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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

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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

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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

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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

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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],

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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

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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].

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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.

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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.

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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.

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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].

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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.

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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.

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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.

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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).

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18 Photoproteins as Reporters in Whole-cell Sensing

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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.

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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.

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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].

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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

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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

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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

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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.

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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

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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

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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

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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

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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

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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.

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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

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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

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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

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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

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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.

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29

30

31

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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.

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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.

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22 The COP9 Signalosome and Its Role in the Ubiquitin System

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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.

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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.

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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.

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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.

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24 The Deubiquitinating Enzymes

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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

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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).

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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].

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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.

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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

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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

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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

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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.

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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.

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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.

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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

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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.

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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.

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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.

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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.

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24 Tailoring Protein Scaffolds for Ligand Recognition

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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

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(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.

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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

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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.

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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

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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]).

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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.

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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.

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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

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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.

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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

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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.

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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.

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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

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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).

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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-

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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

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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.

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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

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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

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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

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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].

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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

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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.

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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

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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