Chemical tools for activity-based proteomics

Journal of Biotechnology 124 (2006) 56–73

Review

Chemical tools for activity-based proteomics Miriam C. Hagenstein, Norbert Sewald ∗ Organic and Bioorganic Chemistry, Department of Chemistry, University of Bielefeld, Universit¨atsstrasse 25, 33615 Bielefeld, Germany Received 22 July 2005; received in revised form 27 October 2005; accepted 1 December 2005

Abstract Several approaches for proteome analysis and the generation of proteome subsets rely on engineered chemical probes that are tailored towards the detection of different protein classes. The concepts are presented in this review covering the literature until mid-2005. © 2005 Elsevier B.V. All rights reserved. Keywords: Proteomics techniques; Activity-based profiling; Chemical probes; Inhibitors; Protein ligands

Contents 1. 2.

3.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Activity-based protein profiling/activity-based proteomics/chemical proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Reporter groups/affinity tags . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Linkers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Strategies for ABPP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.1. Non-directed ABPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.2. Directed ABPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Proteomics technologies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion and outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

57 59 60 60 60 60 63 68 70 71

Abbreviations: ABP, activity-based probe; ABPP, activity-based protein profiling; ␣-CA, ␣-chloroacetamide; CID, collision induced dissociation; 2-DE, 2-dimensional electrophoresis; FAL, fluorescent affinity label; HTS, high throughput screening; IAC, inhibitor affinity chromatography; ICAT, isotope-coded affinity tagging; LC, liquid chromatography; MS, mass spectrometry; MS/MS, tandem mass spectrometry; MudPIT, multidimensional protein identification technology; PAGE, polyacrylamide gel electrophoresis; PEG, poly(ethylene glycol); pI, isoelectric point; PS, phenyl sulfonate ester; PTM, post-translational modification; Rh, rhodamine; RP, reversed phase; SDS, sodium dodecylsulfate ∗ Corresponding author. Tel.: +49 521 106 2051; fax: +49 521 106 8094. E-mail address: [email protected] (N. Sewald). 0168-1656/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.jbiotec.2005.12.005

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1. Introduction The increasing number of completely sequenced genomes of both prokaryotic and eukaryotic organisms confronts researchers with the enormous task of assigning molecular and cellular functions to the overwhelming amount of newly predicted gene products (mRNA and proteins). In order to meet this challenge, the so-called “proteomics” has emerged. The term “proteome” was originally coined by Mark Wilkins and Keith Williams at Macqarie University in Australia, in 1994 to describe the “PROTEin complement of the genOME”. “Proteomics” can be defined as the qualitative and quantitative comparison of proteomes under different conditions (Wasinger et al., 1995). While there exist well-established strategies for gene analysis (“genomics”), which allow the comparative analysis of a given mRNA-complement in a single experiment (e.g. cDNA or oligonucleotide microarrays) (Dharmadi and Gonzalez, 2004), the situation is different when it comes to the investigation of proteins (“proteomics”). Proteome analysis is technically challenging, because the number of different proteins which are expressed at a given time ranges from several thousand for simple prokaryotic organisms up to at least 10 000 in eukaryotic cells. To further complicate proteome investigations, proteins have highly heterogenous physical and chemical properties (membrane-bound versus soluble proteins, extreme pI values, low abundance versus high abundance proteins, posttranslational modifications etc.). There is no amplification method comparable to PCR for DNA/mRNA for low abundance proteins and no general protein binding partners (like, e.g. anti-sense DNA) exist. Therefore, simultaneous characterization of the relative abundance and activity of all proteins of a given proteome is not possible with one single strategy. In contrast to the genome, the proteome is a highly dynamic system, which is influenced by small environmental changes (e.g. changes in bacteria culture conditions, addition of drugs to a cell culture, stress, etc.) in a characteristic manner. A striking example is that both a caterpillar and the accordant butterfly have the same genome, but different proteomes. Proteome analysis is only reasonable in differential approaches, where two or more well-defined states can be compared. Changes in the protein patterns of the

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different states (e.g. cells before and after drug treatment, healthy and pathological states) are monitored and evaluated quantitatively. Hence, the proteome can be considered as a unique, highly sensitive monitor for complex metabolic and regulatory correlations in an organism (Lottspeich, 1999). The most widely used technique for protein separation, quantification and identification is based on two-dimensional polyacrylamide gel electrophoresis (2-DE) (O’Farrell, 1975) in combination with protein staining and mass spectrometry (MS) of excised and digested protein spots. 2-DE usually combines isoelectric focussing with polyacrylamide gel electrophoresis and allows the simultaneous resolution of more than 5000 proteins (depending on gel size and pH gradient), which are separated according to pI and molecular mass (G¨org et al., 2004). The introduction of immobilized pH gradient strips (IPGs) for the first dimension enhanced the capacity/efficiency of this method with respect to former shortcomings like reproducibility, resolution, separation of very acidic/basic proteins and sample loading capacity (G¨org et al., 1988). However, this method still lacks the ability to resolve important protein classes like membrane-associated and low abundance proteins (Gygi et al., 2000; Santoni et al., 2000). Regarding membrane proteins, better results are obtained when SDS-PAGE is combined with reversedphase liquid chromatography–tandem mass spectrometry (LC–MS/MS) analysis of gel lane fractions. This method has been named “geLC–MS/MS” (Rappsilber et al., 2002; Li et al., 2003). Traditionally, analysis of two or more proteomes to identify differentially expressed proteins with 2-DE is done by separation of each protein sample on an individual gel, followed by staining and quantification. The different gels are then compared with computer aided image analysis programs, but since perfect superimposition often cannot be realized, image analysis remains often a time consuming matter. This problem can be avoided by using the fluorescent 2D differential gel electrophoresis (DIGE) developed by Unlu et al. (1997) which allows the comparison of two different samples on the same 2D-gel: two protein mixture samples corresponding to two different proteome states are labeled with different fluorescent cyanine minimal dyes (Cy3 and Cy5, Amersham Biosciences), which are size- and charge-matched, as well

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as spectrally distinct (different excitation and emission wavelength). This allows co-separation of the differently labeled samples in the same gel and ensures that all samples will be subject to exactly the same first and second dimension electrophoresis conditions, limiting experimental variation and ensuring accurate withingel matching. After excitation at the appropriate wavelengths, the images are overlaid and subtracted, visualizing only differences in protein abundance (Gharbi et al., 2002). Alternatively, all protein spots may be excised, digested, and analyzed by MS automatically. “Gel-free” methods such as the multidimensional protein identification technology (MudPIT) or (cleavable) isotope-coded affinity tagging [(c)ICAT] are an alternative to the gel electrophoresis-based strategies. They are based on liquid chromatography (LC) for separation and MS for identification. MudPIT allows the separation and identification of complex protein and peptide mixtures by twodimensional (or multidimensional) liquid chromatography (2D-LC), followed by MS/MS. The proteins are digested by a protease and the peptide mixture is applied to a strong cation exchange column (first dimension), from which the peptides are successively eluted with increasing salt concentrations onto a reversed-phase (RP) column (second dimension), where they are separated according to hydrophobicity. The peptide fractions are submitted online to MS/MS and identification is achieved by sequence database search (Link et al., 1999; Washburn et al., 2001; McDonald et al., 2002). Brittain et al. (2005) recently introduced the concept of “fluorous proteomics”, whereby specific peptide subsets are tagged with perfluorinated moieties that display interesting interaction behavior with aqueous, organic, and perfluorinated solvents or phases, respectively. Subsequently, the tagged peptides are enriched by solid-phase extraction over a fluorousfunctionalized stationary phase. The ICAT-technique is based on differential isotopic labeling of identical proteins or peptides derived from two different proteome states with light or heavy tags. The tags comprise a thiol-specific reactive group, a biotin moiety and – in the case of the heavy variant – several heavy isotopes like deuterium, 13 C or 15 N, providing probes that have different masses, but are chemically identical. The differently tagged sam-

ples are pooled, digested and purified by avidin affinity chromatography. This step reduces the complexity by factor 10 (Gygi et al., 2002), since only Cys-containing peptides are analyzed, and simplifies the separation process. The labeled peptides are then subjected to (multidimensional) LC followed by MS, where relative quantification of simultaneously eluted light- and heavy-labeled peptides is achieved via comparison of the relative signal intensities. Identification of the peptides is achieved with MS/MS and database search (Gygi et al., 1999, 2002; Smolka et al., 2002). The cleavable isotope-coded affinity tagging (cICAT) is a closely related technique, where the tags incorporate a cleavage site. Zhou et al. (2002) describe a solid-phase isotope tagging reagent that contains a photocleavable linker. Cysteine-containing peptides are captured on solid support, removed from the mixture, and released while being isotope-labeled after photoinduced cleavage. Hansen et al. (2003) report the use of a commercial ICAT reagent (Applied Biosystems, Framingham, MA) that contains biotin and an acidlabile linker group, allowing the removal of the biotin moiety after ICAT-labeling and biotin-/avidin-based peptide affinity isolation. This enhances the quality of CID MS spectra (Li et al., 2003). The drawback of the (c)ICAT methodology is that only cysteine-containing proteins and peptides can be analyzed. Therefore, the sequence information of proteins is vastly reduced. Furthermore, (c)ICAT is not a good option for the analysis of posttranslational modifications (PTMs), since the probability of a peptide to contain both a PTM and a cysteine residue is relatively low. Schmidt et al. (2005) presented a method capable of high-throughput proteome profiling on a global scale that circumvents these problems: isotope-coded protein label (ICPL). This method is based on tagging all free amino groups of intact proteins from two different cell states with light or heavy ICPL tags prior to fractionation on the protein and/or after digestion on the peptide level. This method is compatible with all commonly used protein and peptide separation techniques to reduce complexity and provides high protein sequence information, including PTMs and isoforms. In contrast to the relative quantification strategies just described, the use of stable isotopes provides also the means for absolute protein quantification. Gerber et al. (2003) described a strategy named AQUA (Absolute QUAntification). They synthesized peptides cor-

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responding to the native counterparts formed by proteolysis of proteins of interest, but with incorporated stable isotopes. Thereby they obtained an internal peptide standard (AQUA peptides) for MS/MS analysis. Protein mixtures are fractionated by SDS-PAGE, the regions containing the proteins of interest are excised and submitted to in-gel proteolysis in the presence of AQUA peptides. Subsequent LC–MS/MS analysis allows absolute quantification of selected peptides by comparison of the relative signal intensities of native and corresponding AQUA peptides. Bottari et al. (2004) developed the so-called visible isotope-coded affinity tag (VICAT) which also allows absolute quantification of target proteins in complex samples. Like ICAT, VICAT reagents tag thiol groups of cysteines or thioacetylated amino groups. They introduce a biotin affinity handle, a photocleavable linker for removing a portion of the tag, and an isotope tag for distinguishing sample and internal standard peptides. Internal standard peptides are obtained by tagging synthetic peptides (derived from the protein of interest) with the heavy VICAT-variant. The reagent also may include a moiety that can be visualized by autoradiography (14 C) for tracking target peptides in a gel-based separation technique such as isoelectric focusing. Mixtures of tagged sample-derived peptides and internal standard peptides are submitted to isoelectric focussing (IEF). The “visible” moiety reveals the position of the peptides of interest which are eluted and purified by affinity chromatography with streptavidin–agarose followed by photocleavage. The purified peptides are then submitted to micro-LC/ESIMS for quantification. This strategy was used successfully to determine the absolute abundance of human group V phospholipase A (a protein with relatively low abundance) in a cell lysate (Lu et al., 2004). Recently, Ross et al. (2004) published a technique named iTRAQ (Applied Biosystems, Framingham). It is based upon chemically tagging the primary amines (N-terminus or lysine) of peptides generated from protein digests with isobaric tags (isobaric species that have the same atomic mass but different arrangements) that release characteristic low mass reporters ions upon fragmentation (CID, MS/MS). This allows comparison and (absolute) quantification of up to four samples in one experiment. The use of isobaric tags provides a

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sensitivity enhancement over mass difference labeling since there is no splitting of the MS precursor signal and no increase in spectral complexity. The same is true for the MS/MS spectra because the peptide backbone fragment ions are isobaric, too. Since every peptide is labeled, proteome coverage is expanded and tracking of PTMs is possible (although detection may be difficult). One major drawback of this method is that the acquisition of MS/MS spectra is mandatory which requires longer analysis time. The high complexity on the peptide level may cause other problems, since it is impossible to tell how many peptides with coincidentally identical masses are present in the mixture. Because all of them will release their reporter ions, quantification may be inaccurate. All strategies that have been mentioned so far measure changes in protein abundance (abundance-based proteomics), but do not analyze the activity and the function of the enzymes. Since changes in the phenotype may either be due to changes in the overall amount of a given protein, but as well may depend on the fraction of active protein, other investigation methods have been developed, known as activity-based protein profiling (ABPP), activity-based proteomics or chemical proteomics.

2. Activity-based protein profiling/activity-based proteomics/chemical proteomics The basic concept of chemical proteomics comprises the application of relatively small (compared to proteins) molecules which covalently label the active site of enzymes. The use of selective molecules that can be tuned in their affinity to different target enzymes generates “subproteomes” prior to further separation, digestion, and MS analysis, thereby decreasing the amount of data that has to be processed. This method allows for facile screening/detection of the problem childs in protein analysis, low-abundance and membrane-associated proteins, via, e.g. affinity enrichment based on biotin tags. Since often only active proteins are detected while inactive proforms (Kidd et al., 2001) and inhibitor bound enzymes (Liu et al., 1999; Jessani et al., 2002; Saghatelian et al., 2004; Greenbaum et al., 2000; Greenbaum et al., 2002) remain unlabeled, direct assignment of effector

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M.C. Hagenstein, N. Sewald / Journal of Biotechnology 124 (2006) 56–73

Fig. 1. Activity-based probe.

proteins responsible for specific biological events is possible. The tools which constitute chemical proteomics, the so-called activity-based probes (ABP), basically comprise at least one reporter group (which serves for visualization and/or purification) (Adam et al., 2002a) coupled via a linker to a recognition unit and a reactive group that establishes a covalent bond to the protein. In many cases, the recognition unit and the reactive group belong to the same moiety, e.g. irreversible inhibitors that establish a covalent bond to the active site. Other examples are the so-called suicide substrates where the reactive group is created in the course of the enzyme reaction (see below) (Fig. 1). Additional recognition units may be necessary/useful to promote active site-directed interactions of rather unspecific electrophiles with specific enzyme subsets, whereas in the case of reversible inhibitors or ligands as recognition units an additional reactive group (e.g. photocrosslinker) to form the covalent bond (not necessarily to the active site) is required. 2.1. Reporter groups/affinity tags Reporter groups should allow either rapid and sensitive detection of labeled proteins or affinity enrichment prior to subsequent identification. The most commonly used species are fluorescent dyes and radioisotope labels for in-gel visualization, and biotin for visualization after Western blotting as well as for affinity enrichment employing, e.g. streptavidin-coated beads. 2.2. Linkers The main purpose of a linker moiety in an ABP is to connect the probe head (recognition unit) with a reporter group/affinity tag and, depending on the probe head, with an additional reactive group (e.g. photoreactive group). Furthermore, the length of the linker provides sufficient space between probe head

and tag to avoid steric hindrance or interference at the active site. The most simple linker options are either long-chain alkyl or polyethylene glycol (PEG) spacers, which allow tuning of the hydrophobic properties of the ABP according to the requirements (membrane penetration versus solubility in aqueous solutions). The development of cleavable linkers comparable to the ones used in cICAT-technology will play an important role regarding biotin-based analysis methods, since the elution of streptavidin bound biotinylated proteins after affinity enrichment is often problematic due to the strong biotin–streptavidin bond (Zhou et al., 2002; Hansen et al., 2003; Li et al., 2003). 2.3. Strategies for ABPP Two different strategies for activity-based protein profiling (ABPP) have been developed so far: (1) a non-directed (combinatorial) approach in which libraries of active site-directed non-specific electrophiles are screened against complex proteomes for activity-dependent protein reactivity, and (2) a directed approach which relies on well-known inhibitors and/or affinity labels (ligands) to profile the functional state of specific classes of enzymes. 2.3.1. Non-directed ABPs The concept of non-directed ABPs has been introduced by Adam et al. (2001) to expand the number of enzymes which can be addressed by ABPP, since inhibitors or affinity labels (ligands) are not yet known for every enzyme class. They synthesized a small combinatorial library of biotinylated sulfonate esters comprising the following elements: a variable alkyl/aryl binding group to address different enzyme active sites in the proteome, a sulfonate ester reactive group, an aliphatic linker and biotin (Fig. 2). Sulfonate esters were chosen as the reactive group, anticipating that a carbon electrophile may have the ability to label enzymes within as well as across different enzyme classes. Furthermore, moderately reactive electrophiles should react preferentially in enzyme active sites since these are enriched in nucleophilic residues. Each sulfonate was reacted with a native and a heat-denatured version of a proteome. Analysis by SDS-PAGE and avidin blotting revealed heat-sensitive and therefore specific labeling of several proteins as

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Fig. 2. Non-directed sulfonate ester ABPs.

well as considerably different relative reactivities of the sulfonate probes with individual proteins. The authors hypothesized, based on the heat-sensitive labeling, that binding of the sulfonate esters takes place in a structured part of the proteins, often representing either a ligand binding pocket of a receptor or an enzyme active site. The fact that one protein that could be identified as a class I aldehyde hydrolase (ALDH-1) was irreversibly inhibited by some sulfonates is consistent with the assumption that labeling occurs at or near the active site.

Conjugation of the sulfonate ester library to rhodamine instead of biotin (to gain sensitivity and throughput) supplied evidence for labeling of at least six mechanistically distinct enzyme classes, with strong indications that this happens in the active sites (Adam et al., 2002b). To simplify and accelerate the detection and identification of target proteins, trifunctional probes comprising a sulfonate ester group, biotin, and rhodamine were synthesized later on (Fig. 3), consolidating target detection and purification steps.

Fig. 3. Trifunctional sulfonate ester probe.

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Table 1 Enzyme targets of non-directed sulfonate ester ABPs Enzyme

Enzyme class

Aldehyde dehydrogenase 1a Aldehyde dehydrogenase 7b Acetyl-CoA acetyltransferaseb Dihydrodiol dehydrogenaseb

Aldehyde dehydrogenase Aldehyde dehydrogenase Thiolase NAD/NADP-dependent oxidoreductase Enoyl-CoA hydratase Epoxide hydrolase Glutathione S-transferase 3␤-Hydroxysteroid dehydrogenase Phosphofructokinase Transglutaminase

Enoyl-CoA hydratase, peroxisomalb Epoxide hydrolase, cytoplasmicb GSTO1-1b 3␤-Hydroxysteroid dehydrogenase
5-isomerase-1c Platelet phosphofructokinase (pPFK)c Type II tissue transglutaminase (tTG)c a b c

Adam et al. (2001). Adam et al. (2002b). Adam et al. (2002a).

Thereby, three more target proteins could be identified, adding up to a total of nine mechanistically distinct classes (Adam et al., 2002a) (Table 1). None of the enzymes labeled by the sulfonate library represented a target of precedent directed approaches, therefore emphasizing the benefits of chemically and structurally diverse probe libraries to expand the number of enzyme classes susceptible to profiling and to identify novel protein markers (Speers and Cravatt, 2004a). In some cases, sulfonate labeling correlates directly with catalytic activity. Labeling of the tissue transglutaminase in fact occurs in an activity-based manner since it was susceptible to the presence of calcium (physiological activator) and GTP (natural allosteric inhibitor). Similar results were obtained for the labeling of platelet phosphofructokinase (pPFK) which is influenced by the allosteric inhibitor ATP (Adam et al., 2002a). However, in other cases labeling may occur at noncatalytic residues in the active site, therefore not being activity-based from a purely mechanistic standpoint. To determine whether catalytically active residues are affected or not, the respective modification site has to be identified (see below). However, it may be appropriate to generally speak of activity-based profiling if one considers a more biological perspective: any probe that is sensitive to molecular interactions which often regulate enzyme activity in vivo (autoinhibitory domains, protein partners, small molecules that sterically obstruct

the active site) (Kobe and Kemp, 1999) would provide information about the functional state of the enzyme in the context of the cell biology of the proteome (Adam et al., 2002c). Adam et al. (2004) developed a gel-free strategy to identify the sites of labeling on five enzymes targeted by sulfonate ester probes. Proteomes were treated with a rhodamine-tagged phenyl sulfonate ester probe (PS-rhodamine), followed by denaturation, thiol reduction, and alkylation. After tryptic digestion and affinity capture with immobilized anti-rhodamine antibodies, labeled peptides were analyzed by LC–MS/MS and the SEQUEST search algorithm (Eng et al., 1994) to concurrently identify the protein targets as well as the specific labeled residues. It was found that PS-rhodamine labeled conserved active site residues in each enzyme, either playing important roles in catalysis or with unknown function so far. These results suggest that PS-rhodamine indeed acts as an activity-based proteomic probe for most of its enzyme targets. In vivo applications of ABPP are technically challenging due to the bulky reporter tags (molecular mass >700–1000 Da) present in ABPs. Their physical properties may influence the membrane permeability of the probes as well as the probe binding affinity, hampering cellular uptake and distribution. In general small uncharged molecules are supposed to have improved access to intracellular and extravascular compartments. Therefore, it would be advantageous to covalently label proteins first in vivo with an unsuspicious probe, and to attach the reporter group afterwards. Following these considerations, the probe would require only a reactive chemical functionality which remains unchanged under usual physiological conditions but which serves as an anchor to covalently attach the reporter group (fluorophore or biotin or radiolabel) at a later stage. Such bio-orthogonal chemical moieties are, e.g. azido groups that can be selectively modified with phosphines in the course of the Staudinger ligation (Prescher and Bertozzi, 2005) or activated alkynes. Alternatively terminal alkynes may be subsequently ligated with azides to a 1,2,3-triazole in a Cu(I)catalyzed stepwise analogue of Huisgen’s 1,3-dipolar cycloaddition developed by Sharpless and colleagues (“click chemistry”, Fig. 4) (Kolb et al., 2001; Lewis et al., 2002; Rostovtsev et al., 2002; Wang et al., 2003a).

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Fig. 4. Copper(I)catalyzed synthesis of 1,2,3-triazoles.

Speers et al. (2003) exploited the suitability of sulfonate esters for these purposes. They synthesized a rhodamine-alkyne tag (Rh-≡) and an azide-derivatized phenyl sulfonate ester reactive group (PS-N3 ), thereby engineering a pair of biologically inert coupling partners. It was demonstrated that treatment of cells or mice with PS-N3 , followed by reaction of cell or tissue homogenates with Rh-≡ gave strong fluorescent signals, indicating that azide-alkyne-cycloaddition is indeed suited for activity-profiling in vivo. Besides, click chemistry allows easy adaptation of the fluorophore or affinity tag according to the needs, since every alkyne-derivatized reporter group is suitable. Ovaa et al. (2003) reported a comparable tag-free approach for the directed in vivo-labeling of active proteasomes with vinyl sulfones (Table 2), using a modified Staudinger ligation to attach a reporter group to the azide-modified inhibitor. An initial drawback of the click chemistry ABPP was the higher background labeling as compared to conventional ABPP due to non-specific labeling of abundant proteins with Rh-≡. This problem could be overcome by exchanging the azide and alkyne groups (PS-N3 /Rh-≡ to PS-≡/Rh-N3 ), but only at the expense of a lower reaction rate (Speers and Cravatt, 2004b). Recently, Barglow and Cravatt (2004) reported the synthesis of another library of candidate ABPP probes being structurally more diverse than the sulfonate ester library. For this purpose an ␣-chloroacetamide (␣CA) was chosen as non-directed electrophilic reactive group, attached to a variable dipeptide as recognition group (Fig. 5). Among the labeled enzymes, targets of already known ABPs (sulfonate esters (Adam et al., 2002b) and epoxides (Greenbaum et al., 2002)) were found. In contrast, some enzymes exclusively targeted by ␣-CA probes in an activity-based manner were detected as well, while the primary targets of the phenyl sulfonate ester (ALDH-1 and enoyl CoA hydratase 1, see above) remained unlabeled, indicating that the two

Fig. 5. Non-directed ␣-chloroacetamide probe.

probe types target complementary portions of proteomic space. Bioinformatic analysis of the reactivity profiles of ∼25 proteins with the dipeptide ␣-CA probes allowed classification of the probes into discrete subgroups, with subgroup members mostly sharing common features like being positively/negatively charged or hydrophobic, etc. Barglow and Cravatt (2004) assembled an “optimal probe set” containing representative members of each probe subgroup, which allowed to collectively profile the majority of targets labeled by the entire probe library, thereby enabling the screening of proteomes with maximal efficiency and minimal sample consumption. The “optimal probe set”-strategy was successfully applied to profile liver tissue from lean (wt) and obese mice, discovering several obesityassociated enzyme activities. 2.3.2. Directed ABPs A major goal in the development of molecular tools for functional proteomics is the retrieval of molecules that bind to many or all representatives of a class of proteins in a family-specific manner, which would allow to label them with reporter groups or to selectively enrich them during affinity chromatography. The common mechanistic basis is a central feature for such directed molecular tools. Depending on the nature of the recognition unit (ligand, inhibitor) the representatives of this class of compounds are further divided into two sub-groups: directed irreversible and directed reversible inhibitors/ligands. 2.3.2.1. Irreversible Inhibitors. The members of the irreversibly binding recognition units in such conjugates may be sub-divided into two classes comprising (a) reactive mechanism based probes and (b) suicide

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Table 2 Directed irreversible inhibitors Target enzymes

Reactive group

References

Serine hydrolases Liu et al. (1999), Kidd et al. (2001), Patricelli et al. (2001)

Tyrosine phosphatases Kumar et al. (2004)

Lo et al. (2002), Zhu et al. (2003)

Cysteine proteases Schaschke et al. (1998), Schaschke et al. (2000), Greenbaum et al. (2000), Greenbaum et al. (2002)

Bogyo et al. (2000), Nazif and Bogyo (2001), Wang et al. (2003b), Ovaa et al. (2003)

Thornberry et al. (1994), Faleiro et al. (1997), Martins et al. (1997), Kato et al. (2005)

Winssinger et al. (2002)

Liau et al. (2003)

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substrate probes (Jeffrey and Bogyo, 2003). The difference between these two classes is that in the latter case the substrate is transformed in the course of the enzyme reaction into a highly reactive species able to undergo covalent bond formation with the active site of the enzyme. In the former case the high reactivity of the probe, e.g. towards nucleophiles is already present in its original form. The directed irreversible affinity based probes usually are conjugated across a linker to an appropriate reporter group. Most interestingly, in such cases the ability of the probe to bind to the corresponding enzyme family is intimately connected with enzyme activity. Hence, this method can be used for activity profiling because it will detect only active forms of proteins. Representative examples of pairs between protein families and irreversibly binding ligands in such directed probes are compiled in Table 2. Serine proteases can be addressed with fluorophosphonates (Table 2) that are well known to phosphorylate the catalytic serine residue in this class of enzymes. Serine hydrolases form one of the largest families of enzymes present in eukaryotes with an enormous diversity. It has been estimated that this class of enzymes represents approximately 1% of the predicted protein products encoded by the human genome (Lander et al., 2001). Consequently, such conjugates may be of high relevance not only in the field of serine proteases, because lipid hydrolases, esterases and amidases also belong to the family of serine hydrolases. As the fluorophosphonate probes react with the serine residue in the active site of the enzyme, only active forms and not the inactive pro-enzyme or inhibited enzymes are addressed. It has been estimated that fluorescence scanning can detect approximately 0.1 fmol of fluorophosphonate–rhodamine labeled enzyme (Patricelli et al., 2001). Similarly, cysteine proteases and proteasome units are targeted with epoxy succinyl moieties, vinyl sulfones, acyloxymethyl ketones, acrylates and fluoromethyl ketones (Table 2). All compounds mentioned in this context are potent electrophiles that smoothly react with thiol groups even under physiological conditions. Additional selectivity for the ABP can be obtained by the use of peptide or peptide-like linkers, e.g. for certain classes of proteases. Acyloxymethylketones (Table 2) with an aspartate residue neighbor-

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ing the probe head are selective probes for caspases (Thornberry et al., 1994), whereas Nazif and Bogyo (2001) used a library of peptide vinyl sulfones (Table 2) with varying amino acids in the linker to profile the substrate and inhibitor selectivity of the catalytic subunits of the proteasome. The synthesis of different peptide vinyl sulfones on solid support allowing for rapid optimization of the linker moiety was first developed by Overkleeft et al. (2000). They introduced the vinyl sulfone as a modified amino acid in the last step, thereby inducing cleavage from the solid support. Nazif and Bogyo (2001) started with a vinyl sulfone-containing aspartic acid attached on Rink resin via its side-chain carboxylic acid, allowing positional scanning of the following amino acids, but holding the first position constant. Therefore, their method is limited to peptides with aspartate or glutamate at the first position. Wang et al. (2003b) reported a strategy that allows any amino acid to be incorporated at the first position: a phenolic vinylsulfone derivative of an amino acid is immobilized on 2-chlorotrityl resin followed by chain elongation and coupling with CyDye (if desired). Subsequent cleavage gives the vinylsulfones. The epoxy succinyl moiety is derived from a naturally occurring general inhibitor for cysteine proteases (E-64, Fig. 6). Modified epoxy succinyl-derived conjugates have been employed as affinity labels for the papain class of cysteine proteases (Greenbaum et al., 2000). For compound DCG-04 (Fig. 6), a general probe for lysosomal cathepsins, extensive structure-activity relationships with respect to the S2 /P2 position (leucine residue) are reported. In contrast, Schaschke et al. (1998) developed a concept for the specific targeting of single members of the enzyme subclass of cathepsins. Combination of the structural and binding characteristics of E-64 and CA074 (Fig. 6) gave the highly potent and selective chimeric inhibitor NS-134, whose affinity to cathepsin ˇ B was additionally proven by an X-ray structure (Stern et al., 2004). Addition of a reporter group made this chimeric inhibitor suitable for ABPP. The new ‘double-sided modification’ approach of engineering a substrate-like binding moiety addressing the S’ subsite, as well as modifying the propeptidelike part of the inhibitor interacting with the S subsite, generates a platform for selective targeting of specific cathepsins and may be suited to specifically address

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Fig. 6. Epoxysuccinyl-type ABPs derived from naturally occurring protease inhibitors E-64 and CA074.

other cathepsins, e.g. cathepsin X (Schaschke et al., 2000). Halogenomethyl ketones (Table 2) are also potent electrophiles that readily react with thiol nucleophiles. In this context fluoromethyl ketones have been incorporated in an activity-based probe for cysteine proteases like caspases (Liau et al., 2003). Despite successful efforts to map the substrate specificity of caspases many currently available activity-based probes for this protein family are limited by a high level of background labeling when applied to crude proteomes. The acyloxymethyl ketone electrophile (Table 2) is well-known to address cysteine proteases (Verhelst and Bogyo, 2005). This type of electrophile is characterized by low reactivity towards weak nucleophiles but efficiently addresses thiol residues in the active site. A number of probes containing either a single amino acid or an extended peptide sequence have been shown

to target caspases, legumains, gingipains and cathepsins. However, not all caspases were addressed in a similar manner because of the specific subsite requirements (Kato et al., 2005). Acrylates and among this subclass of compounds especially vinylogous amino acids (Table 2) are also suitable electrophiles for targeting cysteine proteases (Winssinger et al., 2002). In a similar fashion vinyl sulfones may be utilized which has been shown for cysteine proteases and the proteasome (Wang et al., 2003b; Bogyo et al., 2000; Nazif and Bogyo, 2001; Bogyo et al., 1998). Two different types of ABPs for proteine tyrosine phosphatases (PTPs) have been reported so far. Kumar et al. (2004) developed a probe based on ␣-bromo benzyl phosphonate (Table 2) that presumably reacts with the active site Cys residue. On the other hand, different phosphatase suicide inhibitors have been employed to construct activity-

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Fig. 7. Suicide substrates.

based probes, among them probes containing a mimetic of tyrosine phosphate. Besides lacking a methylene group, the amino group was replaced by a fluorine substituent as a leaving group (Fig. 7). Phosphatase action basically relies on the nucleophilic attack of a thiolate group present in a cysteine residue in the active site onto the phosphate ester. In addition to the compounds mentioned, appropriately substituted O-phosphorylated o-difluoromethylphenyl derivatives (Zhu et al., 2003) have been proven to be useful as well (Fig. 7). Dephosphorylation (enzymatic hydrolysis) triggers a 1,6-elimination of HF yielding the corresponding quinone methide that readily reacts with surrounding nucleophiles (Fig. 7; Zhu et al., 2003). Nearly the same concept has been developed independently by Lo et al. (2002) and differs basically only in the alternative application of a fluorophore or a biotin residue as reporter group (Fig. 7). A major drawback of this type of suicide inhibitors generating potent electrophiles in the course of the enzyme reaction is that the conjugates have to be employed in a rather high concentration and that obviously the reaction of the quinone methide with nucleophiles is rather slow, which might allow for diffusion of the reactive species out of the active site. Moreover, application in complex proteomes remains to be successfully proven.

The basic principle has also been employed for the design and synthesis of activity probes for glycosidases and different proteases. In the first case glycosidases cleave the O-glycosyl bond of an aryl glycoside (Fig. 7; Tsai et al., 2002), while proteases may be targeted with a linker construct equipped with a moiety that readily undergoes quinone imine methide formation starting from a peptide derivatized at the C-terminus with the corresponding arene moiety (Fig. 7). The selectivity of the cleavage reaction and, hence, the target protease basically is defined by the P1 or P2 subsite residues. In such a case not the cleavage mechanism itself is of importance as the side chains in P1 or P2 position or more positions of the peptidyl residue define specificity (Zhu et al., 2004). 2.3.2.2. Reversible inhibitors/ligands. Inhibitors or ligands binding irreversibly to their target are not known for many enzymes, especially for those that do not employ nucleophilic catalytic residues. More generally, the same situation applies to non-catalytic proteins where irreversibly binding ligands are scarcely known. As mentioned in the introduction, the denaturing conditions of 2D electrophoresis require formation of a covalent bond between the affinity based probe (ABP) and its target.

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This situation calls for the design, synthesis and functional characterization of novel engineered synthetic molecular tools based on reversible inhibitors/ligands that are able to address selectively the protein families in question. The additional chemical reaction that is needed for covalent bond formation can be performed, e.g. photochemically. The principle of photoaffinity labeling has frequently been exploited, e.g. for localization of the ligand binding site in biochemistry. For this purpose, photoreactive groups have been incorporated into the ABP. The different photoreactive groups and their particular applicational (dis-)advantages have been extensively reviewed (Fleming, 1995; Dorman and Prestwich, 2000; Hatanaka and Sadakane, 2002). First attempts following this strategy have been conducted by Li et al. (2000), who used a reversible ␥-secretase inhibitor equipped with a benzophenone moiety to identify within the ␥-secretase complex the protein containing the active site. Although false positive results cannot be excluded, this method is able to reduce the number of proteins in a complex proteome. Incubation of a proteome with such a photoreactive probe and subsequent photocrosslinking will result in the retrieval of proteins able to bind the recognition unit of the molecular tool. Thereby, a discrete proteome subset is labeled with high sensitivity even in a complex protein mixture. Direct labeling of kinases (Hagenstein et al., 2003) with a fluorophore succeeded with a conjugate comprising a general kinase inhibitor (e.g. the isoquinoline sulfone amide H9), a photoreactive group (pbenzoylphenylalanine, Bpa), and a fluorescent label (e.g. carboxyfluorescein). The inhibitor H9 recognizes the ATP-binding site, forming a non-covalent complex with the target kinase. Upon irradiation at 350 nm, photochemical triplet insertion into CH-bonds occurs, resulting in covalent attachment of the conjugate to the target protein. It has been shown that the native folded form of the protein is required for recognition by the protein ligand and, hence, efficient photoaffinity labeling. Practically no photocrosslinking occurs with denatured kinases. The strategy described has been validated subsequently for metalloproteases by different other groups (Saghatelian et al., 2004; Chan et al., 2004; Jenssen, 2004). They conjugated zinc-chelating hydroxamate inhibitors of metalloproteases to a photoreac-

tive group and different reporters. Benzophenone has been attached at different positions by Saghatelian et al. (2004) and Jenssen (2004), while Chan et al. (2004) used a diazirine as the photoreactive group (Table 3). Recently, Chattopadhaya et al. (2005) reported the successful extension of this strategy towards aspartic proteases (Table 3). Analogously, tagging of carbohydrate binding proteins has also been shown to be feasible via this approach. Ballell et al. (2005) developed a probe for lectins comprising a sugar moiety as ligand, benzophenone as photoreactive group and an azide for attachment of a reporter group (an alkyne containing rhodamine derivative) via click chemistry (Table 3). A possible drawback of the photoreactive groups is that they do not necessarily react in the active site, hence identification of catalytic residues is not possible. Reversible inhibitors allow the generation of enzyme-family sub-proteomes via inhibitor affinity chromatography (IAC) as well. Generation of a proteome subset by IAC succeeds by immobilization of small organic compounds (e.g. engineered inhibitors) on an affinity chromatography column. This has been successfully proven by the creation of a metalloprotease sub-proteome (Freije and Bischoff, 2003) or a kinase sub-proteome (Lolli et al., 2003; Daub et al., 2004; Wissing et al., 2004; Brehmer et al., 2004; Brehmer et al., 2005). For the optimization process of the elution conditions in the IAC process surface plasmon resonance (SPR) represents an excellent method (Jenssen et al., 2004). 2.4. Proteomics technologies To date, ABPP has mostly relied on gel-based methods for proteome analysis, putting up with the rather tedious, labor intensive and time-consuming protocol of gel electrophoresis followed by gel excision and tryptic digestion prior to protein identification via LC–MS. Recently, several approaches have been reported that avoid SDS-PAGE. Adam et al. (2004) developed a non-directed approach for the mapping of enzyme active sites in complex proteomes (see above) that basically relies on affinity capture of labeled peptides with immobilized anti-rhodamine antibodies after

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Table 3 ABPs based on reversible inhibitors/ligands Enzyme targets

Structure of the probe (reference)

Kinases

Metalloproteases

Aspartic proteases

Galectins

ABP-incubation and tryptic digestion, followed by LC–MS/MS analysis. A comparable approach with directed ABPs, the socalled “Xsite-platform”, was reported by Okerberg et al. (2005). Proteomes were labeled with fluorescent fluorophosphonate probes, followed by gel filtration and tryptic digestion. The ABP-modified peptides can then

either be rapidly profiled using capillary electrophoresis with laser-induced fluorescence detection (CE-LIF), whilst affinity-capture with anti-fluorophore antibodies on agarose beads followed by LC–MS/MS allows identification of the labeled proteins. Both approaches reveal the identity of the protein targets as well as the precise site of labeling.

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A tool comparable to the high-throughput DNAmicroarray/chip technology in genomics has been missing in proteomics for a long time. Within the last few years, protein microarrays have gained attention due to their high-throughput capability (LaBaer and Ramachandran, 2005), but only few applications have been reported for ABPP. First attempts were conducted by Winssinger et al. (2002). They tethered acrylate moieties to fluorescein-capped peptide nucleic acid (PNA) tags. Inhibitor-protein adducts are hybridized to an oligonucleotide microarray and quantified according to fluorescence intensity. Identification of labeled proteins is achieved by tryptic digestion followed by nanoflow RP-HPLC/micro electrospray ionization (␮ESI)/MS. Since the probes serve as labeling and capture reagents, ABPs with high specificity for individual enzymes are mandatory, thereby to some extent limiting the applicability of this approach. Sieber et al. (2004) developed a more general microarray platform for ABPP, based on orthogonal strategies for labeling and capture of target proteins. Proteomes are first labeled with broad-specificity fluorescent ABPs, e.g. fluorophosphonate-rhodamine. Subsequently, enzymes of interest are captured on glass slides coated with anti-enzyme antibodies. The relative abundance of active enzymes is then determined in a fluorescence scanner. Arraying complementary sets of antibodies allows profiling of several enzyme activities in parallel with enhanced sensitivity and resolution compared to gel-based profiling. Isolation, detection and identification are consolidated in one step. Even though this approach does not require highly selective ABPs, the applicability is somewhat limited due to the dependence on high-quality antibodies to prevent cross-reactivity and false-positives, which are currently available only for a small percentage of the proteome. In contrast, Funeriu et al. (2005) chose enzyme microarrays for activity-based screening of enzyme families. They blotted members of the cathepsin family onto an aldehyde-treated glass slide and treated them with a family-wide activity-based epoxide-containing fluorescent affinity label (FAL), a Cy3-comprising DCG-04 derivative (Fig. 6). Thereby, they could determine the kinetic constants and the reaction mechanism of the FAL. A high throughput screening (HTS) setup allowed quantitative, multiplexed determination of the

inhibition profile against each microarrayed enzyme for members from an inhibitor library. The need for pure protein samples and the uncertainty of whether the proteins will retain integrity and activity under microarray conditions represent drawbacks of this method.

3. Conclusion and outlook The past years have experienced a tremendous increase in activity by chemists in the areas of proteomics and genomics. A broad variety of molecular tools for applications in proteomics has been designed, synthesised and characterised with respect to function. In this context, especially affinity based probes have been proven to represent useful molecular tools for the purification and identification of enzyme targets but also for activity profiling of enzyme activities. However, not only enzymes can now be targeted with the latest generation of affinity based probes, those which are able to be covalently attached to the protein by, e.g. photoaffinity labeling. The scope of targets has been extended from enzymes to the full proteome by using protein ligands instead of enzyme inhibitors. Alternatively, the utilization of non-directed ABP has been proven a viable approach. Several important classes of proteins can be tagged in a family specific manner. Besides the different classes of proteases, sub-proteomes comprising kinases or phosphatases served for the validation of affinity based probes as molecular tools in proteome research. While a lot of work remains to be done in the application of the affinity based probes that have been published until now, new probes with new ligands and new chemistry have to be developed. Chemical proteomics will expand its activities in the field of interaction analysis (interactomics) and in the field of chemistry in living systems where small fluorescent molecules will be used to study the interaction and trafficking of proteins or, more generally, biomolecules in a cell. All these activities perfectly fit into the new challenge of the next decade: molecular systems biology. It aims towards the quantitative understanding of complex controlled biological processes on a cellular level, which requires a deep knowledge of the complex relationship between molecular structure, molecular pro-

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cesses, their mechanisms and functional information of biological networks. General principles behind biological function will be elucidated on the basis of the interactions between components of living systems. Without any doubt this tremendous challenge can only be tackled when scientists from different disciplines efficiently collaborate with each other.

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