITA, T., FUKAYA, M., SONODA, T., OKUNO, T., YAMADA, K., WATANABE, M., NAGASHIMA, Y., AOKI, I., OKUDA, K., MISHINA, M., and KAWAMOTO, S. Delphilin: a novel PDZ and formin homology domain-containing protein that synaptically colocalizes and interacts with glutamate receptor delta 2 subunit. J. Neurosci. 2002, 22, 803–814. YOUNG, P. J., DAY, P. M., ZHOU, J., ANDROPHY, E. J., MORRIS, G. E. and LORSON, C. L. A direct interaction between the survival motor neuron protein and p53 and its relationship to spinal muscular atrophy. J. Biol. Chem. 2002, 277, 2852–2859. LAUGHTON, C. A., TANIOUS, F., NUNN, C. M., BOYKIN, D. W., WILSON, W. D., and NEIDLE, S. A crystallographic and spectroscopic study of the complex between d(CGCGAATTCGCG)2 and 2,5-bis(4-guanylphenyl)furan, an analogue of berenil. Structural origins of enhanced DNA-binding afﬁnity. Biochemistry 1996, 35, 5655–5661. CARRASCO, C., FACOMPRE, M., CHISHOLM, J. D., Van VRANKEN, D. L., WILSON, W. D., and BAILLY, C. DNA sequence recognition by the indolo-carbazole

82

83

84

85

86

87

88

89

90

antitumor antibiotic AT2433-B1 and its diastereoisomer. Nucleic Acids Res. 2002, 30, 1774–1781. CARRASCO, C., VEZIN, H., WILSON, W. D., REN, J., CHAIRES, J. B., and BAILLY, C. DNA binding properties of the indolocarbazole antitumor drug NB-506. Anticancer Drug Des. 2001, 16, 99–107. WANG, L., BAILLY, C., KUMAR, A., DING, D., BAJIC, M., BOYKIN, D. W., and WILSON, W. D. Speciﬁc molecular recognition of mixed nucleic acid sequences: an aromatic dication that binds in the DNA minor groove as a dimer. Proc. Natl Acad. Sci. USA 2000, 97, 12–16. DAVIS, T. M., and WILSON, W. D. Determination of the refractive index increments of small molecules for correction of surface plasmon resonance data. Anal. Biochem. 2000, 284, 348–353. SANDO, S., SAITO, I., and NAKATANI, K. Scanning of guanine–guanine mismatches in DNA by synthetic ligands using surface plasmon resonance. Nature Biotechnol. 2001, 19, 51–55. Kwok, P. Y. REFLECTIONS on a DNA mutation scanning tool. Nature Biotechnol. 2001, 19, 18–19. LACY, E. R., LE, N. M., PRICE, C. A., LEE, M., and WILSON, W. D. Inﬂuence of a terminal formamido group on the sequence recognition of DNA by polyamides. J. Am. Chem. Soc. 2002, 124, 2153–2163. HICKMAN, D. T., KING, P. M., SLATER, J. M., COOPER, M. A., and MICKLEFIELD, J. Kinetically selective binding of single stranded RNA over DNA by a pyrrolidine-amide oligonucleotide mimic (POM). Nucleosides Nucleotides Nucleic Acids 2001, 20, 1169–1172. HOU, M. H., LIN, S. B., YUANN, J. M., LIN, W. C., WANG, A. H., and Kan LS, L. Effects of polyamines on the thermal stability and formation kinetics of DNA duplexes with abnormal structure. Nucleic Acids Res. 2001, 29, 5121– 5128. TSOI, P. Y., and YANG, M. Kinetic study of various binding modes between human DNA polymerase beta and different DNA substrates by

References

91

92

93

94

95

96

97

98

surface-plasmon-resonance biosensor. Biochem. J. 2002, 361, 317–325. SUMMERTON, J., and WELLER, D. Morpholino antisense oligomers: design, preparation, and properties. Antisense Nucleic Acid Drug Dev. 1997, 7, 187–195. MANG’ERA, K. O., LIU, G., YI, W., ZHANG, Y., LIU, N., GUPTA, S., RUSCKOWSKI, M., and HNATOWICH, D. J. Initial investigations of 99mTc-labeled morpholinos for radiopharmaceutical applications. Eur. J. Nucl. Med. 2001, 28, 1682–1689. ASTRIAB-FISHER, A., SERGUEEV, D., FISHER, M., SHAW, B. R., and JULIANO, R. L. Conjugates of antisense oligonucleotides with the Tat and antennapedia cell-penetrating peptides: effects on cellular uptake, binding to target sequences, and biologic actions. Pharm. Res. 2002, 19, 744–754. BATES, P., REDDOCH, J., HANSAKUL, P., ARROW, A., DALE, R., and MILLER, D. Biosensor detection of triplex formation by modiﬁed oligonucleotides. Anal. Biochem. 2002, 307, 235. OKUMOTO, Y., OHMICHI, T., and SUGIMOTO, N. Immobilized small deoxyribozyme to distinguish RNA secondary structures. Biochemistry 2002, 41, 2769–2773. NILSSON, P., PERSSON, B., UHLE´ N, M., and NYGREN, P.-Å. Real-time monitoring of DNA manipulations using biosensor technology. Anal. Biochem. 1995, 224, 400–408. STEINRUCKE, P., ALDINGER, U., HILL, O., HILLISCH, A., BASCH, R., and DIEKMANN, S. Design of helical proteins for real-time endoprotease assays. Anal. Biochem. 2000, 286, 26–34. LOEW, D., PERRAULT, C., MORALES, M., MOOG, S., RAVANAT, C., SCHUHLER, S., ARCONE, R., PIETROPAOLO, C., CAZENAVE, J. P., Van DORSSELAER, A., and LANZA, F.

99

100

101

102

103

104

105

Proteolysis of the exodomain of recombinant protease-activated receptors: prediction of receptor activation or inactivation by MALDI mass spectrometry. Biochemistry 2000, 39, 10812–10822. ZAKO, T., HARADA, K., MANNEN, T., YAMAGUCHI, S., KITAYAMA, A., UEDA, H., and NAGAMUNE, T. Monitoring of the refolding process for immobilized ﬁreﬂy luciferase with a biosensor based on surface plasmon resonance. J. Biochem. (Tokyo). 2001, 129, 1–4. HOUSEMAN, B. T., HUH, J. H., KRON, S. J., and MRKSICH, M. Peptide chips for the quantitative evaluation of protein kinase activity. Nature Biotechnol. 2002, 20, 270–274. KATSAMBA, P. S., BAYRAMYAN, M., HAWORTH, I. S., MYSZKA, D. G., and LAIRD-OFFRINGA, I. A. Complex role of the beta 2–beta 3 loop in the interaction of U1A with U1 hairpin II RNA. J. Biol. Chem. 2002, 277, 33267–33274. NELSON, R. W., NEDELKOV, D., and TUBBS, K. A. Biosensor chip mass spectrometry: a chip-based proteomics approach. Electrophoresis 2000, 21, 1155–1163. KRONE, J. R., NELSON, R. W., DOGRUEL, D., WILLIAMS, P., and GRANZOW, R. BIA/MS: interfacing biomolecular interaction analysis with mass spectrometry. Anal. Biochem. 1997, 244, 124–132. S¨ONKSEN, C. P., NORDHOFF, E., JANSSON, ¨ MALMQVIST, M., and ROEPSTORFF, P. O., Combining MALDI mass spectrometry and biomolecular interaction analysis using a biomolecular interaction analysis instrument. Anal. Chem. 1998, 70, 2731–2736. WILLIAMS, I., and ADDONA, T. A. The integration of SPR biosensors with mass spectrometry: possible applications for proteome analysis (In Process Citation). Trends Biotechnol. 2000, 18, 45–48.

49

1

Cellular Assays and Their Application in Drug Discovery

Hugo Albrecht, Daniela Brodbeck-Hummel, Michael Hoever, Beatrice Nickel, and Urs Regenass Discovery Partners International AG, Allschwil, Switzerland

Originally published in: Molecular Biology in Medicinal Chemistry. Edited by Theodor Dingermann, Dieter Steinhilber and Gerd Folkers. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30431-8

1 Introduction 1.1 Positioning Cellular Assays

Cell-based assays allow to study the function of pharmaceutical or disease targets within complex environments and in the overall biological context. The application of techniques in molecular biology together with the introduction of new tools and analytical devices has moved readouts of cellular assays from phenotypic endpoints such as cell proliferation, cell death, respiration and functional differentiation to the cellular analysis of functional states of speciﬁc signaling molecules and metabolic components. This transition of cellular assays from “black box” systems to speciﬁc targetassociated, mechanistic measurements allows the identiﬁcation of chemical compounds with target-speciﬁc or closely associated modulatory function. It can therefore be assumed that cellular assays will play an increasingly important role in early drug discovery, in particular in discovering compounds for target validation (“chemical genetics”; [1] for review) and in hit-to-lead selection as well as in lead optimization. In contrast to biochemical testing of chemicals for modulatory activity on molecular targets, cellular systems can deliver additional information with respect to drug transport, metabolism and stability. Most importantly, pharmacological targets remain in their natural environment when probed for modulation and data should therefore have a higher predictive value (Fig. 1).

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Cellular Assays and Their Application in Drug Discovery

Tight target coupled and phenotypic cellular assays

Tight target coupled cellular assays

Identification of potential targets

Assay Development

Hit Finding

Lead Finding

Target validation Disease linkage, Functional studies, Chemogenomics

Lead driven Target selection Lead Optimization Drug Candidate Selection

Fig. 1 Chemistry-driven target selection. The increased number of potential pharmaceutical targets emerging from the functional annotation of the human genome requires an efﬁcient process to select those that are valid for modulating diseaserelevant pathways and phenotypes. Cellular assays which are tightly coupled in the readout with the target of interest allow us

•Tight target coupled assays •Phenotypic assays •Loose target coupled or uncoupled assays: Systems effects •Intracellular pharmacology to qualify the hits identiﬁed in cellular or biochemical high-throughput assays with respect to functionality in a physiologically relevant environment. Cellular assays which are not tightly coupled to the target will indicate system effects of compounds and can be interpreted as a cellular “safety” classiﬁcation. Cellular assays can be used for target and compound prioritization.

However, it remains to be said that, in order to use cellular assays in high throughput, cells need to be taken out of their natural habitat and put into culture. This will inevitably lead to alterations in the repertoire of gene and protein expression, and therefore in their phenotype. Cells used in assays should therefore be characterized with respect to the functionality of their intracellular networks, responses to external stimuli, at least for the signaling pathway of interest, and compared with responses and signaling events at the tissue or organ level. In many cases this is unfortunately not yet fully possible. 1.2 Impact on Drug Discovery

Many compounds are lost during the discovery and development process due to undesirable effects or toxicity and lack of required efﬁcacy. A large part of these effects can be related to interactions of the potential drug with targets other than the

1 Introduction 3

one initially selected to modulate disease. Cellular test systems have the potential to elucidate the multiplicity of interactions of drugs, and therefore contribute to a better understanding of drug action and drug selection. The cell is organized in metabolic and signal transduction cascades. Most of these cascades are complex and nonlinear [2]. In addition, different cascades communicate with each other, and form intracellular circuits and domains. Most diseases cannot be explained by one altered step in a pathway, but are due to several alterations that affect networks in a complex way [3, 4]. Similarly, drugs, whether they interact speciﬁcally with one single target or with several molecules, will disturb signaling and metabolic cascades in a complex fashion. In addition, since many cell types make use of the same types of signaling molecules, in a different context, many cell types might be affected to different extents by the same drug. The recent introduction of high-content screening, high-throughput gene and protein expression, as well as metabolite analysis [5, 6], offers the possibility to study system interactions in a multiparallel fashion, and allows a better understanding of the degree and location of interference. Cell-based assays therefore provide a solution to study the modulation of a pharmaceutical target within its complex natural network as well as to assess potential effects on other networks. Human diseases are characterized by one or several alterations in metabolic and signaling pathways important to maintain cellular homeostasis and the differentiated functions. Cellular assays can offer a representation of such alterations and therefore become relevant functional models to assess drug action. Nowadays, cellular assays offer the opportunity to identify the intervention points that lead to phenotypic or endpoint alterations. Often, multiple intervention sites are necessary to alter the outcome of biological networks. Similar rules might apply for the number of drug targets that need to be modulated; in particular, for complex diseases. The right targets and drug combinations can be selected only in the case where each single component of a particular pathway can be individually studied, and this is only rendered possible by the use of cellular systems. The combination of compound that lead to the desired endpoint can be identiﬁed by analyzing multiple parameters. The analysis can, moreover, facilitate the assessment of the effect of individual compounds on multiple pathways. This can be taken as a measure for “unwanted effects” (safety proﬁle), but can also lead to the identiﬁcation of applications in new indications. The analysis of targeted and focused chemical libraries in cellular systems may allow a rapid selection of compounds with various and desired proﬁles. Hierarchical, spatio-temporal readouts will generate system-level insight into mechanisms of drug action (intracellular pharmacology) and allow us to predict systemic effects (Fig. 2). The development and application of computational tools as well as models of intracellular pathways are a prerequisite to secure data evaluation and interpretation [7, 8]. System-level understanding is only possible by seeking experimental results at the cellular level or beyond. The recent developments of cellular high-throughput systems, which allow speciﬁc measurements of molecular events, render applications in primary screening, and in hit-to-lead selection and lead optimization, in particular, possible. This will

4 Cellular Assays and Their Application in Drug Discovery

Extracellular space

• Receptors • Transporters • Ion channels

•pH •pO2

Legend Receptor ligands Phosphorylation Ions, metabolites Translocstions

Cytoplasm

Proteins

with Organelles

Proteins

Sensors fast response

m RNA

medium response slow response

Nucleus with DNA and transcription machinery

+++

+++

Fig. 2 Schematic cellular circuit and cell systems analysis concept. Cells connect to their environment with a wealth of receptors, channels and transporters. Intracellular signaling and metabolic pathways and networks react to intra- and extracellular stimuli in a spatio-temporal manner. Different cell types can react differently to the same stimuli. It can be envisaged that different cell types, cells from normal or pathologi-

cal tissue, or genetically manipulated cells, can be exposed to different stimuli with and without drugs and the cellular reaction can be assessed by sensor systems in a spatiotemporal fashion, from fast responses to phenotypic changes. Multiparametric analysis will identify compounds with targetspeciﬁc, target-unrelated, reversible or irreversible effects on cellular homeostasis.

enable the development of structure–activity relationships at the system level as a new paradigm in compound selection and decision-making. 1.3 Classiﬁcation of Cellular Assays

Cellular assays have the advantage that the pharmacological targets do not need to be puriﬁed. However, in many instances, cellular assays require cell manipulation to allow for a speciﬁc signal readout. Overexpression of cell-surface receptors or intracellular proteins can have a substantial effect on the cell physiology and have to be taken into account when using the systems for the discovery of novel drugs. Early reactions to hormones, growth factors and neurotransmitters are often mediated via second-messenger systems or rapid inﬂux and efﬂux of ions. Posttranslational protein modiﬁcations and translocation of proteins are the next steps that alter the physiological state of cells. Such changes are able to lead to ﬂow changes in metabolic pathways, and to alterations in gene and protein expression, ultimately leading to new cellular phenotypes, including cell proliferation and cell death.

1 Introduction 5 Table 1 Classiﬁcation of functional cellular assay types (see text for references)

Second messengers and channels

Protein dynamics

Gene expression

Protein expression

Metabolites proﬁling Metabolic and phenotypic responses

Ca2+ levels ion channels transporters cyclic nucleotides posttranslational modiﬁcations (e.g. phosphorylation) protein translocations protein–protein interaction chip technologies branched DNA technology oligonucleotide sensors reporter gene technology ELISA assays protein chip arrays quantitative analytical proﬁling cell proliferation, cytotoxicity, cell death/apoptosis respiration, acidiﬁcation cell differentiation

Cellular assays can be classiﬁed by the temporal events occurring when a cell is exposed to alterations in the extra- and intracellular environment (Tab. 1). Fast responses indicated by changes in ion or second-messenger concentrations ultimately lead to functional and phenotypic changes. Some of the cellular events can be recapitulated in so-called model systems, which might be easier to handle and to manipulate genetically than mammalian cells. Yeast has turned out to be an organism of choice, where metabolic signals and protein–protein interactions relevant for signal transduction can be translated into easily measurable growth response (for review, see [9, 10]. Several modiﬁcations of the yeast two-hybrid system have shown utility in ﬁnding compounds that functionally interfere with speciﬁc intracellular mechanisms. Most recently, a sensor system for drug discovery based on conformational activation of a proliferationrelevant enzyme upon drug binding has been described [11]. However, compared to mammalian cells, the yeast cell has some trade-offs, which include the cell wall, lack of the full complement of mammalian genes and alterations in its metabolic systems. The use of yeast two-hybrid and yeast “tribrid” systems [12] are not discussed further in this article. 1.4 Progress in Tools and Technologies for Cellular Compound Proﬁling

The reﬁnement of analytical devices [13] and reader systems today enables investigators to obtain broad proﬁles of alterations in gene and protein expression [14] as well as metabolites [5, 15]. Unless a complete system-wide analysis is required, the present technologies allow a reasonable throughput for drug proﬁling. For gene expression studies, reporter gene assays can substitute for chip-based arrays, at least in selected cases [16, 17].

6 Cellular Assays and Their Application in Drug Discovery

Several cellular imaging or high-throughput microscopy and laser-scanning systems are now becoming available together with the development of new ﬂuorescent tags and speciﬁc antibodies. This combination allows us to study protein movements in cells. The study of protein dynamics and interactions has been largely facilitated by the identiﬁcation of ﬂuorescent proteins [6, 18, 19] and other ﬂuorescent dyes [20] in combination with the development of ﬂuorescence resonance energy transfer (FRET) applications. Other technologies such as multipole coupling spectroscopy (MCS) [21] and biochips for multiparametric cell monitoring [22] are just becoming available. The FLIPRTM (FLuorometric Imaging Plate Reader; Molecular Devices, Sunnyvale, CA) system has been available for several years to study rapid cellular second-messenger responses, such as Ca2+ release, and has been recently upgraded for the analysis of ion channels. The FLIPR applications are discussed in detail below. In the following sections, we will describe a selected set of applications in more detail. The discussion is grouped according to Tab. 1 into fast responses or secondmessenger systems, protein dynamics, alterations in gene and protein expression, and metabolic as well as phenotypic readouts.

2 Membrane Proteins and Fast Cellular Responses

A great variety of membrane proteins have been expressed by using recombinant DNA technology in different eukaryotic cells (e.g. mammalian cell lines, insect cells and yeast) for functional studies of cell-surface receptors, ion channels or transport proteins. 2.1 Receptors

Cell-surface receptors typically mediate cell signaling from the cell surface to the cytoplasm and/or nucleus involving two major principles, i.e. signaling via enzymelinked receptors or trimeric GTP-binding proteins (G-proteins). The enzymelinked receptors include the receptor guanylyl cyclases, receptor tyrosine kinases, tyrosine-kinase-associated receptors, receptor tyrosine phosphatases and receptor serine/threonine kinases [23]. In all cases, a cascade of events is initiated upon activation, which alters receptor status and the concentration of one or more small intracellular signaling molecules. Here, we will discuss assays monitoring signals mediated by G-proteins, which are activated by G-protein-coupled receptors (GPCRs) upon binding of a speciﬁc ligand. Subsequent to stimulation, changes of intracellular cyclic AMP (cAMP) or Ca2+ concentrations are induced. These two signaling molecules are known to be involved in regulatory processes of the cardiovascular and nervous systems, immune mechanisms, cell growth and differentiation, and general metabolism. Both messengers act as allosteric effectors on

2 Membrane Proteins and Fast Cellular Responses

speciﬁc target proteins in the cell including, but not limited to, kinases (e.g. protein kinase A and C), lipases (e.g. phospholipase Cβ), Ca2+ -binding proteins (e.g. calmodulin) and ion channels (e.g. ionotropic glutamate receptors). The cytoplasmic concentrations of the second messengers are regulated via plasma membrane enzymes – in the cAMP pathway the enzyme adenylate cyclase directly produces cAMP and in the Ca2+ pathway lipases are stimulated to produce the soluble messenger inositol triphosphate (IP3 ), which in turn induces the release of Ca2+ from the endoplasmic reticulum. In some cases, ion channels are activated by direct interaction with G-protein subunits. Different technologies have been developed for intracellular monitoring of second messengers in living cells. Detection of Ca2+ is generally based on ﬂuorescent indicators, which have the property of being able to easily penetrate cell membranes and alter their ﬂuorescent emission when bound to Ca2+ . For measurement of a typical fast and transient response, the binding reaction to Ca2+ must be reversible, therefore providing important kinetic data which allow us to distinguish between a typically transient signal due to speciﬁc receptor activation and nonspeciﬁc signals affecting Ca2+ homeostasis or membrane permeability. There are currently many different systems available to detect cAMP in cellbased assays. In most cases cells are lysed after exposure to a speciﬁc ligand and the amount of cAMP is determined with a competitive immunoassay (PerkinElmer Life Sciences, Boston, MA; Molecular Devices; Biomol Research Laboratories, Plymouth Meeting, CA) or a competitive enzyme complementation assay (DiscoveRx, Fremont, CA). Typically, GPCRs are overexpressed in eukaryotic cell lines to monitor the concentration of second messengers upon stimulation with a speciﬁc ligand [24]. In many cases, the co-expression of a suitable G-protein subunit or chimeric G-proteins is necessary in order to elicit a signal sufﬁcient for detection [25, 26].

2.1.1 FLIPR Technology for Detection of Intracellular Calcium Release The FLIPR system provides readouts for high-throughput, cell-based assays that represent information-rich kinetic data in 96- or 384-well format. In brief, cells are loaded with a ﬂuorescent indicator (e.g. Fluo-4 AM for Ca2+ ), which shows an absorption spectrum compatible with excitation at 488 nm with an argon-ion laser source, leading to maximum emission at 516 nm in the presence of Ca2+ . Fluorescence emission is induced simultaneously in all wells of a 96- or 384-well plate and kinetics of intracellular Ca2+ release, triggered by exposure to a speciﬁc ligand, can be monitored at a maximum of 60 sequential measurements per minute. The FLIPR system rapidly discriminates between full agonists, partial agonists and antagonists to accelerate primary and secondary screening. Figure 3 illustrates a typical experiment conducted with a mammalian cell line stably transfected with an expression vector encoding a GPCR gene. Positive compounds depicted in Fig. 3b were subjected to veriﬁcation in duplicate measurement and conﬁrmed hits were further tested for dose-response relationships in EC50 /IC50 experiments.

7

8 Cellular Assays and Their Application in Drug Discovery

2.1.2 Competitive Immunoassay for Detection of Intracellular cAMP Most assays designed for detection of intracellular cAMP are based on monoclonal antibodies speciﬁcally recognizing cAMP. Typically, cell lysates are mixed with exogenously added biotinylated cAMP to form a heterotrimer with a speciﬁc antibody and streptavidin. This complex formation is competed by endogenous non-biotinylated cAMP. Streptavidin and antibodies can be linked covalently to a donor and an acceptor ﬂuorophore (e.g. europium cryptate and allophycocyanin), forming an interacting FRET pair upon complex formation. A decreased signal is observed with an increase in intracellular cAMP produced, due to competition for binding to the speciﬁc antibody. The AlphaScreenTM technology (PerkinElmer Life Sciences) is an alternative technology that allows quantitative detection of a broad range of cellular components,

(a)

20000

Max2

17900

Max1

15800

Fluorescence Change (Counts)

13700 11600 9500 7400 5300 3200 1100 Min -1000 00

20

Compound addition

40

60 80 100 Time (seconds)

120

140

160

180

200

Ligand addition

Fig. 3 Determination of intracellular Ca2+ with the FLIPR system. (a) A ﬂuorescent trace of a typical Ca2+ kinetic is shown and the measured parameters used for calculation of maximal ﬂuorescence signals are depicted. In this example the maxima were measured before and after ligand addition (Max1 and Max2 respectively) and standardized by dividing by the minima measured

before compound addition (Min = baseline). The average of the positive controls was set as 100% control and the average of the negative controls was set as 0% control. For all wells, relative activities with respect to the controls were calculated and the cutoff criterion for both agonists and antagonists was 50%.

2 Membrane Proteins and Fast Cellular Responses

(b)

Agonist

1234567891011121314151617181920212223

151617181920212223

Seed GPCR over-expressing cells into384-well plates

24

A B C D E 1234567891011121314151617181920212223 A

F

B

G

C

H

D

I

E

J

F

24

Incubate over night

K

G

L

H

M

I

N

J

O

K

P

Load cells with Fluo-4 - AM dye

L M N

Screen chemical library

O P

Antagonist

Agonists

NO

Stop

NO

YES

YES

Verification in duplicate

Agonists

Antagonists

NO

YES

EC50 Determination Fig. 3 (Continue)(b) A ﬂow diagram for a typical experiment is shown. Genetically modiﬁed cells overexpressing the target GPCR were seeded into 384-well plates overnight. The cells were washed with suitable buffer prior to adding buffer containing the Fluo-4 AM dye, followed by incubation for 45 min. After an additional wash step, the cell plates were transferred into the FLIPR machine and the chemical com-

Verification in duplicate

Stop

NO

Antagonists YES

IC50 Determination pounds were added followed by ligand addition. Fluorescence emission was monitored starting before compound addition and continued until a substantial decay of the signal was observed. Two 384-well plates from a typical screen are included. Examples of an agonist and an antagonist are shown as enlarged graphs. The 100% and 0% controls are overlaid as red- and blue-hatched lines, respectively.

9

10 Cellular Assays and Their Application in Drug Discovery

including cAMP. AlphaScreen relies on the use of “donor” and “acceptor” beads that are coated with a layer of hydrogel providing functional groups for bioconjugation. When the beads are brought into proximity, a cascade of chemical reactions is initiated to produce a greatly ampliﬁed signal. Upon laser excitation, a photosensitizer in the “donor” bead converts ambient oxygen to a more excited singlet state. The singlet state oxygen molecules diffuse across to react with a chemiluminescent in the “acceptor” bead that further activates ﬂuorophores contained within the same bead. The ﬂuorophores subsequently emit light at 520–620 nm. In the absence of a speciﬁc biological interaction, the singlet state oxygen molecules produced by the “donor” bead go undetected without the close proximity of the “acceptor” bead. AlphaScreen has been used in GPCR functional assays for detection of endogenous cAMP production. In all the above-mentioned systems, the cells have to be subjected to lysis prior to measurement, adding not only an extra step to the assay procedure, but more importantly making kinetic measurements impossible because data can only be generated from endpoints. In addition, whole-cell FRET methods have been developed to determine the temporal and spatial dynamics of cyclic nucleotides (see Section 4.4). 2.1.3 Enzyme Fragment Complementation (EFC) Technology EFC is based on genetically modiﬁed enzymes consisting of two fragments, i.e. the enzyme acceptor (EA) and the enzyme donor (ED). The separated fragments are inactive and only spontaneous formation of heterodimers will lead to an enzymatically active form [27]. The HitHunterTM technology (DiscoveRx) is based on a modiﬁed version of the β-galactosidase enzyme forming an EA/ED pair. The technology has been adapted for use in many different assays including, but not limited to, kinase, progesterone receptor, estrogen receptor and cAMP assays. Here, we only discuss speciﬁc application for intracellular detection of cAMP. In brief, an ED–cAMP peptide conjugate is utilized, in which cAMP is recognized by a speciﬁc antibody. This ED fragment is able to complement EA to form an active complex, but only in the absence of the speciﬁc antibody. In the assay, anti-cAMP antibody is titrated for complete inhibition of enzyme formation. Exposure of a deﬁned antibody aliquot to cell lysate will lead to antibody–cAMP complex formation, therefore reducing the free amount of antibody. Subsequently, addition of ED conjugate in the following assay step will lead to reduced antibody–ED complex formation, therefore promoting complementation of active enzyme. Free ED–conjugate in the assay is proportional to the concentration of intracellular cAMP and the same applies for the measured β-galactosidase activity (see www.discoverx.com for additional information). 2.2 Membrane Transport Proteins

A great number of substances are transported in and out of complex organisms, in some cases using passive transport, driven solely by diffusion down a concentration

2 Membrane Proteins and Fast Cellular Responses

gradient either through the paracellular or transcellular route. Absorption and distribution of many food constituents such as ions, sugars, amino acids and nucleotides in the body are mediated/facilitated by speciﬁc membrane transport proteins, which form channels across cellular membranes. Similar mechanisms are needed for active removal of xenobiotics, waste products and cell metabolites from body tissue. These speciﬁc proteins are divided into two classes, i.e. uniporters and coupled transporters, of which the former transport a single solute from one side of the membrane to the other and, for the latter, the transfer of one solute depends on the simultaneous or sequential transport of a second solute. Coupled transporters are further divided into symporters moving compounds in the same direction and antiporters moving the compounds in opposite directions. Some speciﬁc transporters act in a passive manner moving molecules down a concentration gradient, as is the case with many nutrients such as glucose, which are found at high concentrations in the extracellular space of tissues. A similar situation is found with ligand- and voltage-gated ion channels of the nervous system, which speciﬁcally pass ions down a concentration gradient upon stimulation by neurotransmitters or membrane depolarization, respectively. In contrast, active transporters are linked to hydrolysis of ATP and move compounds in a highly speciﬁc manner against a gradient. The largest known and most diverse group of active transport proteins is the ABC transporter superfamily [28]. Over 50 ABC transporters have been described and the variety of substrates transported is large, including amino acids, sugars, inorganic ions, polysaccharides, peptides and even large proteins. One of the most cited ABC transporters, the multidrug-resistance (MDR1) protein [29], has been in the limelight of many drug discovery efforts in recent years, since overexpression in human cancer cells can render these cells simultaneously resistant to a variety of chemically unrelated drugs that are widely used in cancer chemotherapy. Other important phenomena linked to ABC transporters include resistance to chloroquine in malaria parasites, presentation of antigenic peptides on immune cells and the genetic disease known as cystic ﬁbrosis [30–32]. 2.2.1 Ion Channels Most cells maintain an electropotential across the membrane which has been most extensively studied in neurons and muscle cells where the potential is particularly high, and reaches a value of about −70 mV. The membrane potential is maintained by the ubiquitous Na+ -K+ -ATPase, which operates as an antiporter actively pumping Na+ out of the cell, while pumping K+ into the cell. For every molecule of ATP hydrolyzed, three Na+ are pumped out and two K+ pumped in, therefore creating a negative net charge inside the cell. Ion channel proteins form pores in cell membranes, allowing certain ions to pass through driven by a concentration gradient. A characteristic feature of ion channels is a gating mechanism that controls the movement of ions. Here we focus on ion channels, which are expressed in neurons or muscle cells, and therefore play important roles in neuronal signaling and muscle contraction. Changes of the membrane potential can be induced in a spatially limited fashion by ligand-gated Na+ channels

11

12 Cellular Assays and Their Application in Drug Discovery

sensitive to neurotransmitters. Subsequent opening of voltage-gated Na+ channels induces self-propagation of the depolarization along the cell membrane (action potential), ﬁnally inducing the opening of voltage-gated Ca2+ channels typically at the synapse (end plate). Inﬂux of Ca2+ induces the release of neurotransmitters into the synaptic cleft, further propagating the signal on to the post-synaptic cell. Malfunctions of ion channels are the basis of many ailments, including muscle and cardiac disorders, epilepsy, migraine, depression, and ataxia. Opening of ion channels is in most cases monitored by measuring changes in the membrane potential. The gold standard for this purpose is the patch clamp technology, offering very high sensitivity, good temporal resolution and measurement of single channel conductance at a precise voltage. To date this technology provides the best data for functional assessment of ion channels; however, the massive disadvantages include requirement of skilled operators, laborious work ﬂow and low throughput. Currently, different companies are involved in designing novel devices to achieve automation of the patch clamping procedure. One of the most advanced tools is the IonWorksTM station (Molecular Devices, Sunnyvale, CA), which uses integrated ﬂuidics for automatic positioning of single cells on microapertures on specially fabricated supports [33]. Single ion-channel recordings were demonstrated at a throughput of up to 2000 assay points per day. Higher throughputs can be achieved by using solvatochromatic, cell-permeable dyes like DiBAC (Molecular Probes, Eugene, OR) or the more recently developed FMP dye (Molecular Devices). Combination with a FLIPR system allows monitoring of ion channel function in true high-throughput mode surpassing analysis of 30,000 compounds per day (for more details, see the following paragraph). More details including other well-established ion-channel assay technologies were described in a recent review article [34].

2.2.1.1 FLIPR Technology and Ion Channel Assays To identify compounds that modulate ion channel activity, it is imperative that rapid and economical evaluation of biological activities in a high-throughput manner is available. One of the ﬂuorescence-based assays of membrane potential changes uses a potentiometric ﬂuorescent probe, bis-(1,3-dibutylbarbituric acid) trimethine oxonol [DiBAC4 (3)], that partitions between the intracellular and extracellular compartments depending on the electropotential across the membrane. Upon depolarization of the cells, the dye moves into the cell, and subsequent binding to intracellular hydrophobic sites results in enhanced quantum yield and in an increase in ﬂuorescence intensity. Conversely, membrane hyperpolarization triggers dye extrusion from the cells and consequently decreases ﬂuorescence, because the quantum yield of DiBAC4 (3) is low in an aqueous environment. The partitioning of DiBAC4 (3) is a slow process and therefore the technology is not amenable to measure channels with fast-gating kinetics or with rapid desensitization. Signiﬁcant improvement was achieved with the recent development of the FMP dye [35, 36], which has faster partitioning kinetics. Figure 4 shows a typical experiment conducted with differentiated myotubes (C2C12 cell line).

2 Membrane Proteins and Fast Cellular Responses

Agonist

1

2

3

4

5

6

7

8

9

13

Seed C2C12 Cells into 384-well Plates

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24

A B C D

Differentiate into Myotubes

E F G H I J K L M

Load Cells with FMP-dye

N O P

Screen LOPAC Library Antagonist

Agonists

NO

YES EC50 Determination And Specificity Check Decreasing conc. of agonist

Pacuronium addition

Fig. 4 Ion-channel activity in differentiated myotubes. The murine C2C12 muscle cell line was used to validate the FMP dye. C2C12 cells were seeded into 384-well plates and differentiated into myotubes by exposure to horse serum. Differentiated C2C12 cells strongly express nAChR and depolarization of the cells can be induced

Stop

NO

Antagonists YES

IC50 Determination

Decreasing conc. of antagonist

Carbachol addition

with speciﬁc receptor agonists, such as carbachol, which was applied in the outlined experiment. A 384-well plate from a typical screen for agonists and antagonists is shown. Example of an agonist and an antagonist are depicted as enlarged graphs. The 100% and 0% controls are overlaid as red- and blue-hatched lines, respectively.

14 Cellular Assays and Their Application in Drug Discovery

The LOPAC library (Sigma-Aldrich, St Louis, MO) of pharmacologically active compounds was screened for identiﬁcation of compounds exerting agonistic or antagonistic effects on the nicotinic acetylcholine receptor (nAChR). In brief, the cells were washed with buffer and loaded with the FMP dye for 30 min at room temperature. Six hundred and forty compounds from the LOPAC library were added in the FLIPR followed by stimulation with carbachol. In contrast to a typical Ca2+ signal, the membrane depolarization does not show a transient kinetic. However data evaluation and hit identiﬁcation can be carried out as described in Fig. 3. Identiﬁcation of agonists and antagonists is possible in one experiment. Positives were subjected to EC50 /IC50 determination and, in the case of agonists, the speciﬁc AChR antagonist pancuronium was used to show reversal of the carbachol signal (see ﬂuorescence traces shown for the EC50 determination, Fig. 4). Initially 21 compounds were identiﬁed as agonists and 12 compounds showed speciﬁcity for the AChR. 2.2.1.2 Voltage Ion Probe Reader (VIPR)TM Technology and Ion Channel Assays VIPR technology (Aurora Bioscience, San Diego, CA) is based on FRET and uses two ﬂuorescent molecules. The ﬁrst molecule, oxonol, is a highly ﬂuorescent, negatively charged, hydrophobic ion that can rapidly redistribute between two binding sites on opposite sides of the plasma membrane in response to changes in membrane potential. The second ﬂuorescent molecule, coumarin lipid, binds speciﬁcally to one face of the plasma membrane and functions as a FRET donor to the voltagesensing oxonol acceptor molecule. When the oxonol moves to the intracellular plasma membrane binding site upon depolarization, FRET is decreased, and this results in an increase in the donor ﬂuorescence and a decrease in the oxonol emission. This technology enables research and assay development in the ﬁeld of ion channel biology. VIPR technology can deliver subsecond, real-time readouts from assays in a microplate format (see www.aurorabio.com/ for additional information). 2.2.2 MDR Proteins MDR is mainly linked to overexpression of proteins of the ABC transporter superfamily. The most extensively studied member is the MDR1 gene product (Pglycoprotein), which is expressed in the intestine, kidneys, liver and blood-brain barrier, as well as in many tumor cells. In the latter case, high overexpression is one of the hallmarks of cell resistance to cancer chemotherapy. This observation is a result of anticancer drugs being actively pumped out of cells, resulting in decreased intracellular drug levels. Small molecules capable of reversing efﬂux can restore drug sensitivity in resistant tumor cells. The most commonly used assays to identify MDR pump inhibitors are based on ﬂuorescent P-glycoprotein substrates such as Hoechst 33342, rhodamine 123 and rhodamine 6G. The cells are loaded with the substrate and extracellular dye is removed by washing prior to lysis with detergents [37]. Subsequent measurement of ﬂuorescence emission provides a quantitative readout, which is proportional to the intracellular accumulation of the substrate. Exposure of the cells to MDR pump inhibitors will reduce efﬂux of the substrate

3 Gene and Protein Expression Proﬁling in High-throughput Formats

and therefore leads to an increased ﬂuorescence readout. However, these assays are not homogeneous and involve washing steps after loading of the cells with the corresponding dye. A novel MDR pump substrate was recently developed to run fully homogeneous assays on a FLIPR system. The FLIPR Multidrug Resistance Assay (Molecular Devices) is a fully homogeneous application needing only 15 min of incubation at room temperature with a proprietary dye, which is transported by the MDR1 gene product and changes its ﬂuorescent properties only after modiﬁcation by intracellular enzymes. This allows performance of live-cell kinetic assays for screening MDR pump inhibitors in high-throughput mode. The elimination of wash steps and temperature control means that the assay is highly amenable to automation [37].

3 Gene and Protein Expression Proﬁling in High-throughput Formats

Gene and protein expression proﬁling can be applied in drug discovery for at least two different purposes. The ﬁrst application will be for the discovery of new potential drug leads. In this case, the target of interest is either a component of the expression machinery itself or monitoring of expression serves as an indicator for an upstream target and reﬂects the coupled cellular response [16]. To make high-throughput screening (HTS) possible, the promoter sequences of interest, that have been shown to be coupled to upstream cellular events, are linked to so-called reporter genes, which serve as indicators for transcriptional and translational activation. The second application is the use of expression proﬁling to obtain information on the speciﬁcity of a drug candidate at the cellular level or for the identiﬁcation of surrogate markers for drug action. In this case, gene or protein expression in response to a potential drug candidate of as many genes and proteins as possible should be investigated. Expression patterns can be used as an indicator for the number of interactions within the cell. Such data could be of relevance for getting an understanding of the broader impact of modulating a potential drug target on the whole cellular system. Gene expression chips have been introduced recently for the measurement of hundreds of simultaneously expressed genes at the mRNA level [38]. The utility of the approach has been recently validated in yeast, where expression proﬁles identiﬁed pathways of previously uncharacterized mutations or identiﬁed new potential drug targets [39]. Similarly, protein chip arrays have been introduced to capture a particular fraction of proteins and allow analysis with a reasonable sample throughput [40]. In addition, the technology to analyze the cellular proteome, or parts of it, has been recently introduced [14]. The analysis of thousands of samples becomes a prerequisite if expression proﬁling is to be used in hit identiﬁcation. Hit-to-lead selection and lead optimization will require a throughput which is at the level of hundreds to thousands of compounds. Both can be achieved by focusing on selected indicator genes or their products,

15

16 Cellular Assays and Their Application in Drug Discovery

indicative of the discovery target of interest, rather than on broad expression proﬁling. Several targets can be addressed at once by running established single-readout assays in parallel or by using multiplexing as a tool to combine several readouts in one assay step [41]. In the following section, we will focus on examples that can be applied in early drug discovery and which allow a high sample throughput. Gene expression chips [38, 42] and proteomic technologies [43] are the subject of many recent reviews. However, due to their still limited sample throughput, they will only be mentioned brieﬂy here. 3.1 Reporter Gene Assays in Lead Finding

Transcription factors can be classiﬁed according to their structural nature, but also according to their behavior to respond to modulation of signal transduction pathways. Such a classiﬁcation, which is the basis for reporter gene technology and expression proﬁling, has recently been published by Brivanlou and Darnell [44]. According to this classiﬁcation, transcription factors can be grouped into two large classes – those who are constitutively activated, and those whose activity is regulated in a spatio-temporal, quantitative and qualitative manner. Among the regulated transcription factors is the large class of signal-dependent factors, which are activated upon an intra- or extracellular signal. Genes regulated via the latter are attractive as reporter genes since they couple the signaling pathways with a transcriptional response. Extracellular signals, after interacting with appropriate receptor molecules at the cell surface, initiate a cascade of intracellular events which often leads to alterations in the expression of a set of genes. One or several of these genes can be used as an indicator for the modulation of targets within speciﬁc signaling pathways. The activation of a certain gene can be easily identiﬁed by “tagging” the particular gene with a so-called reporter gene. For this, plasmids need to be constructed which contain the promoter or regulatory region of the gene of interest linked to the reporter gene. These plasmids are then introduced into a cell line which expresses the pathway of interest containing the pharmacological targets and couples the reporter readout with the particular pathway of interest. A number of genes have been described that can serve as reporter genes [16, 45]. Expression can be monitored by ﬂuorescence, e.g. by using green ﬂuorescent protein (GFP) as a reporter, or through the detection with ﬂuorescently labeled antibodies. Alternatively, the enzymatic activity of the reporter is used, as it is the case for luciferase, which converts the substrate to a luminescent product, or the reporter is detected with well-established and characterized secondary readouts producing ﬂuorescence, luminescence or colored reactions. Reporter gene technology has been widely used to establish high-throughput screens for a whole variety of targets of interest in drug discovery (for a review, see [45]). For many targets, such as transmembrane receptors for which other screening methods were difﬁcult to establish, reporter gene technology became the method of

3 Gene and Protein Expression Proﬁling in High-throughput Formats

choice for ﬁnding leads. Selected examples include the use of reporter gene assays to ﬁnd agonists and antagonists for a whole variety of GPCRs [17] stably transfected into Chinese hamster ovary (CHO) cells with a 6 × CRE–luciferase reporter construct. This cell line served as an indicator for GPCRs, such as the melanocortin-1 receptor, which upon stimulation modulates cAMP levels in the cell. The assay can be performed in 384-well plates, therefore offering signiﬁcant savings in cost, time and cell culture efforts. Along the same lines, a HTS system was established for the metabotropic glutamate receptor mGluR7 [46]. The mGluR7 receptor is negatively coupled to adenylyl cyclase and its stimulation decreases cAMP levels. CHO cells were stably transfected with rat mGluR7 cDNA and an AMP-responsive luciferase reporter gene. Suitability for HTS was demonstrated in a line where cAMP levels induced by forskolin (an activator of adenylyl cyclase) were signiﬁcantly reduced by receptor agonists. A direct method to assay receptors negatively coupled to adenylyl cyclase has been described by Xing et al. [47]. The stably transfected CHO reporter cell line contained three separate expression plasmids: (i) a chimeric Gq/i -protein, which redirects a negative Gi signal into a positive Gq signal; (ii) a Gi -coupled GPCR, in this case the µ-opioid receptor or the 5-hydroxytryptamine receptor, and (iii) the 3 × NFAT-β-lactamase reporter. Thus activation of the Gi -coupled GPCR translated into NFAT activation via the Gq -mediated signaling pathway, allowing the monitoring of β-lactamase expression in a HTS format. A wide variety of different cell lines have been used to establish reporter gene assays, a selection of which is indicated here as examples. Human embryonic kidney (HEK) 293-EBNA cells grown in suspension culture were used to assay expression of prostanoid receptors coupled to a CRE-SEAP (secreted human placental alkaline phosphatase) reporter [48]. ECV304 cells, a spontaneously transformed immortal line derived from human umbilical vein endothelial cells, were transfected with an ICAM-1 reporter to screen for compounds which inhibit ICAM-1 expression upon stimulation with interleukin (IL)-1β [49]. The human breast cancer cell line MDA-MB-453 was used as a transfection host to establish stable reporters under the control of a mouse mammary tumor virus (MMTV) promoter to study androgen and glucocorticoid receptor agonists. MDA-MB cells express both the androgen and the glucocorticoid receptors, and therefore serve as an ideal host cell for the reporter constructs [50]. As a practical example, data obtained with a reporter gene assay under HTS conditions is presented in Fig. 5 and Tab. 2. Cells stably transfected with a luciferase reporter gene construct were seeded in 384-well plates and dosed with serial dilutions of a compound known to activate the promoter of the reporter gene construct. After an incubation time of 24 h, the cells were washed, lysed and the luciferase subR ; PerkinElmer Life Sciences) was added. The amount of luciferase strate (LucLite protein expressed correlated directly with the amount of substrate metabolized to a luminescent compound, which was detected in a luminometric plate reader R NXT; PerkinElmer Life Sciences). The data presented here provided (TopCount information as to the minimal concentration of positive reference compound required as standard on each plate for the HTS of 100,000 compounds. To assess the quality of each individual assay plate during HTS, a criterion called the Z factor

17

18 Cellular Assays and Their Application in Drug Discovery 45000 40000

Luminometric counts

35000 30000 25000 20000 15000 10000 5000 0 0

0,1

0,3

1

3

10

30

Concentration of inducer (µM)

Fig. 5 Dose-dependent induction of luciferase reporter gene activity by an inducing substance subsequently used as an agonist control in HTS. Each value represents the average of eight individual determinations, the error bars indicate 1 SD. The mean of luminometric counts measured

with TopCount NXT (PerkinElmer Life Science) is plotted against the concentration of the inducing substance (in :M). Counts obtained in the absence of compound refer to the basal expression of the luciferase reporter gene and constitute the assay background.

is calculated from the dynamic range of the signal (difference between the means of the background and the positive control wells) and the standard deviation (SD) of the control wells on each plate [51]. Since the Z factor takes into account both the signal intensity and the inherent variance, it gives a measure of the stability of an assay. Screening is generally considered possible if 0 < Z < 1 [51]; however, in practice, the minimum value to be met by the Z factor is often set higher. As shown in Fig. 5 and in the values calculated in Tab. 2, a concentration of at least 3 µM of the agonist in the positive control would be required to obtain a Z > 0.5. In effect, the positive control was used at 10 µM during HTS. Table 2 Luciferase reporter gene activation and assay performance

Concentration (l M)

Mean of 8 wells

SD

Z factor

0 0.1 0.3 1 3 10 30

6364 6347 9028 12006 20436 31292 38343

641 314 458 509 1476 1181 2027

n.d. -0.238 0.389 0.549 0.781 0.750

3 Gene and Protein Expression Proﬁling in High-throughput Formats

Reporter gene assays can be highly informative if the gene product of interest and the pathway of induction are well characterized. It needs to be borne in mind that the reporter gene will be modulated by any type of stimulus inﬂuencing any step of the coupled signal transduction cascade. Therefore, the effects on reporter gene expression observed in response to test compounds should always be studied further in assay panels addressing speciﬁc steps in the pathway, to rule out mechanisms not directly associated with the target of interest. This has been shown recently by inhibiting the CXCR1-agonist-stimulated mitogen-activated protein (MAP) kinase reporter gene by pertussis toxin (acting at the G-protein level) and inhibitors of one MAP kinase subfamily, but not by inhibitors of a second subfamily [52]. Even though expression of the reporter genes generally does not inﬂuence homeostasis of the cell, a certain artiﬁcial component is introduced into the system. First and foremost, it needs to be established whether the signal transduction pathway leading to activation of a particular promoter is present and functionally active in the cells in question. The introduction of the reporter gene itself into cells can disturb normal transcription patterns, due to scavenging of particular transcription factors or through positional effects due to integration of the plasmid during the generation of stable cell clones. Thus, great care has to be taken in choosing the promoter and the cell line in which the reporter gene assay is to be established. Limitations can occur regarding the cell lines that can be used, the transfection methods that should be employed and the efﬁciency of expression of the reporter gene. Nevertheless, even though the generation of well-characterized reporter gene cell lines is time-consuming and labor intensive, they constitute a highly efﬁcient system for HTS programmes.

3.2 Reporter Gene Assays in Lead Optimization

The use of gene expression arrays to monitor simultaneously the expression of hundreds or thousands of genes has been mentioned before. Expression arrays are useful tools not only to characterize the expression phenotypes of cells and tissues for their classiﬁcation [53], but they also allow us to monitor the impact of mutations [54] or drugs on the cellular phenotype [55] at a systems level. Arrays are widely used in toxico- and pharmacogenomic studies [56, 57]. Furthermore, the large data sets obtained with gene expression arrays have been used to construct predictive models for diagnostic purposes [58, 59]. The use of expression arrays requires the production of probes suitable for hybridization, which can be fastidious to produce. Furthermore, analysis of the large data sets obtained requires sophisticated software, and a broad special knowledge as to the signiﬁcance and relevance of the multiplicity of the responses observed. In some instances, only a particular fraction of all genes measured is informative for the target in question. Thomas et al. [60] have recently analyzed the expression of 1200 liver transcripts using cDNA arrays, after dosing mice with 24 different

19

20 Cellular Assays and Their Application in Drug Discovery

treatments falling into ﬁve toxicological categories. A set of 12 transcripts was identiﬁed and considered diagnostic for classiﬁcation of the treatments into the toxicological classes. Interestingly, addition of further transcripts lead to a decline in the predictive accuracy, and the “individuality” of the speciﬁc treatment proﬁle began to take over. In a similar study, Bartosiewicz et al. [61] have used a set of 260 genes involved in phase I and II drug metabolism to test the effect of 10 toxicants on expression proﬁles in different tissues and for different exposure times in mice. Distinct groups of chemical classes could be correlated to the speciﬁc expression proﬁle of a relatively low number of transcripts. Similar expression proﬁling studies performed on HepG2 cells [62] and on livers of treated rats [63] show that even with a highly focused set of genes, separate classes of chemicals can be identiﬁed through shared modulation of expression of the selected gene set. Today, the method of choice to assess gene expression proﬁles in vivo consists in the use of oligonucleotide or cDNA microarrays or gene chips. However, due to the high cost involved and the labor-intensive sample preparation, which generally preclude a high-throughput method, these studies are done mostly in a late phase of the drug discovery process. Thus, one or a few compounds are analyzed for the induced expression patterns of many genes. To reduce late-phase attrition of potential drug candidates due to unforeseen side-effects, expression-proﬁling systems which are amenable to HTS are needed. An approach making use of reporter gene assays has been described [64, 65]. Albano et al. [65] selected a set of oxidative stress-responsive promoters from Escherichia coli genes to generate GFP–reporter gene constructs, as a tool for mechanistic screening of antitumor drugs in 96-well format. Zhu and Fahl [66] generated stable HepG2 cell clones expressing GFP under the control of the electrophile response element (EpRE), which allow HTS for induction of phase II drug-metabolizing enzymes. In addition to high-throughput, a further advantage of GFP–reporter gene constructs is that the expression can be detected in live cells, allowing easy monitoring of time courses and reversibility of induction. The use of reporter gene cell lines for expression proﬁling has further been documented by Farr and Todd [64], who established a panel of cell lines containing the bacterial chloramphenicol N-acetyltransferase (CAT) reporter gene under the control of promoters indicative of speciﬁc cellular stress responses. Induction of reporter constructs by chemical compounds can give an indication concerning the action of compounds at the subcellular level. Thus, promoter–reporter gene constructs expressed in well-characterized cell lines can provide useful model systems to proﬁle compounds with respect to activation of canonical signal transduction pathways associated with deﬁned cell fates or phenotypes. The assay system will detect compounds which modulate reporter gene expression directly and those which interfere with the expression elicited by external stimuli such as growth factors or cytokines. When a panel of reporter gene assays is used for compound proﬁling, similar expression patterns can point to comparable mechanisms of action of test compounds in intact cells. Changes of expression patterns in response to speciﬁc signaling pathways and in different cell lines can provide information on the speciﬁcity of test compounds. These systems can easily be adapted to HTS, and the analysis of time

3 Gene and Protein Expression Proﬁling in High-throughput Formats

course and dose dependency of induction. Therefore, the proﬁling of hit collections or compound sets resulting from lead optimizations might allow the clustering of compounds into groups with pharmacologically similar mechanistic responses. For example, activation of reporter genes coupled to the p53-responsive element (p53RE) [67] would indicate that the test compound leads to DNA damage, and activation of reporters under the control of the xenobiotic response element (XRE) [68, 69] would point to the transcriptional induction of drug-metabolizing enzymes, such as cytochrome P450s and others. Thus, in vitro reporter gene proﬁling with a well-deﬁned and validated set of genes can contribute not only to compound classiﬁcation, but also to the identiﬁcation of possible side-effect proﬁles of the compounds tested. More recently, with the advance of novel and more sensitive techniques of monitoring gene expression, an entirely new panel of assays has been brought forward. While reporter gene assays were devised with the idea of visualizing a particular gene expression event through the coupling to a protein which can be easily monitored, the novel techniques directly monitor gene expression by quantitation of the mRNA levels for several to up to thousands of genes simultaneously. Furthermore, these techniques do not require the prior introduction of a detection system, allowing unlimited choice with respect to cell type or tissue derived from organs. Table 3 gives an overview of a number of techniques developed for HTS applications that are currently available. This list is by no means complete, but should serve as a general reference to the different approaches taken. With respect to the HTS suitability of these novel technologies, two major categories can be differentiated. There are methodologies like serial analysis of gene expression (SAGE) [70], GeneCalling (transcript proﬁling by restriction endonuclease ﬁngerprinting) [71] and total gene expression analysis (TOGA) [72], where initial sample preparation is very fastidious, but where high-speed parallel detection of many different genes can be achieved. These techniques are collectively called “open” systems – no prior sequence information is required and virtually all genes induced by a particular compound can be detected. Open systems challenge the DNA microarray technology, providing an alternative to the high cost and high demand of starting material associated with the latter. In practice, they will prove useful if only a few samples need to be analyzed, but where the high throughput (HT) refers to the detection of the different gene products. Quantitative reverse transcription-polymerase chain reaction (RT-PCR) [73], the branched DNA technology promoted by Bayer (Emeryville, CA) [74, 75] and the high-performance signal ampliﬁcation (HPSATM [76]) system promoted by Chromagen (San Diego, CA) can detect one or a few mRNAs at a time. They are called “closed” system technologies, as are the DNA and oligonucleotide microarrays and the reporter gene technology, since they can be applied only to genes with known sequence. However, these methodologies can be used for HTS, since the assays can be run in a microtiter plate format, allowing the simultaneous processing of many samples. Moreover, they provide an advantage to the traditional reporter gene

21

22 Cellular Assays and Their Application in Drug Discovery Table 3 Differential gene expression technologies (adapted from [155])

Technology

Methodology

Target genes

Open systems: no prior sequence information required HT-compatible SAGE transcript capture by “ﬁngerprint” SAGE tags, identiﬁcation and quantitation by sequencing of serial concatemers 96 reactions GeneCalling restriction (transcript proﬁling) endonuclease “ﬁngerprinting” of cDNA, conﬁrmation by competitive PCR TOGA PCR-based cDNA 256 reactions display, identiﬁcation of mRNAs with 3 poly(A) tail by sequencing Closed systems: analysis of genes with known sequence information HT-compatible hybridization of total DNA microarray/oligoarrays RNA to gene probes arrayed on silicone chips or glass slides Quantitative RT-PCR PCR-mediated release few probes (TaqmanTM ) of gene-speciﬁc ﬂuorogenic probes; threshold of detection is inversely proportional to concentration Branched DNA 1 probe direct mRNA (QuantigeneTM ) detection by hybridization with gene-speciﬁc linker and enzyme-linked ampliﬁcation probes 2 probes HPSATM direct mRNA detection by hybridization with gene-speciﬁc capture and enzyme-linked ampliﬁcation probes

Samples

References

few

70

few

71

few

72

few

156, 157

96-well format

73

96-well format

74, 75

96-well format

158

format, because they are more ﬂexible and can be performed on virtually any source material, and the detection of a novel gene requires only the design and synthesis of gene-speciﬁc probes. These novel technologies promise increased sensitivity, with a readout not compromised by secondary effects due to the reporter gene, but they are not yet as widely established as the reporter gene systems.

4 Spatio-temporal Assays and Subpopulation Analysis 23

4 Spatio-temporal Assays and Subpopulation Analysis

Cellular assays addressing protein expression levels, spatial and temporal distribution of proteins within the cell, and intracellular signaling dynamics are currently exploited to better understand pharmaceutical target functions in cells. To date, there is an increasing number of labeling reagents and ﬂuorescent probes for imaging applications, including target-speciﬁc antibodies, complementing enzymes [77, 78], dyes (e.g. Alexa Fluor dye series; Molecular Probes), afﬁnity reagents [79] and genetically encoded ﬂuorescent proteins [18, 80, 81]. Systems that allow high-throughput analysis of cells include ﬂow cytometrybased devices, such as the recently described high-throughput pharmacological system [82]. This system allows multifactorial characterization of cells, but does not allow us to quantify spatial rearrangements inside cells. Recently developed microscopic imaging technologies have broad applications. They allow analysis of a wide range of cellular functions including cell motility, cell morphology, protein movement inside cells, protein localization, intracellular second messengers and endpoint measurements of signaling cascades. The development of automated imaging systems has made compound screening in high-content format possible. The great advantage of automated imaging and scanning techniques is the tremendous increase in sample throughput compared to manual microscopy. In the case of ﬂuorescence microscopy, multiple ﬂuorescent probes can be detected and quantiﬁed simultaneously. R The ArrayScan II system was introduced in 1999 by Cellomics [83] (http://www .cellomics.com) and was one of the ﬁrst automated imaging instruments for highcontent screening on the market. Meanwhile, other instruments, software and hardware were developed by several companies including Acumen-Bioscience (http://www.acusite.co.uk), Amersham Bioscience (http://www.amersham.com), Applied Biosystems (http://www.appliedbiosystems.com), Q3DM (http://www .q3dm.com), CompuCyte (http://www.compucyte.com), Kinetics Imaging (http:// www.kinetikimaging.com), Universal Imaging (http://www.imagel.com), Axon Instruments (http://www.axon.com) and Evotec OAI (http://www.evotecoai.com). The rapid progress in the development of analytical devices for subcellular analysis in combination with labeling reagents for speciﬁc identiﬁcation of the molecules of interest will allow us to analyze speciﬁc signaling and metabolic steps in a highthroughput mode. 4.1 Phosphorylation Stage-speciﬁc Antibodies

Of particular interest for automated imaging are activity-dependent reagents, such as phosphospeciﬁc antibodies that allow selective analysis of signaling pathways within cells. Phosphorylation and dephosphorylation of proteins by kinases and phosphatases, respectively, are part of a complex signaling network of tightly regulated dynamic processes. Protein phosphorylation frequently results in sub-cellular

24 Cellular Assays and Their Application in Drug Discovery

redistribution of key signaling molecules, which is critical for their biological activity [84–86]. The spatial distribution of two key signaling proteins upon phosphorylation, extracellular signal-regulated kinase (Erk) and p38 MAP kinase, was analyzed with phosphotyrosine-speciﬁc antibodies recognizing only the phosphorylated form of the protein [87]. Exposure of cells to a potent Cdc25 phosphatase inhibitor resulted in a concentration-dependent nuclear accumulation of phospho-Erk and phospho-p38, but not nuclear factor-κB (NF-κB). These experiments revealed that Cdc25 inhibition increased Erk phosphorylation and nuclear accumulation, and indicated that the phosphorylation status of Erk is regulated by a Cdc25 phosphatase. A phospho-histone H3 antibody that targets the phosphorylation site on Ser10 of histone H3 and a phospho-histone H1 antibody were used to detect mitogenand oncogene-stimulated phosphorylation in small nucleosomal regions that are sensitive to hyperacetylation [88, 89]. Active kinase states have been analyzed with a panel of phosphospeciﬁc monoclonal anti-kinase antibodies. The speciﬁcity of the antibodies for the speciﬁc kinase has been veriﬁed by Western blot analysis and with speciﬁc inhibitors for upstream kinases. Using multiparameter ﬂow cytometry, Perez and Nolan [90] could identify lymphocyte subsets with as many as four different intracellular active kinases. This technique allows us for the ﬁrst time to investigate simultaneously different active kinases in one individual cell. While this technology can be best applied to cells in suspension, high-throughput microscopy is the technology of choice for adherent cells. In addition microscopic applications allow analysis of intracellular spatial distributions which is not possible with ﬂow cytometry. Mitotic cells are characterized by an increase in phosphorylated nucleolin, a nuclear protein phosphorylated in cells entering mitosis. Using a monoclonal antibody that speciﬁcally recognizes phosphorylated nucleolin, Mayer et al. [91] have developed a high-throughput whole-cell immunoblot assay. Compounds arresting cells in mitosis by novel modes of action could be identiﬁed after sorting compounds with secondary assays. Other protein modiﬁcations like acetylation can be detected with speciﬁc antibodies as well. The organization of transcriptionally active chromatin and activity of nuclear histone acetyltransferase and deacetylase was investigated by using an acetyl-histone H3-speciﬁc antibody [92, 93]. Compounds affecting ligandstimulated changes in chromatin structure can be screened with such antibodies by automated imaging techniques.

4.2 Target-protein-speciﬁc Antibodies

Many proteins play important roles in both the cytoplasm and nucleus. Therefore, they need to move from one compartment to the other in order to perform their functions. Typical examples are latent cytoplasmic transcription factors [44] or nuclear transport factors [94]. Assays for nuclear transport factors using heterokaryon formation in combination with ﬂuorescent proteins have been described [95].

4 Spatio-temporal Assays and Subpopulation Analysis 25

Nuc-CytoInten Difference

800 Exp.1

700

Exp.2

600 500 400 300 200 100 0 -100 0

20

40

60

80

Fig. 6 Time course of NF-6B nuclear translocation. HeLa cells were stimulated with 1 ng/ml IL-1" for the indicated times at 37◦ C. Mean values of 100 counted cells per well of two different experiments (Exp.1 and Exp.2) are shown. Nuc-CytoInten = nuclear minus cytoplasmic ﬂuorescence intensity.

The movement of latent cytoplasmic transcription factors upon activation to the nucleus can be monitored and quantiﬁed by automated ﬂuorescence imaging. IL-1induced and tumor necrosis factor-α-induced translocation of NF-κB to the nucleus has been quantiﬁed with the application of the Cellomics ArrayScan II technology [96]. Unstimulated cells contain inactive NF-κB distributed within the cytoplasm that can be detected by a NF-κB-speciﬁc antibody combined with a ﬂuorescently labeled secondary antibody. In IL-1α-stimulated cells, most NF-κB is activated and translocates to the nucleus in a time- and dose-dependent manner (Figs. 6 and 7). Upon IL-1α stimulation, HeLa cells were analyzed with the ArrayScan II system using the speciﬁc software that allows quantitative analysis of ﬂuorescence in the nucleus and in the adjacent cytoplasm. Cells are identiﬁed as objects by the software through the nuclear staining with Hoechst 33324 or DAPI, and are overlaid with masks for nuclear and cytoplasm ﬂuorescence measurements (Fig. 8). 4.3 Protein–GFP Fusions

Tagging technologies have become popular to monitor the movement or transport of molecules into cells and within cells. Extracellular ligands can be conjugated to biotin and later visualized with enzyme-conjugated streptavidin. A HTS assay

Nuc-CytoInten Difference

26 Cellular Assays and Their Application in Drug Discovery

1000 900 800 700 600 500 400 300 200 100 0 -100 0.0001

Exp.1 Exp.2

0.001

0.01

0.1

1

10

ng/ml IL-1α Fig. 7 Dose-response relationship of IL-1"-induced NF-6B nuclear translocation. HeLa cells were stimulated with indicated concentrations of IL-1" for 30 min at 37◦ C. Mean values of 100 counted cells per well of one block two different experiments (Exp.1 and Exp.2) are shown. Nuc-CytoInten = nuclear minus cytoplasmic ﬂuorescence intensity.

has been described on this basis for epidermal growth factor (EGF) internalization [97]. Fluorescent proteins signiﬁcantly contribute to the understanding of protein dynamics within the cell. They represent powerful tools and ideal tags to assess realtime analysis of molecular events in living cells in the form of reporters or fusion proteins. The most prominent ﬂuorescent protein is GFP, originally isolated and cloned from the jelly ﬁsh Aequorea victoria [98]. A number of GFP variants have been developed with different excitation and emission spectra, including enhanced blue ﬂuorescent protein (eBFP), enhanced yellow ﬂuorescent protein (eYFP) and enhanced cyan ﬂuorescent protein (eCFP). The nature of their structures renders these proteins extremely stable and resistant to degradation by most proteases [99]. This high level of stability does not necessarily represent an advantage for cellular applications. If changes of gene expression with transcription reporter assays are to be monitored, a shorter half-life of the reporter is desired. Therefore, short half-life forms of eGFP were developed in order to be used as transcription reporter proteins and these are termed destabilized eGFP [80]. Meanwhile, other ﬂuorescent proteins from marine invertebrates have been cloned, like the ﬂuorescent proteins from reef corals [81]. The combination of distinct spectral variants of ﬂuorescent proteins

4 Spatio-temporal Assays and Subpopulation Analysis 27

Fig. 8 IL-1"-induced NF-6B translocation in HeLa cells. Cells were stained with rabbit anti-p65 NF-6B antibody and goat antirabbit Alexa Fluor 488 antibody. Nuclear DNA was stained with Hoechst 33342. NF6B is quantiﬁed by measurement of green

ﬂuorescence intensity; nuclei appear blue. Unstimulated cells (left) show most green ﬂuorescence in the cytoplasm, whereas cells stimulated with 2 ng/ml of IL-1" for 30 min at 37◦ C (right) show translocation of NF-6B to the nucleus.

enables a wide range of applications, such as the simultaneous analysis of multiple gene expression cascades and localization or translocation of different proteins. Measurement of co-internalization of β-arrestin bound to GPCRs can be used to monitor receptor activation and recycling. Barak et al. [100] demonstrated, using confocal microscopy, the translocation of β-arrestin-2–GFP fusion proteins from the cytoplasm to membrane-anchored GPCRs. This was demonstrated for more than 15 different ligand-activated GPCRs in HEK-293 cells. In the absence of receptor activation, β-arrestins are distributed throughout the cytosol and are absent in the nucleus. Upon addition of ligand to the cells, β-arrestin-1–GFP fusion proteins translocate to the plasma membrane and further to endocytic vesicles [101]. The accumulation of the GFP-tagged proteins can then be quantiﬁed using a ﬂuorescence imaging technology. With a β-arrestin-1–GFP fusion protein, co-internalization of arrestin with thyrotropin-releasing hormone receptor 1 in response to agonist was monitored and imaged [102, 103]. NORAK Biosciences (Research Triangle Park, NC; http://www.norakbio.com) exploits arrestin-GFP movement under the term TransﬂuorTM technology as a broadly applicable, stable HTS platform to identify compounds modulation GPCR signaling. Receptor

28 Cellular Assays and Their Application in Drug Discovery

internalization and trafﬁcking was also exploited by the use of ﬂuorescently labeled ligands such as ligand of transferrin receptor [104]. High-content screening was conducted on the ArrayScan II system, and internalization and recycling of the transferrin receptor with bound ligand could be monitored and quantiﬁed. GPCR activation and internalization has also been monitored using GPCR-GFP ﬂuorescent protein fusions [104, 105]. Conway et al. [105] have used the ArrayScan high-content screening system to monitor parathyroid hormone receptor and β 2 adrenergic receptor internalization upon stimulation by speciﬁc ligands. Other examples where fusions with ﬂuorescent proteins were used to quantitate protein movement include nuclear translocation of calmodulin [106] or the membrane-bound transcriptional regulator tubby after phosphoinositide hydrolysis [107]. In other applications, translocation domains such as the Ca2+ -sensitive C2 domain of protein kinase Cδ have been fused to YFP and expressed in rat leukemia cells. Evanescent wave single-cell array technology has been applied as an imaging technology to monitor protein movement upon addition of platelet-activating factor [108]. Simultaneous analysis of mixed cell populations can be performed by transfecting each cell line with a single ﬂuorescent fusion protein. Vectors of BFP and YFP were used to stably transfect two different colon cancer lines, the parental line with a mutant K-ras gene and a variant where the mutant gene has been deleted. To screen for compounds that selectively kill only the tumorigenic variant with the mutant ras gene, both populations were mixed and simultaneously assayed for cell survival using ﬂuorescence intensity measurements [109]. Compounds that preferentially affect the cells with the mutant ras allele could be identiﬁed. 4.4 Fluorescence Resonance Energy Transfer (FRET)

FRET has become a popular method to study protein dynamics in cells [110]. FRET is based on the transfer of energy absorbed by one ﬂuorescent molecule to another, causing excitation of this second molecule. To achieve FRET, the two ﬂuorescent molecules with different excitation/emission spectra need to come into close proximity of each other. The most general strategy for creating such indicators is to sandwich the conformationally sensitive domain of interest between two mutants of GFP. In case of ligand binding or modiﬁcation of the domain, the two ﬂuorescent moieties get into close proximity, which enables FRET. In a pioneering study, Mahajan et al. [111] used plasmid constructs that expressed the fusion proteins GFP-Bax and BFP-Bcl-2. Bax and Bcl-2 are two key modulators of apoptosis, but their direct interaction had then never been shown at the cellular level. Using FRET, the direct interaction of the two proteins in mitochondria could be demonstrated. FRET has also been applied to determine kinase activities in living cells. For this purpose, reporters were constructed, consisting of fusions of CFP, a phosphoamino acid-binding domain, a consensus substrate for the relevant kinase of interest and YFP. In cells transfected with the reporter constructs and stimulated

4 Spatio-temporal Assays and Subpopulation Analysis 29

for the particular kinase activity, the fusion protein after phosphorylation underwent a conformational change, and allowed FRET between the YFP and CFP. The change in the ratio of yellow to cyan emission could be quantiﬁed, and temporally as well as spatially determined. This technology has been successfully applied for protein kinase A [112], and the three tyrosine kinases Abl, Scr and EGF-receptor (EGF-R) [113]. In a similar approach, Honda et al. [114] demonstrated the use of a genetically encoded ﬂuorescent indicator to investigate the dynamics of guanosine 3 ,5 -cyclic monophosphate (cGMP) in single cells. The cGMP indicator was constructed by bracketing deletion mutants of the cGMP-dependent protein kinase between cyan and yellow mutants of GFP. Binding of cGMP to the fusion protein indicator altered FRET and the ratio of cyan to yellow emissions selectively over cAMP. By using this method, the temporal and spatial dynamics of intracellular cGMP levels in rat fetal lung ﬁbroblasts and in primary rat Purkinje neurons was monitored upon addition of different stimuli. Similar sensors for cAMP have also been published [115]. FRET-based sensors were also used to demonstrate the spatio-temporal behavior of GTB-binding proteins after cellular stimulation with growth factors [116]. A fusion protein was engineered which contained H-Ras, the Ras-binding domain of Raf, and was ﬂanked by a pair of yellow and cyan mutants of GFP. H-Ras was replaced in a second construct by the analogue protein Rap1. Activation of the Ras/Rap pathway by growth factors induced FRET. Spatio-temporal localization of the fusion protein indicated that EGF stimulation of COS-1 cells activated Ras at the peripheral plasma membrane, but Rap-1 in the perinuclear region. In addition, this study showed that growth factors activate only a subpopulation of Ras and Rap, and only at very restricted areas in the cells. This technology in combination with cellular imaging will greatly increase the sensitivity and facilitate cellular HTS approaches for speciﬁc intracellular signaling events. 4.5 GPCR Activation using Bioluminescence Resonance Energy Transfer (BRET)

BRET is a naturally occurring process. The jelly ﬁsh A. victoria or the sea pansy Renilla reniformes create green ﬂuorescence by BRET. In the former blue-lightemitting case, aequorin associates with GFP, whereas in the latter case, the energy generated by degradation of coelanterazine by luciferase is transferred to GFP. This transfer, however, only occurs if luciferase and GFP form a heterodimer. BRET has been used to demonstrate the interaction of circadian clock proteins from cyanobacteria expressed in E. coli [117]. More recently, BRET has been applied to study GPCR activation by several groups [118–120]. To demonstrate β 2 -adrenergic receptor (β 2 AR) dimerization, β 2 AR-Renilla luciferase and β 2 AR-red-shifted GFP (YFP) constructs were coexpressed in HEK-293 cells. Occurrence of BRET was an indication of receptor dimerization. In the same study, β-arrestin, which is known to associate with the intracellular domain of activated G protein-coupled receptors, was expressed as an arrestin-YFP construct together with β 2 AR-Renilla luciferase. BRET between the two proteins occurred, but was entirely dependent upon receptor

30 Cellular Assays and Their Application in Drug Discovery

activation and indicated the association of arrestin with the intracellular domain of the β 2 AR in the active state of the receptor. This assay has been developed as a HTS assay technology by PerkinElmer Life Sciences. 4.6 Protein Fragment Complementation Assays (PCA)

The methods described above to study protein-protein interactions are associated with certain shortcomings. The yeast two-hybrid system is based on transcription, which requires proteins to be available in the nucleus. In FRET, ﬂuorescent proteins need to be expressed in cells at relatively high levels and the interaction must be such that the proteins are in close proximity to allow energy transfer. Enzyme complementation and its use in studying protein-protein interactions have been developed for E. coli β-galactosidase [121], dihydrofolate reductase (DHFR) [122] and most recently for β-lactamase [123]. In the case of β-galactosidase, two weakly complementing deletion mutants are each fused to the two proteins to be probed for interaction. Only if the two proteins interact are the two parts of the galactosidase forced to complement each other and exert enzymatic activity. The system has been developed by demonstrating FRAP and FKBP12 interaction in the presence of rapamycin. More recently, β-galactosidase complementation was used to establish a HTS system to screen for antagonists of EGF-mediated EGF-R dimerization [77, 124]. For this purpose, the complementing deletion mutants of β-galactosidase were fused to the intracellular domains of the EGF-R and cotransfected into C2C12 mouse myoblasts. Enzyme activity was reconstituted after adding EGF to the cells in culture. With this assay adapted to a 384-well format, primary hits identiﬁed in the EGF-R assay were screened for speciﬁcity with two complementing deletion mutants of β-galactosidase fused to human β 2 -adrenoceptor and β-arrestin-2. By the use of sensitive chemiluminescent and ﬂuorescent galactosidase reagents, this technology enables the measurement of protein-protein interactions in living cells for HTS applications. The reassembly of active DHFR from rationally designed fragments was established by Pelletier et al. in a prokaryotic system [122]. E. coli DHFR is selectively inhibited by the antifolate trimethoprim. Murine DHFR can rescue E. coli cells. Complementing fragments of murine DHFR fused to homo- or heterodimerizing proteins were used to transform E. coli, which were then grown in the presence of trimethoprim. Survival was taken as a measure for protein interactions and enzyme complementation. Besides GCN4 leucine zipper fusions, raf-ras interaction, and rapamycin-induced interaction of FKBP and TOR2 were used in survival studies to demonstrate interaction and complementation. In another study, protein partners fused to complementing fragments of murine DHFR were introduced into DHFR-negative mammalian cells grown in the absence of nucleotides. Cells where the proteins associate and DHFR can complement are able to survive under these conditions [125]. As an alternative, protein interaction and DHFR complementation can be monitored by feeding cells ﬂuorescein-conjugated methotrexate. Methotrexate only binds

5 Phenotypic Assays 31

to DHFR when the complementary fragments are coexpressed and reassembled. Fluorescence signals can then be monitored by ﬂuorescence microscopy, FACS or by spectroscopical devices used in HTS. The FKBP-rapamycin-FRAP complex and the activation of the erythropoietin receptor were used as model systems [125]. Most recently, DHFR-PCA assays were used to analyze whole biochemical networks in living cells [78]. Cells expressing associating protein pairs were selected and ﬂuorescein-conjugated methotrexate, which binds to reconstituted DHFR, was applied to localize interacting protein partners and to study inhibitors preventing the interaction. The same method has been successfully applied in plant protoplasts expressing complementary fragments of DHFR fused to interacting proteins. The binding of ﬂuorescein-conjugated methotrexate as measure of protein–protein interaction was monitored by ﬂuorescence microscopy [126].

5 Phenotypic Assays 5.1 Proliferation/Respiration/Toxicity

Many cytotoxicity and proliferation assays for mammalian cells rely on radioactivity and measure the incorporation of radiolabeled nucleotides such as [3 H]thymidine into cellular DNA. Although very sensitive, the [3 H]thymidine assay [127] is relatively expensive and labor intensive, and requires handling and disposal of the radioactive waste. Other cytotoxicity assays, such as MTT [3-(4,5-dimethylthiazol-2-yl)2,5-diphenyltetrazolium bromide] [128], Alamar Blue [129], lactate dehydrogenase [130] and ATP content [131], require the addition of speciﬁc reagents to generate a signal. Among those, the most commonly used is MTT [132]. This tetrazolium salt is reduced within the mitochondria of metabolically active cells to form a colored formazan dye precipitate. The broad utility of MTT is limited by the fact that it may be susceptible to interference from drugs, either by reacting with reducing groups on the drugs or by scatter of absorbance resulting from the precipitation of drugs absorbing light in the visible spectrum. Cell viability measurement by the MTT assay relies on its reaction with mitochondrial succinate dehydrogenase, which may itself perturb the cells. Furthermore, the test itself is not reversible and, because it is an endpoint reading, each time point reading requires a separate assay. It has to be pointed out that toxicity and proliferation assays are used separately and independently of each other as well as in combination. The combination of both assays allows a thorough investigation of a compounds effect on a particular cell line. There is a considerable need for assays that are homogeneous and reversible. The Oxygen Biosensor System (OBSTM ; BD Biosciences, Bedford, MA) can be seen as a prototype representative. Unlike dye conversion methods, there is no need to add any reagent in order to generate a signal. The OBS is a microplate-based assay platform that enables kinetic monitoring of dissolved oxygen. The oxygen-sensitive dye is embedded in a silicone matrix,

32 Cellular Assays and Their Application in Drug Discovery

16000

Relative fluorescence units

14000 12000 16µM Sodium azide

10000

80µM Sodium azide 400µM Sodium azide

8000

2000µM Sodium azide 10000µM Sodium azide

6000 4000 2000 0

3

30

50

73

102

126

Time after drug addition [h] Fig. 9 Oxygen consumption of HeLa cells after exposure to sodium azide. Cells (1 × 105 ), growing on beads, were seeded in individual wells of an oxygen biosensor plate and treated with sodium azide at the concentrations indicated. Cells were cultivated and ﬂuorescence was measured at the times indicated. Low micromolar concentrations of sodium azide did not affect

the growth of HeLa cells, as judged from the increased oxygen consumption during cells growth. On the other hand, millimolar concentrations of sodium azide are toxic and the relative ﬂuorescence is maintained at a constant (background) level. The decrease of oxygen consumption at 126 h after drug addition indicates that the cells became conﬂuent and stopped proliferating.

which is permanently attached to the bottom of each well of a microplate. The gas-permeable matrix allows oxygen in the vicinity of the dye to be in equilibrium with the oxygen in the liquid medium. Oxygen quenches the ability of the dye to ﬂuoresce in a predictable, concentration-dependent manner. Hence, as oxygen is depleted from a well (e.g. as cells grow and consume oxygen in the surrounding medium), the concentration of oxygen in the matrix decreases and the ﬂuorescence increases. The amount of ﬂuorescence correlates directly to oxygen consumption in the well. This, in turn, relates to the number or relative viability of cells in this particular well. Any reaction that can be linked to oxygen consumption or generation can potentially be measured by using this system (Fig. 9). The signal is dynamic and is read in real-time. Samples may be read as frequently and as many times as desired. The signal is completely reversible, and will rise as the cells proliferate and then fall as they die, giving a more complete picture of proliferation and toxicological kinetics. 5.2 Apoptosis

Apoptosis, or programmed cell death, is one of the most complex cellular processes. Early and late events occurring in cells undergoing apoptosis comprise activation

5 Phenotypic Assays 33

of different caspases, changes in the cytoskeleton, changes of the mitochondrial membrane potential and mitochondrial mass [133], nuclear condensation or fragmentation [134], ﬂipping of phosphatidylserine to the outer membrane layer and blebbing of the plasma membrane. Changes in nuclear morphology, actin reorganization, and mitochondrial potential and mass can be detected and quantiﬁed by a multiparameter apoptosis assay [135]. Increase or shrinkage of the nuclear area and chromatin distribution within the nucleus is analyzed by staining cells with a ﬂuorescent DNA dye, such as DAPI or Hoechst 33342. Bundling of intracellular F-actin ﬁlaments and increase of the actin content in apoptotic cells [136, 137] is detected by the use of ﬂuorescent phalloidin. This phallotoxin, isolated from the mushroom Amanita phalloides, binds highly speciﬁcally to F-actin and can be labeled with any ﬂuorescent dye. However, increase in F-actin content is not observed in all cases of apoptotic cells, depending on the cell type or inducer [138, 139]. The mitochondrionR Red, which accumulates in active mitochondria speciﬁc cationic dye Mitotracker is used to measure mitochondrial membrane potential and mass. Apoptotic cells can exhibit loss of mitochondrial membrane potential or increase of total mitochondrial mass [140]. In both cases, the quantity of mitochondrial stain is a measure for the degree of apoptosis. In the case of the multiparameter apoptosis application the ArrayScan II algorithm (see Section 4.2) not only quantiﬁes the label, but also takes into account the distribution of ﬂuorescent dyes within cellular compartments, and calculates nuclear fragmentation and mitochondrial mass increase. This algorithm has been successfully applied in detection of compound-induced apoptosis [135] and can be used for high-throughput compound screening. 5.3 Differentiation

Human stem cells of various types have the ability to differentiate into a number of mature cell types [141], and such stem cells are currently used in bone marrow transplants and in drug research. New drugs such as erythropoietin and granulocyte colony-stimulating factor have been developed that act directly on such cells, and are essential to the success of bone marrow transplantation [142]. The expansion of bone marrow progenitors is extremely sensitive to a wide range of chemotherapeutics and other toxic agents. In some cases, the toxic side-effects of drugs are speciﬁc to particular hematopoietic lineages [143]. In the past, use of hematopoietic progenitors for high-throughput drug discovery and toxicology screening was limited by tissue availability. Given the current availability of high-quality primary human hematopoietic progenitors, the current bottleneck in the high-throughput use of these cells is the assay of their differentiation phenotypes. The most commonly used in vitro assay in order to determine the differentiation response of hematopoietic progenitor cells is the colony assay [144]. The colony assay uses selected progenitor populations from human bone marrow, umbilical cord blood or mobilized peripheral blood in clonogenic assays in a semi-solid medium. Colonies of differentiating blood cells appear after a period of 14 days and

34 Cellular Assays and Their Application in Drug Discovery

are counted manually. Different types of colonies are identiﬁed by their morphology (e.g. myeloid and erythroid) or cytochemical staining techniques (megakaryocytes). Although the parameters of these assays have been optimized to obtain quantitative data, the extremely low throughput and subjective nature of this assay cause some problems when it is used for chemical compound library screening, early drug discovery and high-throughput toxicity screening. Differentiative responses can also be assayed by the transfection of a target cell with a receptor gene tied to the expression of a differentiation-speciﬁc marker [145]. It requires, however, the transfection of a cell containing the many intact signal transduction cascades characteristic of hematopoietic progenitors. Another approach was described by Warren et al. [146]. The authors described a cell-based immunoassay in which the cells were transferred to a ﬁxative-treated assay plate, and ﬁxed in place prior to incubations with primary and secondary antibodies. Although this assay has been used successfully, transfer of cells from the culture plate to the assay plate is time-consuming and not amenable to HTS. Recently [147], this assay design was revised and improved in order to reduce the amount of manipulations required. On termination of the culture period, cell culture supernatant is removed via vacuum ﬁltration and a labeled speciﬁc antibody is added. In order to obtain the sensitivity required, the monoclonal antibodies were conjugated to a DELFIA europium chelate [148]. In addition to its sensitivity, the relatively long emission time of the DELFIA europium chelate allowed the use of time-resolved ﬂuorescence (TRF) to avoid the endogenous background ﬂuorescence that is often a signiﬁcant problem in biological assays. Removal of the cell culture supernatant, unbound antibodies and subsequent wash steps are performed by using a 96-well ﬁlter plate. Because there is little carryover from one wash cycle to the next, removal of unbound antibody is very efﬁcient. Repeated washes without cell loss are possible. For signal detection, an enhancement solution was added and europium ﬂuorescence (excitation at 340 nm and emission at 615 nm) was measured over a 400-µs time period after an initial delay of 400 µs using a suitable spectroﬂuorometer.

5.4 Monitoring Cell Metabolism

Direct determination of metabolites at the cellular or in vivo level provides a sensitive system for assessing drug action because metabolites represent end products of cellular regulatory processes. In addition, intracellular metabolites are generally far less abundant than proteins. In a cellular environment, signal transduction, initiation of gene expression, protein synthesis and metabolic responses to a stress factor must be essentially sequential. The ﬁelds of metabolomics – which investigates metabolic regulation and ﬂuxes in individual cells or cell types – and metabonomics – the determination of systemic biochemical proﬁles and regulation of function in whole organisms by analyzing bioﬂuids and tissues – have therefore been developed recently [15, 149].

5 Phenotypic Assays 35

Studying the effects of drugs on cells or whole organisms by metabonomics relies on multiparametric measurements of alterations in metabolism over time in response to a stress factor. Samples are mapped according to their biochemical composition [150]. Metabolites proved particularly useful in the characterization of mutant phenotypes as shown recently in plants [151] and in the characterization of so-called “silent mutations” in yeast [6]. Many genes are “silent”, which means that if mutated they do not lead to an easily visible phenotypic change, such as alterations in growth rate. The growth rate may not be changed because the concentrations of different intracellular metabolites are adapted in a way that compensates for the effect of the mutation. Accordingly, metabolome analysis can reveal a phenotype at the level of metabolites in many mutants previously scored as silent. In addition, quantifying the relative changes of metabolite concentrations caused by a mutation makes it possible to identify the site of action of the altered gene product as demonstrated by Raamsdonk et al. [6]. By using nuclear magnetic resonance or mass spectrometry (MS)/liquid chromatography/MS applications, a reasonable sample throughput is possible. This will allow the use of metabolome and metabonome analysis in drug discovery as a sensitive measure of drug action [15]. An emerging approach, which allows the analysis of the metabolic state in intact cells, is the functional online analysis of living cells in physiologically controlled environments for extended periods of time (“biochips”). Efforts are being directed towards the parallel development, adaptation and integration of different microelectronic sensors into miniaturized biochips for a multiparametric cellular monitoring (“Cell Monitoring System” [22]). Some possible readouts and the corresponding cellular activities are summarized in Tab. 4. Monitoring of cellular oxygen exchange [152] can be employed for the assessment of mitochondrial activities. Other sinks or sources of oxygen do not contribute signiﬁcantly in most cell types. Mitochondria, apart from supplying the cell with energy, are suggested to be involved in cell ageing accelerated by reactive oxygen species, in apoptosis and in the toxic mechanisms of numerous drugs. Monitoring mitochondrial activity by lipophilic dyes such as rhodamine 123 suffers from the

Table 4 Readouts for multiparametric cell monitoring systems

Readout

Corresponding cellular activity

Changes in oxygen level Acidiﬁcation of the medium Changes in impedance

mitochondrial activity (energy metabolism; cell ageing; apoptosis) global indicator of metabolic activity; fast changes: cellular activation events; slow changes: cell growth or death cell adhesion: matrix attachment, tumor metastasis; cell outgrowth, e.g. neurite differentiation inﬂux and efﬂuxofCa2+ , Na+ , K+ e.g. glucose uptake; lactate excretion e.g. induction of reporter gene expression coupled to the generation of bioluminescence

Ion ﬂow Metabolites Light emission

36 Cellular Assays and Their Application in Drug Discovery

risk of potential artifacts, and therefore, nontoxic and noninvasive techniques are particularly valuable. The OBS has been described above as one of the ﬁrst noninvasive techniques to monitor oxygen consumption. One limitation of the OBS, however, is the requirement for a large number of cells. The system needs at least 50,000 cells per well in order to produce viable results (less cells do not consume enough oxygen in order to (over)compensate the diffusion of oxygen back from the outside environment to the oxygen biosensor). In comparison, microsensor-based technologies are so small in scale that, under ideal circumstances, signals can be obtained from a single cell. Microsensor-based pH recording [153] is predominantly used for determinations of extracellular acidiﬁcation rates. Several metabolic pathways contribute to extracellular acidiﬁcation and, thus, can be regarded as a global indicator of metabolic activity. In this sense, fast changes (within minutes) tend to reﬂect cellular activation events (e.g. receptor-mediated signaling), while slow changes (several hours) are usually attributable to cell growth or cell death [154], respectively. Monitoring of cell adhesion (e.g. tumor metastasis) or outgrowth (e.g. for neurites) can be accomplished by growing the cells directly on pairs of interdigitated electrodes. The cellular impedance signal results from insulation by the cell membranes. If cells are placed on the electrodes, they block the current ﬂow in a passive way and the impedance increases. Therefore, the impedance of the system gives insight into the adhesive behavior of the cells. This methodology is based on on-line data acquisition using multiparametric microsensor devices. A ﬂuid system for the supply of culture media/drug solutions and for the realization of metabolic measurements is connected to the chip. The precise maintenance of in vitro microenvironmental conditions is also necessary for the cellular specimen. The cellular parameters currently amenable by sensor chip readings are metabolic activity (rates of extracellular acidiﬁcation and cellular respiration), morphologic properties and some ion ﬂuxes. Small-scale sensor chip areas require very small amounts of cellular specimen. This may allow the utilization use of primary cells. Methods have been developed which allow testing of dozens of compounds, but 96-well formats might become available soon [22]. 5.5 Other Phenotypic Assays

There is an increased need for real-time, label-free observations of proteins and cells under their natural (unperturbed) conditions due to some limitations associated with the currently existing technologies. A major drawback of today’s technologies is the need to label and overexpress the protein of interest. Due to its overexpression and modiﬁcation, the protein’s biochemical activity and localization may not reﬂect the natural (unperturbed) conditions, and therefore the results have to be interpreted with caution. One technology that addresses those issues uses radiofrequency waves [21] to probe the physiological state of a protein in the context of the whole cell. MCS uses microwaves, which correspond to the natural frequency of

References

protein dynamics, allowing the measurement of electrodynamic properties of proteins and cells. MCS does not use any tags or markers and may be performed in any environment, from simple aqueous solutions to very complex mixtures (i.e. cells), thereby allowing a direct approach to determine changes in cellular phenotypes (i.e. differentiation). Isolated proteins in physiological buffers exposed to a range of microwave frequencies, demonstrate two properties: the spectral response (the proteins “signature”) and the biophysical proﬁle. The ﬁrst of these determines the raw response of each protein over a range of frequencies. The biophysical proﬁle describes the way in which a protein interacts with the environment – molecular volume, complexation of water, interaction with other compounds/proteins – as well as how it compares to other proteins belonging to similar classes. By scanning a range of frequencies and evaluating different environments (e.g. buffer conditions, presence of cofactors), this analysis deﬁnes a multidimensional signature to each protein even in complex (cellular) environments. This would allow the development of cell-based assays where the biophysical proﬁle of target proteins can be analyzed in an environment that closely mimics the situation in primary disease tissues.

Acknowledgments

We thank Dr. Guy Marais for carefully reading and reviewing the manuscript.

References 1 TAN D. S. 2002. Sweet surrender to chemical genetics. Nat Biotechnol 20, 561–3. 2 STEPHANOPOULOS G., J. KELLEHER. 2001. Biochemistry. How to make a superior cell. Science 292, 2024–5. 3 HANAHAN D., R. A. WEINBERG. 2000. The hallmarks of cancer. Cell 100, 57–70. 4 SOMOGYI R., L. D. GRELLER. 2001. The dynamics of molecular networks: applications to therapeutic discovery. Drug Discov Today 6, 1267–77. 5 GLASSBROOK N., C. BEECHER, J. RYALS. 2000. Metabolic proﬁling on the right path. Nat Biotechnol 18, 1142–3. 6 RAAMSDONK L. M., B. TEUSINK, D. BROADHURST, N. ZHANG, A. HAYES, M. C. WALSH, J. A. BERDEN, K. M. BRINDLE, D. B. KELL, J. J. ROWLAND, H. V. WESTERHOFF, K. van DAM, S. G. OLIVER. 2000. A functional genomics strategy

7

8

9

10

11

12

that uses metabolome data to reveal the phenotype of silent mutations. Nat Biotechnol 19, 45–50. GIFFORD D. K. 2002. Blazing pathways through genetic mountains. Science 293, 2049–51. KARP P. D. 2001. Pathway databases: a case study in computational symbolic theories. Science 293, 2040–4. WHITE M. A. 1996. The yeast two-hybrid system: forward and reverse. Proc Natl Acad Sci USA 93, 10001–3. ITO T., T. CHIBA, M. YOSHIDA. 2001. Exploring the protein interactome using comprehensive two-hybrid projects. Trends Biotechnol 19, S23–7. TUCKER C. L., S. FIELDS. 2001. A yeast sensor of ligand binding. Nat Biotechnol 19, 1042–6. OSBORNE M. A., S. DALTON, J. P. KOCHAN. 1995. The yeast tribrid system – genetic

37

38 Cellular Assays and Their Application in Drug Discovery

13

14

15

16

17

18

19

20

21

22

23

detection of trans-phosphorylated ITAM–SH2-interactions. Biotechnology 13, 1474–8. STRAUB B., E. MEYER, P. FROMHERZ. 2001. Recombinant maxi-K channels on transistor, a prototype of ionoelectronic interfacing. Nat Biotechnol 19, 121–124. CELIS J. E., M. KRUHOFFER, I. GROMOVA, C. FREDERIKSEN, M. OSTERGRAAD, T. THYKJAER, P. GROMOV, J. VU, H. PALSDOTTIR , N. MAGNUSSON, T. F. ` ORTNTOFT. 2000. Gene expression proﬁling: monitoring transcription and translation products using DNA micro-arrays and proteomics. FEBS Lett 480, 2–16. NICHOLSON J. K.g, J. CONNELLY, C. LINDON, E. HOLMES. 2002. Metabonomics: a platform for studying drug toxicity and gene function. Nat Rev Drug Discov 1, 153–61. Broach JR, J. THORNER. 1996. High throughput screening for drug discovery. Nature 384, 14–6. GOETZ A. S., J. L. ANDREWS, T. R. LITTLETON, D. M. IGNAR. 2000. Development of a facile method for high throughput screening with reporter gene assays. J Biomol Screen 5, 377–84. REMINGTON S. J. 2002. Negotiating the speed bumps to ﬂuorescence. Nat Biotechnol 20, 28–9. SCHAUFELE F. 2001. Shedding light on molecular timing. Nat Biotechnol 19, 21–2. MITCHELL P. 2001. Turning the spotlight on cellular imaging. Nat Biotechnol 19, 1013–7. HEFTI J. P., A. KUMAR. 1999. Sensitive detection method of dielectric dispersions in aqueous based, surface bound macromolecular structures using microwave spectroscopy. Appl Phys Lett 75, 1802–4. BAUMANN W., M. LEHMANN, M. BITZENHOFER, A. SCHWINDE, M. BIRSCHWEIN, R. EHRET, B. WOLF. 1999. Microelectronic sensor system for microphysiological application on living cells. Sensors and Acuators B 55, 77–89. SCHLESSINGER J. 1993. Cellular signaling by receptor tyrosine kinases. Harvey Lect 89, 105–23.

24 SULLIVAN E., E. M. TUCKER, I. L. DALE. 1999. Measurement of [Ca2+ ] using the Fluorometric Imaging Plate Reader (FLIPR). Methods Mol Biol 114, 125–33. 25 MILLIGAN G., S. REES. 1999. Chimaeric G alpha proteins: their potential use in drug discovery. Trends Pharmacol Sci 20, 118–24. 26 NEER E. J. 1995. Heterotrimeric G proteins: organizers of transmembrane signals. Cell 80, 249–57. 27 ZABIN I. 1982. A model of protein interaction. Mol Cell Biochem 49, 87–96. 28 DEAN M., Y. HAMON, G. CHIMINI. 2001. The human ATP-binding cassette (ABC) transporter super-family. J Lipid Res 42, 1007–17. 29 BRINKMANN U., M. EICHELBAUM. 2001. Polymorphisms in the ABC drug transporter gene MDR1. Pharmacogenomics 1, 59–64. 30 HWANG L. Y., P. T. LIEU, P. A. PETERSON, Y. YANG. 2001. Functional regulation of immunoproteasomes and transporter associated with antigen processing. Immunol Res 24, 245–72. 31 KO Y. H., P. L. PEDERSEN. 2001. Cystic ﬁbrosis: a brief look at some highlights of a decade of research focused on elucidating and correcting the molecular basis of the disease. J Bioenerg Biomembr 33, 513–21. 32 OUELLETTE M., D. LEGARE, B. PAPADOPOULOU. 2001. Multidrug resistance and ABC transporters in parasitic protozoa. J Mol Microbiol Biotechnol 3, 201–6. 33 SCHMIDT C., M. MAYER, H. VOGEL. 2000. A chip-based biosensor for the functional analysis of single ion channels. Angew Chem Int 39, 3137–40. 34 XU J. Z., X. WANG, B. ENSIGN, M. LI, L. WU, A. GUIA, J. XU. 2001. Ion-channel assay technologies: quo vadis? Drug Discov Today 6, 1278–87. 35 BAXTER D. F., M. KIRK, A. F. GARCIA, A. RAIMONDI, M. H. HOLMQVIST, K. K. FLINT, D. BOJANIC, P. S. DISTEFANO, R. CURTIS, Y. XIE. 2002. A novel membrane potential-sensitive ﬂuorescent dye improves cell-based assays for ion channels. J Biomol Screen 7, 79–85.

References 36 WHITEAKER K. L., S. M. GOPALAKRISHNAN, D. GROEBE, C. C. SHIEH, U. WARRIOR, D. J. BURNS, M. J. COGHLAN, V. E. SCOTT, M. GOPALAKRISHNAN. 2001. Validation of FLIPR membrane potential dye for high throughput screening of potassium channel modulators. J Biomol Screen 6, 305–12. 37 SARVER J. G., W. A. KLIS, J. P. BYERS, P. W. ERHARDT. 2002. Microplate screening of the differential effects of test agents on Hoechst 33342, rhodamine 123, and rhodamine 6G accumulation in breast cancer cells that overexpress P-glycoprotein. J Biomol Screen 7, 29–34. 38 EPSTEIN C. B., R. A. BUTOW. 2000. Microarray technology–enhanced versatility, persistent challenge. Curr Opin Biotechnol 11, 36–41. 39 HUGHES T. R., M. J. MARTON, A. R. JONES, C. J. ROBERTS, R. STOUGHTON, C. D. ARMOUR, H. A. BENNETT, E. COFFEY, H. DAI, Y. D. HE, M. J. KIDD, A. M. KING, M. R. MEYER, D. SLADE, P. Y. LUM, S. B. STEPANIANTS, D. D. SHOEMAKER, D. GACHOTTE, K. CHAKRABURTTY, J. SIMON, M. BARD, S. H. FRIEND. 2000. Functional discovery via a compendium of expression proﬁles. Cell 102, 109–26. 40 WEINBERGER S. R., E. A. DALMASSO, E. T. FUNG. 2002. Current achievements using ProteinChip Array technology. Curr Opin Chem Biol 6, 86–91. 41 GOETZ A. S., J. LIACOS, J. YINGLING, D. M. IGNAR. 1999. A combination assay for simultaneous assessment of multiple signaling pathways. J Pharmacol Toxicol Methods 42, 225–35. 42 FREEMAN T. 2000. High throughput gene expression screening: its emerging role in drug discovery. Med Res Rev 20, 197–202. 43 PANDEY A., M. MANN. 2000. Proteomics to study genes and genomes. Nature 405, 837–46. 44 BRIVANLOU A. H., J. E. DARNELL, Jr. 2002. Signal transduction and the control of gene expression. Science 295, 813–8. 45 NAYLOR L. H. 1999. Reporter gene technology: the future looks bright. Biochem Pharmacol 58, 749–57.

46 TERSTAPPEN G. C., A. GIACOMETTI, E. BALLINI, L. ALDEGHERI. 2000. Development of a functional reporter gene HTS assay for the identiﬁcation of mGluR7 modulators. J Biomol Screen 5, 255–62. 47 XING H., H. C. TRAN, T. E. KNAPP, P. A. NEGULESCU, B. A. POLLOK. 2000. A ﬂuorescent reporter assay for the detection of ligands acting through Gi protein-coupled receptors. J Recept Signal Transduct Res 20, 189–210. 48 DUROCHER Y., PERRET S., THIBAUDEAU E., GAUMOND M. H., KAMEN A., STOCCO R., ABRAMOVITZ M. 2000. A reporter gene assay for high-throughput screening of G-protein-coupled receptors stably or transiently expressed in HEK293 EBNA cells grown in suspension culture. Anal Biochem 284, 316–26. 49 DAVIS G. F., T. R. DOWNS, J. A. FARMER, C. R. PIERSON, J. T. ROESGEN, E. J. CABRERA, S. L. NELSON. 2002. Comparison of high throughput screening technologies for luminescence cell-based reporter screens. J Biomol Screen 7, 67–77. 50 WILSON V. S., K. BOBSEINE, C. R. LAMBRIGHT, L. E. GRAY, Jr. 2002. A novel cell line, mda-kb2, that stably expresses an androgen- and glucocorticoid-responsive reporter for the detection of hormone receptor agonists and antagonists. Toxicol Sci 66, 69–81. 51 ZHANG J. H., T. D. CHUNG, K. R. OLDENBURG. 1999. A simple statistical parameter for use in evaluation and validation of high throughput screening assays. J Biomol Screen 4, 67–73. 52 REES S., D. P. MARTIN, S. V. SCOTT, S. H. BROWN, N. FRASER, C. O’SHAUGHNESSY, I. J. BERESFORD. 2001. Development of a homogeneous MAP kinase reporter gene screen for the identiﬁcation of agonists and antagonists at the CXCR1 chemokine receptor. J Biomol Screen 6, 19–27. 53 POMEROY S. L., P. TAMAYO, M. GAASENBEEK, L. M. STURLA, M. ANGELO, M. E. MCLAUGHLIN, J. Y. KIM, L. C. GOUMNEROVA, P. M. BLACK, C. LAU, J. C. ALLEN, D. ZAGZAG, J. M. OLSON, T.

39

40 Cellular Assays and Their Application in Drug Discovery

54

55

56

57

58

59

60

CURRAN, C. WETMORE, J. A. BIEGEL, T. POGGIO, S. MUKHERJEE, R. RIFKIN, A. CALIFANO, G. STOLOVITZKY, D. N. LOUIS, J. P. MESIROV, E. S. LANDER, T. R. GOLUB. 2002. Prediction of central nervous system embryonal tumour outcome based on gene expression. Nature 415, 436–42. IDEKER T., V. THORSSON, J. A. RANISH, R. CHRISTMAS, J. BUHLER, J. K. ENG, R. BUMGARNER, D. R. GOODLETT, R. AEBERSOLD, L. HOOD. 2001. Integrated genomic and proteomic analyses of a systematically perturbed metabolic network. Science 292, 929–34. ULRICH R. G., S. H. FRIEND. 2002. Toxicogenomics and drug discovery: will new technologies help us produce better drugs? Nat Rev Drug Discov 1, 84–5. NUWAYSIR E. F., M. BITTNER, J. TRENT, J. C. BARRETT, C. A. AFSHARI. 1999. Microarrays and toxicology: the advent of toxicogenomics. Mol Carcinog 24, 153–9. ROTHBERG B. E.. 2001. The use of animal models in expression pharmacogenomic analyses. Pharma-cogenomics 1, 48–58. SHIPP M. A., K. N. ROSS, P. TAMAYO, A. P. WENG, J. L. KUTOK, R. C. AGUIAR, M. GAASENBEEK, M. ANGELO, M. REICH, G. S. PINKUS, T. S. RAY, M. A. KOVAL, K. W. LAST, A. NORTON, T. A. LISTER, J. MESIROV, D. S. NEUBERG, E. S. LANDER, J. C. ASTER, T. R. GOLUB. 2002. Diffuse large B-cell lymphoma outcome prediction by gene-expression proﬁling and supervised machine learning. Nat Med 8, 68–74. VAN’T VEER L. J., H. DAI, M. J. van de VIJVER, Y. D. HE, A. A. HART, M. MAO, H. L. PETERSE, K. K. van DER, M. J. MARTON, A. T. WITTEVEEN, G. J. SCHREIBER, R. M. KERKHOVEN, C. ROBERTS, P. S. LINSLEY, R. BERNARDS, S. H. FRIEND. 2002. Gene expression proﬁling predicts clinical outcome of breast cancer. Nature 415, 530–6. THOMAS R. S., D. R. RANK, S. G. PENN, G. M. ZASTROW, K. R. HAYES, K. PANDE, E. GLOVER, T. SILANDER, M. W. CRAVEN, J. K. REDDY, S. B. JOVANOVICH, C. A. BRADFIELD. 2001. Identiﬁcation of toxicologically predictive gene sets using

61

62

63

64

65

66

67

68

69

cDNA microarrays. Mol Pharmacol 60, 1189–94. BARTOSIEWICZ M. J., D. JENKINS, S. PENN, J. EMERY, A. BUCKPITT. 2001. Unique gene expression patterns in liver and kidney associated with exposure to chemical toxicants. J Pharmacol Exp Ther 297, 895–905. BURCZYNSKI M. E., M. MCMILLIAN, J. CIERVO, L. LI, J. B. PARKER, R. T. DUNN, S. HICKEN, S. FARR, M. D. JOHNSON. 2000. Toxicogenomics-based discrimination of toxic mechanism in HepG2 human hepatoma cells. Toxicol Sci 58, 399–415. WARING J. F., R. CIURLIONIS, R. A. JOLLY, M. HEINDEL, R. G. ULRICH. 2001. Microarray analysis of hepatotoxins in vitro reveals a correlation between gene expression proﬁles and mechanisms of toxicity. Toxicol Lett 120, 359–68. FARR S. B., M. D. TODD. 1998. Methods and kits for eukaryotic gene proﬁling. President and Fellows of Harvard College (Cambridge, MA, Xenometrix, Inc.). US Patent 5,811,231. ALBANO C. L., W. E. BENTLEY, G. RAO. 2001. High throughput studies of gene expression using green ﬂuorescent protein–oxidative stress promoter probe constructs. J Biomol Screen 6, 421–8. ZHU M., W. E. FAHL. 2000. Development of a green ﬂuorescent protein microplate assay for the screening of chemopreventive agents. Anal Biochem 287, 210–7. ZHAN Q., F. CARRIER, A. J. FORNACE, Jr. 1993. Induction of cellular p53 activity by DNA-damaging agents and growth arrest. Mol Cell Biol 13, 4242–50. DELESCLUSE C., N. LEDIRAC, R. LI, M. P. PIECHOCKI, R. N. HINES, X. GIDROL, R. RAHMANI. 2001. Induction of cytochrome P450 1A1 gene expression, oxidative stress, and genotoxicity by carbaryl and thiabendazole in transfected human HepG2 and lymphoblastoid cells. Biochem Pharmacol 61, 399–407. WATKINS R. E., G. B. WISELY, L. B. MOORE, J. L. COLLINS, M. H. LAMBERT, S. P. WILLIAMS, T. M. WILLSON, S. A. KLIEWER, M. R. REDINBO. 2001. The

References

70

71

72

73

74

75

76

77

78

human nuclear xenobiotic receptor PXR: structural determinants of directed promiscuity. Science 292, 2329–33. BERTELSEN A. H., V. E. VELCULENSCU. 1998. High-throughput gene expression analysis using SAGE. Drug Discov Today 3, 152–9. SHIMKETS R. A., D. G. LOWE, J. T. TAI, P. SEHL, H. JIN, R. YANG, P. F. PREDKI, B. E. ROTHBERG, M. T. MURTHA, M. E. ROTH, S. G. SHENOY, A. WINDEMUTH, J. W. SIMPSON, J. F. SIMONS, M. P. DALEY, S. A. GOLD, M. P. MCKENNA, K. HILLAN, G. T. WENT, J. M. ROTHBERG. 1999. Gene expression analysis by transcript proﬁling coupled to a gene database query. Nat Biotechnol 17, 798–803. SUTCLIFFE J. G., P. E. FOYE, M. G. ERLANDER, B. S. HILBUSH, L. J. BODZIN, J. T. DURHAM, K. W. HASEL. 2000. TOGA: an automated parsing technology for analyzing expression of nearly all genes. Proc Natl Acad Sci USA 97, 1976–81. HEID C. A., J. STEVENS, K. J. LIVAK, P. M. WILLIAMS. 1996. Real time quantitative PCR. Genome Res 6, 986–94. TODD M. D., X. LIN, L. F. STANKOWSKI, Jr., M. DESAI, G. H. WOLFGANG. 1999. Toxicity screening of a combinatorial library: correlation of cytotoxicity and gene induction to compound structure. J Biomol Screen 4, 259–68. WARRIOR U., Y. FAN, C. A. DAVID, J. A. WILKINS, E. M. MCKEEGAN, J. L. KOFRON, D. J. BURNS. 2000. Application of QuantiGene nucleic acid quantiﬁcation technology for high throughput screening. J Biomol Screen 5, 343–52. CONRAD M. J.. 1999. Measurement of speciﬁc gene transcripts in 96-well plate assays. Genet Engng News 19. GRAHAM D. L., N. BEVAN, P. N. LOWE, M. PALMER, S. REES. 2001. Application of beta-galactosidase enzyme complementation technology as a high throughput screening format for antagonists of the epidermal growth factor receptor. J Biomol Screen 6, 401–11. REMY I., S. W. MICHNICK. 2001. Visualization of biochemical networks in living cells. Proc Natl Acad Sci USA 98, 7678–83.

79 GRIFFIN B. A., S. R. ADAMS, R. Y. TSIEN. 1998. Speciﬁc covalent labeling of recombinant protein molecules inside live cells. Science 281, 269–72. 80 KAIN S. R.. 1999. Green ﬂuorescent protein (GFP): applications in cell-based assays for drug discovery. Drug Discov Today 4, 304–12. 81 MATZ M. V., A. F. FRADKOV, Y. A. LABAS, A. P. SAVITSKY, A. G. ZARAISKY, M. L. MARKELOV, S. A. LUKYANOV. 1999. Fluorescent proteins from nonbioluminescent Anthozoa species. Nat Biotechnol 17, 969–73. 82 EDWARDS B. S., F. W. KUCKUCK, E. R. PROSSNITZ, J. T. RANSOM, L. A. SKLAR. 2001. HTPS ﬂow cytometry: a novel platform for automated high throughput drug discovery and characterization. J Biomol Screen 6, 83–90. 83 TAYLOR D. L., E. S. WOO, K. A. GIULIANO. 2001. Real-time molecular and cellular analysis: the new frontier of drug discovery. Curr Opin Biotechnol 12, 75–81. 84 BRUNET A., D. ROUX, P. LENORMAND, S. DOWD, S. KEYSE, J. POUYSSEGUR. 1999. Nuclear translocation of p42/p44 mitogen-activated protein kinase is required for growth factor-induced gene expression and cell cycle entry. EMBO J 18, 664–74. 85 DALAL S. N., C. M. SCHWEITZER, J. GAN, J-A DECAPRIO. 1999. Cytoplasmic localization of human cdc25C during interphase requires an intact 14-3-3 binding site. Mol Cell Biol 19, 4465–79. 86 LIU F., C. ROTHBLUM-OVIATT, C. E. RYAN, H. PIWNICA-WORMS. 1999. Overproduction of human Myt1 kinase induces a G2 cell cycle delay by interfering with the intracellular trafﬁcking of Cdc2–cyclin B1 complexes. Mol Cell Biol 19, 5113–23. 87 VOGT A., T. ADACHI, A. P. DUCRUET, J. CHESEBROUGH, K. NEMOTO, B. I. CARR, J. S. LAZO. 2001. Spatial analysis of key signaling proteins by high-content solid-phase cytometry in Hep3B cells treated with an inhibitor of Cdc25 dual-speciﬁcity phosphatases. J Biol Chem 276, 20544–50.

41

42 Cellular Assays and Their Application in Drug Discovery

88 CHADEE D. N., M. J. HENDZEL, C. P. TYLIPSKI, C. D. ALLIS, D. P. BAZETT-JONES, J. A. WRIGHT, J. R. DAVIE. 1999. Increased Ser-10 phosphorylation of histone H3 in mitogen-stimulated and oncogenetransformed mouse ﬁbroblasts. J Biol Chem 274, 24914–20. 89 BOGGS B. A., C. D. ALLIS, A. C. CHINAULT. 2000. Immunoﬂuorescent studies of human chromosomes with antibodies against phosphorylated H1 histone. Chromosoma 108, 485–90. 90 PEREZ O. D., G. P. NOLAN. 2002. Simultaneous measurement of multiple active kinase states using polychromatic ﬂow cytometry. Nat Biotechnol 20, 155–62. 91 MAYER T. U., T. M. KAPOOR, S. J. HAGGARTY, R. W. KING, S. L. SCHREIBER, T. J. MITCHISON. 1999. Small molecule inhibitor of mitotic spindle bipolarity identiﬁed in a phenotype-based screen. Science 286, 971–4. 92 BOGGS B. A., B. CONNORS, R. E. SOBEL, A. C. CHINAULT, C. D. ALLIS. 1996. Reduced levels of histone H3 acetylation on the inactive X chromosome in human females. Chromosoma 105, 303–9. 93 HENDZEL M. J., M. J. KRUHLAK, D. P. BAZETT-JONES. 1998. Organization of highly acetylated chromatin around sites of heterogeneous nuclear RNA accumulation. Mol Biol Cell 9, 2491–507. 94 YONEDA Y. 2000. Nucleocytoplasmic protein trafﬁc and its signiﬁcance to cell function. Genes Cells 5, 777–87. 95 HOWELL J. L., R. TRUANT. 2002. Live-cell nucleocytoplasmic protein shuttle assay utilizing laser confocal microscopy and FRAP. Biotechniques 32, 80–7. 96 DING G. J., P. A. FISCHER, R. C. BOLTZ, J. A. SCHMIDT, J. J. COLAIANNE, A. GOUGH, R. A. RUBIN, D. K. MILLER. 1998. Characterization and quantitation of NF-kappaB nuclear translocation induced by interleukin-1 and tumor necrosis factor-alpha. Development and use of a high capacity ﬂuorescence cytometric system. J Biol Chem 273, 28897–905. 97 De WIT R., C. M. HENDRIX, J. BOONSTRA, A. J. VERKLEU, J. A. POST. 2000. Large-scale screening assay to measure

98

99

100

101

102

103

104

105

epidermal growth factor internalization. J Biomol Screen 5, 133–40. PRASHER D. C., V. K. ECKENRODE, W. W. WARD, F. G. PRENDERGAST, M. J. CORMIER. 1992. Primary structure of the Aequorea victoria green-ﬂuorescent protein. Gene 111, 229–33. CODY C. W., D. C. PRASHER, W. M. WESTLER, F. G. PRENDERGAST, W. W. WARD. 1993. Chemical structure of the hexapeptide chromophore of the Aequorea green-ﬂuorescent protein. Biochemistry 32, 1212–8. BARAK L. S., S. S. FERGUSON, J. ZHANG, M. G. CARON. 1997. A beta-arrestin/ green ﬂuorescent protein biosensor for detecting G protein-coupled receptor activation. J Biol Chem 272, 27497–500. OAKLEY R. H., S. A. LAPORTE, J. A. HOLT, M. G. CARON, L. S. BARAK. 2000. Differential afﬁnities of visual arrestin, beta arrestin1, and beta arrestin2 for G protein-coupled receptors delineate two major classes of receptors. J Biol Chem 275, 17201–10. GROARKE D. A., S. WILSON, C. KRASEL, G. MILLIGAN. 1999. Visualization of agonist-induced association and trafﬁcking of green ﬂuorescent protein-tagged forms of both beta-arrestin-1 and the thyrotropin-releasing hormone receptor-1. J Biol Chem 274, 23263–9. GROARKE D. A., T. DRMOTA, D. S. BAHIA, N. A. EVANS, S. WILSON, G. MILLIGAN. 2001. Analysis of the C-terminal tail of the rat thyrotropin-releasing hormone receptor-1 in interactions and cointernalization with beta-arrestin 1-green ﬂuorescent protein. Mol Pharmacol 59, 375–85. GHOSH R. N., Y. T. CHEN, R. DEBIASI, R. L. DEBIASIO, B. R. CONWAY, L. K. MINOR, K. T. DEMAREST. 2000. Cell-based, high-content screen for receptor internalization, recycling and intracellular trafﬁcking. Biotechniques 29, 170–5. CONWAY B. R., L. K. MINOR, J. Z. XU, J. W. GUNNET, R. DEBIASIO, M. R. D’ANDREA, R. RUBIN, R. DEBIASIO, K. GIULIANO, L. DEBIASIO, K. T. DEMAREST.

References

106

107

108

109

110

111

112

113

114

1999. Quantiﬁcation of G-protein coupled receptor internalization using G-protein coupled receptor-green ﬂuorescent protein conjugates with the ArrayScanTM high-content screening system. J Biomol Screen 4, 75–86. MERMELSTEIN P. G., K. DEISSEROTH, N. DASGUPTA, A. L. ISAKSEN, R. W. TSIEN. 2001. Calmodulin priming: nuclear translocation of a calmodulin complex and the memory of prior neuronal activity. Proc Natl Acad Sci USA 98, 15342–7. SANTAGATA S., T. J. BOGGON, C. L. BAIRD, C. A. GOMEZ, J. ZHAO, W. S. SHAN, D. G. MYSZKA, L. SHAPIRO. 2001. G-protein signaling through tubby proteins. Science 292, 2041–50. TERUEL M. N., T. MEYER. 2002. Parallel single-cell monitoring of receptor-triggered membrane translocation of a calcium-sensing protein module. Science 295, 1910–2. TORRANCE C. J., V. AGRAWAL, B. VOGELSTEIN, K. W. KINZLER. 2001. Use of isogenic human cancer cells for high-throughput screening and drug discovery. Nat Biotechnol 19, 940–5. LIPPINCOTT-SCHWARTZ J., E. SNAPP, A. KENWORTHY. 2001. Studying protein dynamics in living cells. Nat Rev Mol Cell Biol 2, 444–56. MAHAJAN N. P., K. LINDER, G. BERRY, G. W. GORDON, R. HEIM, B. HERMAN. 1998. Bcl–2 and Bax interactions in mitochondria probed with green ﬂuorescent protein and ﬂuorescence resonance energy transfer. Nat Biotechnol 16, 547–52. ZHANG J., Y. MA, S. S. TAYLOR, R. Y. TSIEN. 2001. Genetically encoded reporters of protein kinase A activity reveal impact of substrate tethering. Proc Natl Acad Sci USA 98, 14997– 5002. TING A. Y., K. H. KAIN, R. L. KLEMKE, R. Y. TSIEN. 2001. Genetically encoded ﬂuorescent reporters of protein tyrosine kinase activities in living cells. Proc Natl Acad Sci USA 98, 15003–8. HONDA A., S. R. ADAMS, C. L. SAWYER, V. LEV-RAM, R. Y. TSIEN, W. R. DOSTMANN. 2001. Spatiotemporal dynamics of guanosine 3 ,5 -cyclic monophosphate

115

116

117

118

119

120

121

122

revealed by a genetically encoded, ﬂuorescent indicator. Proc Natl Acad Sci USA 98, 2437–42. ZACCOLO M., F. De GIORGI, C. Y. CHO, L. FENG, T. KNAPP, P. A. NEGULESCU, S. S. TAYLOR, R. Y. TSIEN, T. POZZAN. 2000. A genetically encoded, ﬂuorescent indicator for cyclic AMP in living cells. Nat Cell Biol 2, 25–9. MOCHIZUKI N., S. YAMASHITA, K. KUROKAWA, Y. OHBA, T. NAGAI, A. MIYAWAKI, M. MATSUDA. 2001. Spatio-temporal images of growth-factor-induced activation of Ras and Rap1. Nature 411, 1065–8. XU Y., D. W. PISTON, C. H. JOHNSON. 1999. A bioluminescence resonance energy transfer (BRET) system: application to interacting circadian clock proteins. Proc Natl Acad Sci USA 96, 151–6. ANGERS S., A. SALAHPOUR, E. JOLY, S. HILAIRET, D. CHELSKY, M. DENNIS, M. BOUVIER. 2000. Detection of beta 2-adrenergic receptor dimerization in living cells using bioluminescence resonance energy transfer (BRET). Proc Natl Acad Sci USA 97, 3684–9. AYOUB M. A., C. COUTURIER, E. LUCAS-MEUNIER, S. ANGERS, P. FOSSIER, M. BOUVIER, R. JOCKERS. 2002. Monitoring of ligand-independent dimerization and ligand-induced conformational changes of melatonin receptors in living cells by bioluminescence resonance energy transfer. J Biol Chem 277, 21522–8. RAMSAY D., E. KELLETT, M. MCVEY, S. REES, G. MILLIGAN. 2002. Homo- and hetero-oligomeric interactions between G protein-coupled receptors in living cells monitored by two variants of bioluminescence resonance energy transfer. Biochem J 365, 429–40. ROSSI F., C. A. CHARLTON, H. M. BLAU. 1997. Monitoring protein–protein interactions in intact eukaryotic cells by beta-galactosidase complementation. Proc Natl Acad Sci USA 94, 8405–410. PELLETIER J. N., F. X. CAMPBELL-VALOIS, S. W. MICHNICK. 1998. Oligomerization domain-directed reassembly of active dihydrofolate reductase from rationally

43

44 Cellular Assays and Their Application in Drug Discovery

123

124

125

126

127

128

129

130

131

designed fragments. Proc Natl Acad Sci USA 95, 12141–6. GALARNEAU A., M. PRIMEAU, L. E. TRUDEAU, S. W. MICHNICK. 2002. Beta-lactamase protein fragment complementation assays as in vivo and in vitro sensors of protein–protein interactions. Nat Biotechnol 20, 619–22. BLAKELY B. T., F. M. ROSSI, B. TILLOTSON, M. PALMER, A. ESTELLES, H. M. BLAU. 2000. Epidermal growth factor receptor dimerization monitored in live cells. Nat Biotechnol 18, 218–22. REMY I., S. W. MICHNICK. 1999. Clonal selection and in vivo quantitation of protein interactions with protein-fragment complementation assays. Proc Natl Acad Sci USA 96, 5394–9. SUBRAMANIAM R., D. DESVEAUX, C. SPICKLER, S. W. MICHNICK, N. BRISSON. 2001. Direct visualization of protein interactions in plant cells. Nat Biotechnol 19, 769–72. MARSHALL E. S., K. M. HOLDAWAY, J. H. SHAW, G. J. FINLAY, J. H. MATTHEWS, B. C. BAGULEY. 1993. Anticancer drug sensitivity proﬁles of new and established melanoma cell lines. Oncol Res 5, 301–9. PETTY R. D., L. A. SUTHERLAND, E. M. HUNTER, I. A. CREE. 1995. Comparison of MTT and ATP-based assays for the measurement of viable cell number. J Biolumin Chemilumin 10, 29–34. AHMED S. A., R. M. GOGAL, Jr., J. E. WALSH. 1994. A new rapid and simple non-radioactive assay to monitor and determine the proliferation of lymphocytes. An alternative to [3 H]thymidine incorporation assay. J Immunol Methods 170, 211–24. SKAANILD M. T., J. CLAUSEN. 1989. Estimation of LC50 values by assay of lactate dehydrogenase and DNA redistribution in human lymphocyte cultures. ATLA, 293–6. CROUCH S. P., R. KOZLOWSKI, K. J. SLATER, J. FLETCHER. 1993. The use of ATP bioluminescence as a measure of cell proliferation and cytotoxicity. J Immunol Methods 160, 81–8.

132 MOSMANN T. 1983. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J Immunol Methods 65, 55–63. 133 GREEN D. R., J. C. REED. 1998. Mitochondria and apoptosis. Science 281, 1309–12. 134 MAJNO G., I. JORIS. 1995. Apoptosis, oncosis, and necrosis. An overview of cell death. Am J Pathol 146, 3–15. 135 WOO E. S., G. LAROCCA, O. LAPETS, G. R. BRIGHT. 1999. Multiparametric high content screening for apoptotic signals. High Throughput Screening Supplement to Biomed Products. 136 LEVEE M. G., M. I. DABROWSKA, J. L. LELLI, Jr., D. B. HINSHAW. 1996. Actin polymerization and depolymerization during apoptosis in HL-60 cells. Am J Physiol 271, C1981–92. 137 MAEKAWA T. L., T. A. TAKAHASHI, M. FUJIHARA, J. AKASAKA, S. FUJIKAWA, M. MINAMI, S. SEKIGUCHI. 1996. Effects of ultraviolet B irradiation on cell–cell interaction; implication of morphological changes and actin ﬁlaments in irradiated cells. Clin Exp Immunol 105, 389–96. 138 ENDRESEN P. C., P. S. PRYTZ, J. AARBAKKE. 1995. A new ﬂow cytometric method for discrimination of apoptotic cells and detection of their cell cycle speciﬁcity through staining of F-actin and DNA. Cytometry 20, 162–71. 139 van ENGELAND M., H. J. KUIJPERS, F. C. RAMAEKERS, C. P. REUTELINGSPERGER, B. SCHUTTE. 1997. Plasma membrane alterations and cytoskeletal changes in apoptosis. Exp Cell Res 235, 421–30. 140 CAMILLERI-BROET S., H. VANDERWERFF, E. CALDWELL, D. HOCKENBERY. 1998. Distinct alterations in mitochondrial mass and function characterize different models of apoptosis. Exp Cell Res 239, 277–92. 141 ANDERSON D. J., F. H. GAGE, I. L. WEISSMAN. 2001. Can stem cells cross lineage boundaries? Nat Med 7, 393–5. 142 HENRY D. 1997. Haematological toxicities associated with dose-intensive chemotherapy, the role for and use of

References

143

144

145

146

147

148

149

150

151

152

recombinant growth factors. Ann Oncol 8 (Suppl. 3), S7–S10. SCHEDING S., J. E. MEDIA, A. NAKEFF. 1994. Acute toxic effects of 3 -azido-3 -deoxythymidine (AZT) on normal and regenerating murine hematopoiesis. Exp Hematol 22, 60–5. EAVES C. J.. 1995. Assay of hematopoietic progenitor cells. In BEUTLER E., M. A. LICHTMAN, B. S. COLLIER, T. J. KIPPS (eds), Williams Hematology, 5th edn. New York: McGraw-Hill, 22–6. BRASIER A. R., D. RON. 1992. Luciferase reporter gene assay in mammalian cells. Methods Enzymol 216, 386–97. WARREN M. K., W. L. ROSE, L. D. BEALL, J. CONE. 1995. CD34+ cell expansion and expression of lineage markers during liquid culture of human progenitor cells. Stem Cells 13, 167–74. GREENWALT D. E., J. SZABO, I. MANCHEL. 2001. High throughput cell-based assay of hematopoietic progenitor differentiation. J Biomol Screen 6, 383–92. SAARINEN K., K. KIVISTO, K. BLOMBERG, K. PUNNONEN, L. LEINO. 2000. Time-resolved ﬂuorometric assay for leukocyte adhesion using a ﬂuorescence enhancing ligand. J Immunol Methods 236, 19–26. FIEHN O. 2002. Metabolomics – the link between genotypes and phenotypes. Plant Mol Biol 48, 155–71. NICHOLSON J. K., J. C. LINDON, E. HOLMES. 1999. “Metabonomics”: understanding the metabolic responses of living systems to pathophysiological stimuli via multivariate statistical analysis of biological NMR spectroscopic data. Xenobiotica 29, 1181–9. FIEHN O. 2002. Metabolomics –the link between genotypes and phenotypes. Plant Mol Biol 48, 155–71. LEHMANN M., W. BAUMANN, M. BRISCHWEIN, H. GAHLE, I. FREUND, R. EHRET, S. DRECHSLER, H. PALZER, M. KLEINTGES, U. SIEBEN, B. WOLF. 2001.

153

154

155

156

157

158

Simultaneous measurement of cellular respiration and acidiﬁcation with a single CMOS ISFET. Biosens Bioelectron 16, 195–203. LEHMANN M., W. BAUMANN, M. BRISCHWEIN, R. EHRET, M. KRAUS, A. SCHWINDE, M. BITZENHOFER, I. FREUND, B. WOLF. 2000. Non-invasive measurement of cell membrane associated proton gradients by ion-sensitive ﬁeld effect transistor arrays for microphysiological and bioelectronical applications. Biosens Bio-electron 15, 117–24. HENNING T., M. BRISCHWEIN, W. BAUMANN, R. EHRET, I. FREUND, R. KAMMERER, M. LEHMANN, A. SCHWINDE, B. WOLF. 2001. Approach to a multiparametric sensor-chip-based tumor chemosensitivity assay. Anticancer Drugs 12, 21–32. RININGER J. A., V. A. DIPIPPO, B. E. GOULD-ROTHBERG. 2000. Differential gene expression technologies for identifying surrogate markers of drug efﬁcacy and toxicity. Drug Discov Today 5, 560–8. SCHENA M., SHALON D., DAVIS R. W., BROWN P. O.. 1995. Quantitative monitoring of gene expression patterns with a complementary DNA microarrays. Science 270, 368–71. LOCKHART D. J., DONG H., BYRNE M. C., FOLLETTIE M. T., GALLO M. V., CHEE M. S., MITTMANN M., WANG C., KOBAYASHI M., HORTON H., BROWN E. L.. 1996. Expression monitoring by hybridization to high-density oligonucleotide arrays. Nat Biotechnol 14, 1649. GONZALES F., MORITA K., SEKI T., MIYATA N., UEKI T., LOIS A., MILAN L., MUNDAY C., GARCIA E., NAKAO T. 2001. Simultaneous HTS measurement of heme oxygenase and GAPDH mRNAs by multiStarTM High Performance Signal Ampliﬁcation (HPSATM ). Presented at SBS 7th Annual Conference & Exhibition, Baltimore, MD.

45

1

Phosphoproteome Analysis in Medical Research Elke Butt

Universit¨atsklinik W¨urzburg, W¨urzburg, Germany

Katrin Marcus Ruhr-University Bochum, Bochum, Germany

Originally published in: Proteomics in Drug Research. Edited by Michael Hamacher, Katrin Marcus, Kai St¨uhler, Andr´e van Hall, Bettina Warscheid and Helmut E. Meyer. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31226-9

1 Introduction

Proteomics has entirely shifted the paradigm for drug discovery and novel diagnostics, as it is an effective means to rapidly identify new biomarkers and therapeutic targets for the diagnosis and treatment of various diseases, while increasing our understanding of biological processes. Clinical applications of proteomics involve the use of proteomic technologies with the ﬁnal goal being to characterize the information ﬂow through the intra- and extracellular molecular protein networks that interconnect organ and circulatory systems. Protein phosphorylation, a reversible posttranslational modiﬁcation, occurs principally on serine, threonine, and tyrosine residues and plays a central role in cellular regulation either by altering a protein’s activity directly or by inducing speciﬁc protein–protein interactions, which in turn affect localization, binding speciﬁcity or activity. Abnormal kinase or phosphatase function plays a role in several pathologies and tumorigenesis. The phosphoproteome especially will play a key role in clinical proteomics as the aberrant function of protein kinases and phosphatases are often the center of many diseases, e.g., cancer, stroke, inﬂammatory and viral pathologies. The new focus of narrowly focused molecular targeted therapeutics addresses this concept [1]. A comprehensive analysis of protein phosphorylation includes the identiﬁcation of the phosphoprotein or phosphopeptide, the localization of the modiﬁed amino acid, and if possible, the quantiﬁcation of phosphorylation.

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Phosphoproteome Analysis in Medical Research

Several methods have been used to detect and identify phosphorylated residues in a protein. The ﬁrst step in a proteome study is the separation of a highly complex protein mixture resulting from a cell, tissue or organ lysate. In general, methods applied to the analysis of the proteome and protein identiﬁcation are also well suited for the analysis of the phosphoproteome, including the localization of speciﬁc phosphorylation sites. The most prominent method for this purpose today is twodimensional polyacrylamide gel electrophoresis (2D PAGE), which separates the proteins according to their isoelectric points in the ﬁrst and their molecular weights in the second dimension [2, 3]. The phosphoproteins can be visualized using different methods such as radiolabeling with [32 P]-orthophosphate combined with autoradiography [4], immunodetection with phosphospeciﬁc antibodies, and/or staining using phosphospeciﬁc dyes [5]. The speciﬁc isolation of phosphorylated proteins/peptides from a complex protein mixture is facilitated by different methods such as immunoprecipitation [6–8], immobilized metal afﬁnity chromatography [9, 10], or chemical modiﬁcation followed by speciﬁc puriﬁcation [11, 12]. Mass spectrometric identiﬁcation and localization of the phosphorylated amino acid residue can be done by several different methods such as mass spectrometry (MS)/MS-based peptide sequencing, precursor ion or neutral loss scanning [13–15] Indeed, the analysis of phosphoproteins and the detection and localization of the phosphorylated residues are not straightforward for various reasons: Ĺ The site of phosphorylation in a protein may vary resulting in a very heterogeneous mixture of different phosphorylation forms. Ĺ Phosphorylation in most instances emerges as a transient modiﬁcation within signaling cascades and is accompanied by dephosphorylation. Hence, the stoichiometry is generally relatively low and the phosphoprotein detection method must be very sensitive. Ĺ Most analytical techniques for studying protein phosphorylation have a limited dynamic range and only major phosphorylation sites may be detected, whereas minor sites might be difﬁcult to localize. Ĺ Phosphorylated peptides are generally suppressed in mass spectrometric analysis because of their low ionization rates.

In consequence, the localization of the phosphorylated residue often requires the speciﬁc isolation, detection and analysis of the modiﬁed region, which is complicated by the physicochemical conditions and complexity of the analyzed protein sample. The following three sections show concrete examples of the analysis of phosphoproteins as potential drug targets and demonstrate the applicability and enormous power of proteomics in this ﬁeld.

2 Phosphoproteomics of Human Platelets

Blood platelets arise as pinch-offs of megakaryocytes in the bone marrow, playing a critical role in wound healing. In their inactive, nonadhesive state they circulate

2 Phosphoproteomics of Human Platelets 3

freely in the blood. Interactions with structures in the subendothelial matrix caused by injuries of the blood vessels initiate a rapid platelet activation resulting in the formation of vascular plugs and release of intracellular substances (e.g., thrombin) that initiate repair processes. Platelet adhesion at the vessel wall is the ﬁrst step in platelet activation in vivo. Starting from this adhesion, intracellular signaling cascades are initiated resulting in secretion of activating substances and aggregation of the platelets. One important event within these intracellular signaling cascades is the phosphorylation/dephosphorylation of multiple proteins on various tyrosine, serine and threonine residues [16, 17]. Platelets are important components of hemostasis underlying cardiovascular diseases such as myocardial infarction or stroke [18, 19]. Genetic defects may result in bleeding disorders like Glanzmann thrombasthenia and Bernard-Soulier syndrome [20]. The high clinical relevance of arterial thrombosis, the limited knowledge about the underlying mechanisms on the molecular level, and the small number of available antiplatelet drugs [21] show the necessity for gaining more profound insights into the system. Although in recent years inhibitors of platelet aggregation have been established as the outstanding principle for secondary, and partially for primary prevention of cardiovascular diseases [22, 23], only a few effective drugs exist so far [21]. Hence, the search for better and perhaps safer antiplatelet drugs is of particular clinical interest, and it is therefore imperative to understand the exact molecular mechanisms in platelet activation, to ﬁnd new potential components of the signal transduction pathways and to determine the role of protein phosphorylation. In the study described, a differential phosphoproteomic approach using the combination of several methods – 2D PAGE, 1D PAGE, autoradiography, immunoblotting, and nano high-performance liquid chromatography (LC)-electrospray ionization (ESI) MS/MS – was employed, resulting in a comprehensive analysis of the platelet phosphoproteome. After ex vivo stimulation with thrombin and radiolabeling of the platelet (resting platelets served as control), proteins were separated using 2D PAGE and phosphorylated proteins were directly assigned by detection of [32 P] via autoradiography (Figure 1). Discrimination of serine/threonine-phosphorylated proteins on the one hand and tyrosine-phosphorylated proteins on the other was done by detection of phosphotyrosine-containing proteins via immunoblotting with an antiphosphotyrosine-speciﬁc antibody. Phosphorylated protein spots/bands were analyzed by nano-LC MS/MS. Several of the identiﬁed proteins were known to be involved in signal transduction and platelet function, e.g., cortactin, myosin regulatory light chain (myosin RLC), and protein disulﬁde isomerase (PDI). 2.1 Cortactin

In several spots of the immunoblots and the autoradiogram (Figure 2, white arrow) the actin-binding protein cortactin was identiﬁed. The human form of cortactin is known as an oncogene, often ampliﬁed in human colon carcinomas and tumor cells [24] and seems to be involved in the signal transduction caused by extracellular

4 Phosphoproteome Analysis in Medical Research

Fig. 1 Autoradiography of a twodimensional separation of phosphorylated human platelet proteins after labeling with [32 P] in the pH range 3–10. Top: Autoradiography of the 2D gel with pH 3–10 after application of 400 :g protein lysate of acti-

vated (A) and resting (B) human platelets. Below: Scaled-up sections I-V of the 2D gels. This ﬁgure clearly demonstrates the power of autoradiography in the differential phosphoproteome analysis.

stimuli such as thrombin, integrins, or the EGF [25]. On the basis of the MS/MS spectra of the nonphosphorylated and the phosphorylated peptide, a phosphorylation of Ser-418 was detected (Figure 3A, B). Additionally, two further phosphorylation sites were localized at Thr-401 and Ser-405 (results not shown). These sites were already described to be phosphorylated by the extracellular signal-regulated kinase (ERK) in vivo and in vitro [26]. Although the exact role of this phosphorylation is not yet clear, there is evidence for an important function in the regulation of the subcellular localization of cortactin [27]. Additionally, a tyrosine-phosphorylation of cortactin by the tyrosine kinase pp60c-src at different tyrosine residues (Tyr-421, Tyr-466, and Tyr-482) is known to be responsible for its reduced actin-crosslink activity [28].

2 Phosphoproteomics of Human Platelets 5

Fig. 2 Extracts of an anti-phosphotyrosine immunoblot (top) and autoradiography (bottom) after two-dimensional separation in the pH range 4–7 of resting and activated human platelets. (For better orientation a black circle is drawn in.) The extracts of the regions between pI 5.4–5.9 and 40–55 kDa clearly demonstrate the differences in the phosphorylation patterns of resting and thrombin-activated platelets. An intensity

increase of different protein spots identiﬁed as, e.g., cortactin (three different phosphorylated forms; white arrows) and protein disulﬁde-isomerase (black arrow) could be detected after stimulation with thrombin in both the immunoblot and the autoradiogram. Several of the identiﬁed proteins were known to be involved in signal transduction and platelet function.

2.2 Myosin Regulatory Light Chain

The regulatory light chains of myosin are known to be phosphorylated at different serine and threonine residues. Thr-18 and Ser-19 are substrates of the myosin light chain-kinase (MLCK) whereas Ser-1, Ser-2 und Ser-9 are phosphorylated by the protein kinase C (PKC) [29–31]. As the ﬁrst slow phosphorylation by the MLCK is already present in the resting platelets the second fast phosphorylation by the PKC occurred after thrombin stimulation. This second Ca2+ -dependent phosphorylation plays an important role in the secretion of substances from platelet intracellular granules [32]. In 1988, Naka and coworkers [29] described the existence of three different forms of myosin RLC in thrombin–activated platelets (non-, mono-, and bis-phosphorylated RLC) by nonproteomic studies. These results were conﬁrmed unequivocally and were even augmented by this proteomic study (Figure 3C, D) [7]. 2.3 Protein Disulﬁde Isomerase

PDI is a noncovalent homodimer with a molecular weight of 57 kDa for each subunit. The protein is generally involved in the formation, reduction and

6 Phosphoproteome Analysis in Medical Research

Fig. 3 Localization of the phosphorylation sites of cortactin (A and B) and the myosin regulatory light chain (RLC; C, D). A: Fragment ion spectrum of the nonphosphorylated peptide LPSpSPVYEDAASFK of cortactin. B: Fragment ion spectrum of the nonphosphorylated peptide LPSSPVYEDAASFK of cortactin. Due to the secessions of −98 Da from y11 Ser-418 (Ser-4 in the spectrum) was unambiguously detected to be phosphorylated. C: Fragment ion spectrum of the nonphosphorylated peptide AT-

SNVFAMFDQSQIQEFK of the RLC. D: Fragment spectrum of the phosphorylated peptide ApTSNVFAMFDQSQIQEFK or ATpSNVFAMFDQSQIQEFK, respectively. This peptide contained three possible phosphory-lation sites (Thr-2, Ser-3, and Ser-12). The comparison of the two spectra revealed Thr-2 or Ser-3 as phosphorylated residues. As no secession of −98 Da was observed till y13 a phosphorylation of Ser13 could be excluded. On the basis of the MS/MS spectrum the exact site of phosphorylation was not determinable.

rearrangement of disulﬁde bridges and plays an important role in platelet function. PDI is localized at the extracellular side of the plasma membrane and is secreted after platelet activation [33]. In platelets, PDI catalyzes the formation of disulﬁde-bridged complexes of thrombospondin 1 and thrombin-antithrombin III and is thus involved in hemostasis and wound healing [34]. In addition, it is part of the integrin-mediated platelet adhesion [35]. A threonine phosphorylation has been described for the PDI localized in the endoplasmatic reticulum [36].

3 Identiﬁcation of cAMP- and cGMP-Dependent 7

All proteins identiﬁed could be classiﬁed into different categories based on their function: 23 cytoskeletal proteins, 25 signaling molecules, 19 metabolic proteins, 5 vesicular proteins, 9 mitochondrial proteins, 6 proteins with extracellular function, 12 proteins involved in protein processing, 23 unknown proteins, and 12 miscellaneous. Overall, this proteome study resulted in a successful identiﬁcation of 134 proteins and the localization of eight partly unknown phosphorylation sites, clearly demonstrating the power of proteomics. Methods, procedures and results are described in detail in [7]. The identiﬁcation of phosphorylated proteins from such global approaches is only the ﬁrst step in characterizing these signaling molecules. It is important to verify the involvement of the proteins identiﬁed and to prove the localized phosphorylation sites in regard to their functionality. Such studies may be performed by using antibodies against the protein of interest and/or overexpression of wild-type and mutant forms of epitope-tagged proteins to examine their role in signal transduction pathways.

3 Identiﬁcation of cAMP- and cGMP-Dependent Protein Kinase Substrates in Human Platelets

The activation process of human platelets and vessel wall–platelet interactions are tightly regulated under physiological conditions and are often impaired in thrombosis, arteriosclerosis, hypertension, and diabetes. Platelet activation can be inhibited by a variety of agents, including aspirin and Ca2+ antagonists, as well as cGMP- and cAMP-elevating agents such as sodium nitroprusside (NO) and prostaglandin I2 [37]. The inhibitory effects of cGMP and cAMP are mediated by cAMP- and cGMP-dependent protein kinase (PKA and PKG), respectively [38]. For example, platelets from certain patients with chronic myelocytic leukemia showed decreased expression of PKG but a normal level of PKA. Phosphorylation of vasodilator-stimulated phosphoprotein (VASP) a speciﬁc substrate of both PKA and PKG, was severely impaired in these cGMP-dependent protein-kinase-deﬁcient platelets, despite an exaggerated cGMP response to sodium nitroprusside, whereas the cAMP-mediated VASP phosphorylation was functionally intact [39]. In PKGdeﬁcient mice cGMP-mediated inhibition of platelet aggregation is impaired [40]. However, the molecular mechanisms of platelet inhibition by cGMP signaling downstream of cGK activation are only partially understood and to date, only a few substrates for PKA and PKG have been identiﬁed and characterized in intact human platelets. In this respect, we used differential phosphoproteomic display of radiolabeled human platelets to identify speciﬁc substrates of PKG and PKA. Therefore, human platelets were labeled with 32 P, stimulated with 500 µM of the speciﬁc PKG activator 8-pCPT-cGMP, and proteins of the resulting platelet lysate were separated by 2D PAGE. The two-dimensional phosphoproteomes resulting from this experiment (Figure 4) demonstrate the phosphorylation/dephosphorylation of several proteins after stimulation with 8-pCPT-cGMP.

8 Phosphoproteome Analysis in Medical Research

Fig. 4 PKG-dependent phosphorylation in intact human platelets. Human platelets were incubated in the presence of [32 P]orthophosphate and treated with buffer alone (control) or with 500 :M PKG activator 8pCPT-cGMP for 30 min (stimulation). Platelet homogenate was separated by

two-dimensional gel electrophoresis and an autoradiogram was obtained. Spot 1: LASP; spots 2a, b and c: VASP; spot 3: rap 1b; spots 4, 5 and 8 have not yet been identiﬁed. The pH gradient is indicated at the top of the gel.

For identiﬁcation of these proteins, the spots were excised from the gel, digested with trypsin, and the resulting peptides were analyzed by nano LC-ESI MS/MS [41]. Alongside the increase of phosphorylation in several spots, dephosphorylation is also observed (Spots 4, 5 and 8) indicating the activation of phosphatases by activated PKG (Table 1). All PKG substrates identiﬁed so far in human platelets interact directly or indirectly with the actin ﬁlament and the cytoskeleton of the cell and are involved in the negative regulation of platelet activation after phosphorylation by PKG. The proteomic approach in combination with functional studies is a very successful tool for studying phosphorylation-dependent effects in platelets. Ongoing proteome studies will now focus on membrane proteins for new therapeutic targets to develop antithrombotic drugs.

4 Identiﬁcation of a New Therapeutic Target for Anti-Inﬂammatory Therapy by Analyzing Differences in the Phosphoproteome of Wild Type and Knock Out Mice

Genetically modiﬁed mice either overexpressing or knocking out speciﬁc components have provided exciting advances in our knowledge of diseases. Recently, we focused on the p38 mitogen-activated protein kinase (MAPK) pathway and its physiological effect on inﬂammation. An intact cytokine response is essential for efﬁcient host defense against invading pathogens. Inhibition of the p38 MAPK pathway that is involved in a number of cellular stress responses has been used successfully to decrease cytokine production in

4 Identiﬁcation of a New Therapeutic Target for Anti-Inﬂammatory Table 1 Identiﬁcation and function of cAMP- and cGMP-dependent protein kinase substrates

in human platelets Spot No.

Identiﬁcation

Function

1

LIM and SH3 domain protein LASP

2a, b and c

VASP

3

Rap 1b

Phosphorylation of LASP at Ser-146 leads to a redistribution of the actin-bound protein from the tips of the cell membrane to the cytosol accompanied with a reduced cell migration [41, 42] Three distinct phosphorylation sites (Ser-157, Ser-239 and Thr-278) with different speciﬁcities for PKA and PKG [43]. VASP phosphorylation is thought to be involved in the negative regulation of integrin α IIb bIII [45]. Experiments in vitro revealed reduced F-actin binding and actin polymerization of phosphorylated VASP [44]. Phosphorylation of Rap 1b at Ser-179 is associated with translocation of the protein from the membrane to the cytosol [46]

4, 5, 8 Not identiﬁed yet Gel not shown Heat shock protein 27 Hsp27

Phosphorylation of Hsp27 at Thr-143 by PKG reduced the stimulatory effect of MAPKAP kinase 2-phosphorylated Hsp27 on actin polymerization [47]

vitro [48], suggesting a key role for this MAPK pathway in inﬂammation. However, the central role of p38 MAPK, which is responsible for the activation of a number of kinases and transcription factors, limits the use of p38 inhibitors as a selective antiinﬂammatory strategy. Thus, downstream substrates of the MAPK pathway could represent promising targets for a speciﬁc suppression of cytokine production. In mouse and human, three MAPKAP kinases (MK2, MK3, and MK5) have been identiﬁed and are activated by p38. The substrate speciﬁcities of these kinases also seem very similar and the small heat shock protein HSP25/27 is an efﬁcient substrate for all three kinases in vitro [49]. Therefore, it was rather surprising that lack of MK2 resulted in a defect of cytokine biosynthesis and decreased levels of p38 MAPK. This was unexpected since MK3 and MK5 should be able to compensate for the loss of MK2, at least to a certain degree. To analyze these signiﬁcant differences, we deleted the MK2 and MK5 gene from mice and compared the phenotypes of the resulting MK5 knockout (KO) mice and the MK2-deﬁcient mice with the wild type (WT) [50]. For this approach WT, MEK2 -/- and MEK5 -/- mouse embryonic cardiac ﬁbroblasts were labeled with 32 P, stimulated by 100 µM arsenite and proteins of the cell lysates were separated by 2D PAGE. The differential phosphoproteomes (Figure 5) demonstrate the phosphorylation of a protein identiﬁed by MS as HSP25 (arrow) in wild type and MK5-deﬁcient cells indicating that in vivo, MK2 was the only kinase phosphorylating HSP25 in these cells stimulated by arsenite [50].

9

10 Phosphoproteome Analysis in Medical Research

Fig. 5 Autoradiograms of 2D phosphoproteomics from wildtype (WT), MAPKAP kinase 2 deﬁcient (MK2 −/−) and MAPKAP kinase 5 deﬁcient (MK5 −/−) mouse cardiac ﬁbroblasts before and after stimulation with 100 :M arsenite. The spot for phospho-HSP 25 was identiﬁed by mass spectrometry and is indicated by an arrow.

Obviously, MK5 is not able to compensate for this effect, making MK2 a unique pharmacological target for anti-inﬂammatory therapy. 5 Conclusions

In summary, modern proteomics approaches offer unprecedented opportunities to identify novel phosphorylated proteins for diagnosis and therapy of major illnesses. The development of new clinical proteomic tools away from 32 P labeling methods and the monitoring of protein expression by two-dimensional gels towards MSbased methods using phosphospeciﬁc isotope-tagged labeled peptides followed by one-dimensional electrophoresis will gain in importance. In the future, better protein enrichment strategies and quantiﬁcation of phosphorylation will allow the clinical transfer to functional application and early detection of pathological organ conditions, as a supplement to existing and coevolving advances in diagnostic imaging. Acknowledgements

Parts of the authors’ work were funded by the German Ministry of Education and Research (BMBF).

References

References 1 PETRICOIN, E., WULFKUHLE, J., ESPINA, V., LIOTTA, L. A. (2004). Clinical proteomics: revolutionizing disease detection and patient tailoring therapy. J. Proteome Res. 3, 209–217. 2 G¨ORG, A., POSTEL, W., G¨UNTHER, S., WESER, J. (1985). Improved horizontal two-dimensional electrophoresis with hybrid isoelectric focusing in immobilized pH gradients in the ﬁrst dimension and laying-on transfer to the second dimension. Electrophoresis 6, 599–604. 3 KLOSE, J. (1999). Large-gel 2-D electrophoresis. Methods Mol. Biol. 112, 147–172. 4 GARRISON, J. C. (1993). Protein phosphorylation. In HARDIE, D. G. (Ed.), A Practical Approach. Oxford University Press, Oxford, pp. 1. 5 SCHULENBERG, B., GOODMAN, T. N., AGGELER, R., CAPALDI, R. A., PATTON, W. F. (2004). Characterization of dynamic and steady-state protein phosphorylation using a ﬂuorescent phosphoprotein gel stain and mass spectrometry. Electrophoresis 15, 2526–2532. 6 MARCUS, K., IMMLER, D., STERNBERGER, J., MEYER, H. E. (2000). Identiﬁcation of platelet proteins separated by two-dimensional gel electrophoresis and analyzed by matrix assisted laser desorption/ionization-time of ﬂightmass spectrometry and detection of tyrosine-phosphorylated proteins. Electrophoresis 21, 2622–2636. 7 MARCUS, K., MOEBIUS, J., MEYER, H. E. (2003). Differential analysis of phosphorylated proteins in resting and thrombin-stimulated human platelets. Anal. Bioanal. Chem. 276, 973–993. 8 GRONBORG, M., KRISTIANSEN, T. Z., STENSBALLE, A., ANDERSEN, J. S., OHARA, O., MANN, M., JENSEN, O. N., PANDEY, A. (2002). A mass spectrometry-based proteomic approach for identiﬁcation of serine/threonine-phosphorylated proteins by enrichment with phospho-speciﬁc antibodies: identiﬁcation of a novel protein, Frigg, as a protein kinase A substrate. Mol. Cell Proteomics 1, 517–527.

9 SALOMON, A. R., FICARRO, S. B., BRILL, L. M., BRINKER, A., PHUNG, Q. T., ERICSON, C., SAUER, K., BROCK, A., HORN, D. M., SCHULTZ, P. G., et al. (2003). Proﬁling of tyrosine phosphorylation pathways in human cells using mass spectrometry. Proc. Natl. Acad. Sci. USA 100, 443–448. 10 FICARRO, S. B., MCCLELAND, M. L., STUKENBERG, P. T., BURKE, D. J., ROSS, M. M., SHABANOWITZ, J., HUNT, D. F., WHITE, F. M. (2002). Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nat. Biotechnol. 20, 301–305. 11 ODA, Y., NAGASU, T., CHAIT, B. T. (2001). Enrichment analysis of phosphorylated proteins as a tool for probing the phosphoproteome. Nat. Biotechnol. 19, 379–382. 12 ZHOU, H., WATTS, J. D., AEBERSOLD, R. (2001). A systematic approach to the analysis of protein phosphorylation. Nat. Biotechnol. 19, 375–378. 13 STEEN, H., MANN, M. (2002). A new derivatization strategy for the analysis of phosphopeptides by precursor ion scanning in positive ion mode. J. Am. Soc. Mass Spectrom. 13, 996–1003. 14 BATEMAN, R. H., CARRUTHERS, R., HOYES, J. B., JONES, C., LANGRIDGE, J. I., MILLAR, A., VISSERS, J. P. (2002). A novel precursor ion discovery method on a hybrid quadrupole orthogonal acceleration time-of-ﬂight (Q-TOF) mass spectrometer for studying protein phosphorylation. J. Am. Soc. Mass Spectrom. 13, 792–803. 15 MANN, M., ONG, S. E., GRONBORG, M., STEHEN, H., JENSEN, O. N., PANDEY, A. (2002). Analysis of protein phosphorylation using mass spectrometry: deciphering the phosphoproteome. Trends Biotechnol. 20, 261–268. 16 WATSON, S. P. (1997). Protein phosphatases in platelet function. In VON BRUCHHAUSEN, F., WALTER, U. (Eds.), Platelets and their Factors. Springer-Verlag, Berlin, pp. 297–325. 17 PRESEK, P., MARTINSON, E. A. (1997). Platelet protein tyrosine kinases cyclases. In VON BRUCHHAUSEN, F., WALTER, U.

11

12 Phosphoproteome Analysis in Medical Research

18

19

20

21

22

23 24

25

26

27

28

(Eds.), Platelets and their Factors. Springer-Verlag, Berlin, pp. 263–296. BROWN, A. S., ERUSALIMSKY, J. D., MARTIN, J. F. (1997). Megakaryocytopoiesis: the megakaryocyte/platelet haemi-static axis. In VON BRUCHHAUSEN, F., WALTER, U. (Eds.), Platelets and their Factors. Springer-Verlag, Berlin, pp. 3–19. GLASS, C. K., WITZUM, J. L. (2001). Atherosclerosis: The Road Ahead. Cell 104, 503–516. NURDEN, A. T. (1999). Inherited abnormalities of platelets. Thromb Haemost 82, 468–480. FITZGERALD, D. J. (2001). Vascular biology of thrombosis: the role of platelet–vessel wall adhesion. Neurology 57, S1–S4. LEFKOWITZ, R. J., WILLERSON, J. T. (2001). Prospects for cardiovascular research. JAMA 285, 581–587. BENNETT, J. S. (2001). Platelet-ﬁbrinogen interactions. Annu. Rev. Med. 52, 161–184. SCHUURING, E., VERHOEVEN, E., LITVINOV, S., MICHALIDES, R. J. (1993). The product of the EMS1 gene, ampliﬁed and over-expressed in human carcinomas, is homologous to a v-src substrate and is located in cell-substratum contact sites. Mol. Cell Biol. 13, 2891–2898. HUANG, C., NI, Y., WANG, T., GAO, Y., HAUDENSCHILD, C. C., ZHAN, X. (1997). Down-regulation of the ﬁlamentous actin cross-linking activity of cortactin by Src-mediated tyrosine phosphorylation. J. Biol. Chem. 272, 13911–13915. CAMPBELL, D. H., SUTHERLAND, R. L., DALY, R. J. (1999). Signaling pathways and structural domains required for phosphorylation of EMS1/cortactin. Cancer Res. 59, 5376–5385. van DAMME, H., BROK, H., SCHUURING-SCHOLTES, E., SCHUURING, E. (1997). The redistribution of cortactin into cell-matrix contact sites in human carcinoma cells with 11q13 ampliﬁcation is associated with both overexpression and posttranslational modiﬁcation. J. Biol. Chem. 272, 7374–7380. RAFFEL, G. D., PARMAR, K., ROSENBERG, N. (1996). In vivo association of v-Abl with Shc mediated by a non-phosphotyrosinedependent SH2 interaction. J. Biol. Chem. 271, 4640–4645.

29 NAKA, M., SAITOH, M., HIDAKA, H. (1988). Two phosphorylated forms of myosin in thrombin-stimulated platelets. Arch. Biochem. Biophys. 261, 235–240. 30 KAWAMOTO, S., BENGUR, A. R., SELLERS, J. R., ADELSTEIN, R. S. (1989). In situ phosphorylation of human platelet myosin heavy and light chains by protein kinase C. J. Biol. Chem. 264, 2258–2265. 31 MOUSSAVI, R. S., KELLEY, C. A., ADELSTEIN, R. S. (1993). Phosphorylation of vertebrate nonmuscle and smooth muscle myosin heavy chains and light chains. Mol. Cell Biochem. 127–128, 219–227. 32 NISHIKAWA, M., TANAKA, T., HIDAKA, H. (1980). Ca2+ -calmodulin-dependent phosphorylation and platelet secretion. Nature 287, 863–865. 33 CHEN, K., LIN, Y., DETWILER, T. C. (1992). Protein disulﬁde isomerase activity is released by activated platelets. Blood 79, 2226–2228. 34 MILEV, Y., ESSEX, D. W. (1999). Protein disulﬁde isomerase catalyzes the formation of disulﬁde-linked complexes of thrombospondin-1 with thrombin – antithrombin III. Arch. Biochem. Biophys. 361, 120–126. 35 LAHAV, J., GOFER-DADOSH, N., LUBOSHITZ, J., HESS, O., SHAKLAI, M. (2000). Protein disulﬁde isomerase mediates integrin-dependent adhesion. FEBS Lett. 475, 89–92. 36 QUEMENEUR, E., GUTHAPFEL, R., GUEGUEN, P. (1994). J. Biol. Chem. 269, 5485–5488. 37 KOESLING, D., NUERNBERG, B. (1997). Platelet G proteins and adenylyl and guanylyl cyclases. In VON BRUCHHAUSEN, F., WALTER, U. (Eds.), Platelets and their Factors. Springer-Verlag, Berlin, pp. 181–218. 38 M¨UNZEL, T., FEIL, R., M¨ULSCH, A., LOHMANN, S. M., HOFMANN, F., WALTER, U. (2003). Physiology and pathophysiology of vascular signaling controlled by guanosine 3 ,5 -cyclic monophosphate-dependent protein kinase. Circulation 108, 2172–2183. 39 EIGENTHALER, M., ULLRICH, H., GEIGER, J., HORSTRUP, K., HONIG-LIEDL, P., WIEBECKE, D., WALTER, U. J. (1993). Defective nitrovasodilator-stimulated protein phosphorylation and calcium regulation

References

40

41

42

43

44

45

in cGMP-dependent protein kinase-deﬁcient human platelets of chronic myelocytic leukemia. Biol. Chem. 268, 13526–13531. MASSBERG, S., SAUSBIER, M., KLATT, P., BAUER, M., PFEIFER, A., SIESS, W., FASSLER, R., RUTH, P., KROMBACH, F., HOFMANN, F. (1999). Increased adhesion and aggregation of platelets lacking cyclic guanosine 3 ,5 -monophosphate kinase I. J. Exp. Med. 189, 1255–1264. BUTT, E., GAMBARYAN, S., G¨OTTFERT, N., GALLER, A., MARCUS, K., MEYER, H. E. (2003). Actin binding of human LIM and SH3 protein is regulated by cGMP- and cAMP-dependent protein kinase phosphorylation on serine 146. J. Biol. Chem. 278, 15601–15607. KEICHER, C., GAMBARYAN, G., SCHULZE, E., MARCUS, K., MEYER, H. E., BUTT, E. (2004). Phosphorylation of mouse LASP-1 on threonine 156 by cAMP-and cGMP-dependent protein kinase. Biochem. Biophys. Res. Commun. 324, 308–316. BUTT, E., ABEL, K., KRIEGER, M., PALM, D., HOPPE, V., HOPPE, J., WALTER, U. (1994). cAMP- and cGMP-dependent protein kinase phosphorylation sites of the focal adhesion vasodilator-stimulated phosphoprotein (VASP) in vitro and in intact human platelets. J. Biol. Chem. 269, 14509–14517. HARBECK, K., H¨UTTELMAIER, S., SCHLU¨ TER, K., JOCKUSCH, B. M., ILLENBERGER, S. (2000). Phosphorylation of the vasodilator-stimulated phosphoprotein regulates its interaction with actin. J. Biol. Chem. 275, 30817–30825. HORSTRUP, K., JABLONKA, B., H¨ONIG-LIEDL,

46

47

48

49

50

P., JUST, M., KOCHSIEK, K., WALTER, U. (1994). Phosphorylation of focal adhesion vasodilator-stimulated phosphoprotein at Ser157 in intact human platelets correlates with ﬁbrinogen receptor inhibition. Eur. J. Biochem. 225, 21–27. TORTI, M., LAPETINA, E. G. (1994). Structure and function of rap proteins in human platelets. Thromb Haemost 71, 533–543. BUTT, E., IMMLER, D., MEYER, H. E., KOTLYAROV, A., LAASS, K., GAESTEL, M. (2001). Heat shock protein 27 is a substrate of cGMP-dependent protein kinase in intact human platelets: phosphorylation-induced actin polymerization caused by HSP27 mutants. J. Biol. Chem. 276, 7108–7113. LEE, J. C., LAYDON, J. T., MCDONNEL, P. C., GALLAGHER, T. F., KUMAR, S., GREEN, D., MCNULTY, D., BLUMENTHAL, M. J., HEYS, J. R., LANDVATTER, S. W. (1994). A protein kinase involved in the regulation of inﬂammatory cytokine biosynthesis. Nature 372, 739–746. STOKOE, D., ENGEL, K., CAMPBELL, D. G., COHEN, P., GAESTEL, M. (1992). Identiﬁcation of MAPKAP kinase 2 as a major enzyme responsible for the phosphorylation of the small mammalian heat shock proteins. FEBS Lett. 313, 307–313. SHI, Y., KOTLYAROV, A., LAASS, K., GRUBER, A. D., BUTT, E., MARCUS, K., MEYER, H. E., FRIEDRICH, A., VOLK, H. D., GAESTEL, M. (2003). Elimination of protein kinase MK5/PRAK activity by targeted homologous recombination. Mol. Cell Biol. 23, 7732–7741.

13

1

Mass Spectrometry of Peptides and Proteins Bettina Warscheid Ruhr-University Bochum, Bochum, Germany

Originally published in: Proteomics in Drug Research. Edited by Michael Hamacher, Katrin Marcus, Kai St¨uhler, Andr´e van Hall, Bettina Warscheid and Helmut E. Meyer. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31226-9 Abstract

Rapid advances in biological mass spectrometry have promoted proteomics to being a key technology in molecular cell biology and biomedical research. Current mass spectrometry-based proteomics provides the capability for the identification of cellular and subcellular proteomes, the study of changes in protein concentrations via stable isotope labeling methods and the mapping of functional protein modules isolated by means of affinity purification. The large potential of proteomics techniques to identify and accurately quantify proteins from complex biological samples has been well recognized in the larger scientific community and it will in all probability have a great impact on future drug research.

1 Introduction

In 1981, Barber et al. [1] demonstrated the analysis of a peptide of about 1300 Da in a mass spectrometer. To ionize the native peptide, the molecules were embedded in a glycerol matrix and bombarded with a high energy beam of argon atoms, a technique referred to as fast atom bombardment (FAB). Since ionization of hydrophobic peptides in the FAB process is favored compared to hydrophilic ones, the applicability of this technique to peptide mixture analysis is limited. Yet an avenue toward ionizing large biomolecules was apparent and encouraged the development of new ionization techniques, which also triggered the design of commercial mass spectrometers with expanded mass ranges. In 1988, Karas and Hillenkamp embedded proteins in a large molar excess of a UV-absorbing crystal matrix and irradiated the sample with a laser beam at suitable wavelengths [1]. In this process of matrix-assisted laser desorption/ionization Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Mass Spectrometry of Peptides and Proteins

(MALDI), large proteins can be transferred into the gas phase as intact molecules and become, for the most part, singly protonated. Since MALDI relies on the use of a pulsed laser, mass analysis is usually performed with a time-of-flight (TOF) analyzer of theoretically unlimited mass range. Since the mass spectrometer determines the mass-to-charge (m/z) ratio of the molecules, the ability to generate multiply charged ions would allow for the use of instruments with limited mass ranges in peptide and protein analysis. This was realized by Fenn and coworkers in the 1980s when they sprayed a diluted solution of protein in a high voltage