CHAPTER TWO
Photoreactive Stapled Peptides to Identify and Characterize BCL-2 Family Interaction Sites by Mass Spectrometry Susan Lee*,†, Craig R. Braun*,†,{, Gregory H. Bird*,†, Loren D. Walensky*,†,1
*Department of Pediatric Oncology and the Linde Program in Cancer Chemical Biology, Dana-Farber Cancer Institute, Boston, Massachusetts, USA † Department of Pediatrics, Children’s Hospital Boston, Harvard Medical School, Boston, Massachusetts, USA { Department of Cell Biology, Harvard Medical School, Boston, Massachusetts, USA 1 Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. Overview of Photoreactive Stabilized Alpha-Helices Methodology 3. Design and Synthesis of Photoreactive Stapled Peptides 4. Photoaffinity Labeling and Retrieval 5. Mass Spectrometry Analysis 6. Computational Docking 7. Mapping Novel Binding Sites 8. Conclusions and Future Directions Acknowledgments References
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Abstract Protein interactions dictate a myriad of cellular activities that maintain health or cause disease. Dissecting these binding partnerships, and especially their sites of interaction, fuels the discovery of signaling pathways, disease mechanisms, and next-generation therapeutics. We previously applied all-hydrocarbon peptide stapling to chemically restore a-helical shape to bioactive motifs that become unfolded when taken out of context from native signaling proteins. For example, we developed stabilized alphahelices of BCL-2 domains (SAHBs) to dissect and target protein interactions of the BCL-2 family, a critical network that regulates the apoptotic pathway. SAHBs are a-helical surrogates that bind both stable and transient physiologic interactors and have effectively uncovered novel sites of BCL-2 family protein interaction. To leverage stapled peptides for proteomic discovery, we describe our conversion of SAHBs into photoreactive agents that irreversibly capture their protein targets and facilitate rapid
Methods in Enzymology, Volume 544 ISSN 0076-6879 http://dx.doi.org/10.1016/B978-0-12-417158-9.00002-9
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2014 Elsevier Inc. All rights reserved.
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identification of the peptide helix binding sites. We envision that the development of photoreactive stapled peptides will accelerate the discovery of novel and unanticipated protein interactions and how they impact health and disease.
1. INTRODUCTION Alpha-helical interactions are found throughout the cell and govern critical biological processes, such as infection (Weissenhorn, Dessen, Harrison, Skehel, & Wiley, 1997), the immune response (Blum, Stevens, & DeFranco, 1993), apoptosis (Sattler et al., 1997), and transcription (Kussie et al., 1996; Fig. 2.1). The natural complexity of peptide a-helices enables them to engage diverse cellular targets with high affinity and selectivity. Indeed, helical peptides can effectively discriminate among homologous proteins owing to the high fidelity key-in-lock specificity afforded by their amino acid composition. The conserved BCL-2 homology 3 (BH3) domain of BCL-2 family proteins is a critical interaction module that mediates
A. Cell surface
B. Plasma membrane
C. Cytosol and mitochondrion
D. Nucleus
Figure 2.1 The peptide a-helix is a ubiquitous secondary structural motif that mediates a host of biomedically relevant protein interactions. Hydrocarbon-stapled peptides modeled after bioactive helices can be generated to target and modulate protein interactions from the surface to the inner nuclear core of the cell.
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member-specific communication. The canonical complex between a proapoptotic BH3 a-helix and a surface groove on antiapoptotic members reflects the molecular wrestling match between pro- and antiapoptotic signals (Sattler et al., 1997). If antiapoptotic grooves are sufficient in number to bind and sequester the proapoptotic BH3 signals, cell survival prevails. In contrast, if the capacity to withstand proapoptotic assault is breached, cell death ensues (Fig. 2.2). Not only did the discovery of this protein interaction paradigm provide a mechanism for apoptotic regulation, but it also informed the development of drugs to reactivate cell death through targeted inhibition of antiapoptotic BH3-binding pockets (Oltersdorf et al., 2005). Given the critical roles of amphipathic alpha-helices in mediating signal transduction, we advanced all-hydrocarbon stapling to refold bioactive Multi-BH domain antiapoptotic proteins
Multi-BH domain proapoptotic proteins
Heavy cell stress
BAX/BAK activation by BH3-only BAX
Anti-apoptotic inhibition by BH3-only
BH3-only displacement of sequestered BAX/BAK
BH3-only proapoptotic proteins
Weak cell stress
Anti-apoptotic sequestration of BH3-only
Anti-apoptotic sequestration of BAX/BAK
BAX/BAK remain inactive BAX
BAK
BAK
Oligomerization
Cytochrome c release
Cell death
Cell survival
Figure 2.2 BCL-2 family proteins regulate the life and death decision of stressed cells. If mitochondrial antiapoptotic proteins (orange) can effectively harness their C-terminal binding pockets to trap and sequester the BH3-signaling helices of proapoptotic proteins, cell survival prevails. However, with increased cellular stress, proapoptotic signals overwhelm the antiapoptotic reserve. The BH3 domain helices of BH3-only proteins (blue) can directly activate the essential executioner proteins BAX and BAK (gray) and also release trapped forms from antiapoptotic inhibition by targeting the antiapoptotic groove.
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alpha-helical peptides for use as research tools and prototype therapeutics (Fig. 2.3A). Depending on their amino acid composition and design, “stapled peptides” have proven to be structurally stable, protease resistant, and cell permeable agents capable of interrogating and modulating protein interactions in vitro and in vivo (Bernal et al., 2010; Kim et al., 2013; LaBelle et al., 2012; Takada et al., 2012; Walensky et al., 2004). Such stapled peptides have revealed unanticipated and functionally relevant helix/target interactions, including engagement of (1) antiapoptotic MCL-1 by its own MCL-1 BH3 helix (Stewart et al., 2010; Fig. 2.3B), (2) a novel “trigger site” on proapoptotic BAX by the BIM BH3 helix (Gavathiotis et al., 2008; Fig. 2.3C), and (3) glucokinase by a BH3-phosphorylated form of the BH3-only protein BAD (Danial et al., 2008). The capacity of these stabilized alpha-helices of BCL-2 domains (SAHBs) to access canonical and noncanonical protein interactors, and reveal sites of interaction by their use in NMR and X-ray crystallography studies, prompted us to consider whether they could be harnessed for higher throughput binding site discovery. Here, we describe our approach to converting stapled peptides into high fidelity photoaffinity reagents and the methodologies developed to rapidly identify their sites of target protein interaction.
2. OVERVIEW OF PHOTOREACTIVE STABILIZED ALPHA-HELICES METHODOLOGY We generate photoreactive stabilized alpha-helices or pSAHs by substituting into the peptide sequence a nonnatural amino acid bearing a benzophenone moiety (4-benzoylphenylalanine, Bpa) (Fig. 2.4A), which covalently crosslinks to protein targets upon exposure to ultraviolet (UV) light (Dorman & Prestwich, 1994; Saghatelian, Jessani, Joseph, Humphrey, & Cravatt, 2004; Vodovozova, 2007). We then insert an (i, i + 4) or (i, i + 7) pair of nonnatural amino acids bearing olefin tethers, followed by ruthenium-catalyzed ring-closing metathesis (RCM), to generate a hydrocarbon-stapled peptide with reinforced a-helical structure (Blackwell et al., 2001; Schafmeister, Po, & Verdine, 2000; Fig. 2.4B). For binding site identification, pSAHs containing alternatively placed Bpa moieties are individually incubated with a protein-of-interest and UV irradiated (Fig. 2.4C). The mixture is subjected to electrophoresis and the crosslinked species excised, trypsinized, and prepared for mass spectrometry analysis, which is designed to identify peptidic fragments bearing Bpa crosslinks and thus the explicit sites of target protein intercalation (Fig. 2.4D).
A
Insert nonnatural amino acids Cy3P CI
Ph
Ru
Chemically “staple” the peptide back into shape
O NH
O O
*
OH CH3
CI Cy3P
[ ]n
Unstructured peptide - Loss of functional shape - Easily degraded - Does not enter cells Hydrophobic Positive charge
Stapled peptide - Stabilized structure - Not degraded - Enters cells Hydrophilic Negative charge
B
C R214 D218
MCL-1 SAHBD BIM BH3 K21 R263 N260 D256
S255 Q32
Staple
R134 E131
MCL-1Δ NΔ C
BAX
Figure 2.3 Synthetic overview of all-hydrocarbon stapling and its applications in developing a-helical peptides for structural studies of protein interactions. (A) Pairs of a,a-disubstituted nonnatural amino acids bearing olefin tethers are substituted into the peptide sequence at discrete locations (e.g., i, i + 4), followed by rutheniumcatalyzed ring-closing metathesis (RCM) to generate “stapled peptides.” (B) Crystal structure of a stapled MCL-1 BH3 helix in complex with antiapoptotic MCL-1 (Stewart, Fire, Keating, & Walensky, 2010). (C) Calculated model structure of a BIM BH3 helix engaging the N-terminal trigger site of full-length BAX, as derived from paramagnetic relaxation enhancement NMR analyses using full-length 15N-BAX- and MTSL-labeled BIM SAHBs (aa 145–164) (Gavathiotis et al., 2008).
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A
B
Cy3P
O O
NH O
CI *
OH CH3
Ph
Ru CI
Cy3P
[ ]n
O
UV light
Trypsinolysis
D
O
O
C
O
O
E
F
Figure 2.4 Design of photoreactive stabilized alpha-helices (pSAHs) for binding site identification by mass spectrometry. A photoreactive Bpa residue (A) and a pair of stapling amino acids (B) are inserted into the peptide template followed by RCM to generate a pSAH. Upon exposure to UV light, the bound pSAH covalently crosslinks to the target protein (C) and, following electrophoresis of the mixture, crosslinked protein is excised from the gel and subjected to in-gel digestion with trypsin (D). LC–MS/MS analysis identifies the explicit sites of covalent modification, which when mapped on to the protein structure reveal the region of pSAH interaction (E). Top scoring crosslinks from a series of experiments employing sequentially placed Bpa residues can provide interaction restraints for calculating model structures (F).
By mapping the crosslink sites of pSAHs bearing alternatively placed Bpa moieties onto the protein structure (if known), the location of the interaction site can be determined (Fig. 2.4E). Furthermore, if the structure of the protein target has been solved and a series of pSAHs with sequentially localized Bpa residues map sites of intercalation with sufficient resolution,
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computational docking can be applied to calculate a model structure of the complex for biochemical and functional validation (Fig. 2.4F).
3. DESIGN AND SYNTHESIS OF PHOTOREACTIVE STAPLED PEPTIDES The design of photoreactive stapled peptides is ideally based on a structure involving the template a-helix, so that the structurally reinforcing staples can be placed on the noninteracting surface and Bpa residues located at or near the putative interaction surface to facilitate UV crosslinking. Ideally, Bpa residues are substituted at positions of homologous, bulky, and hydrophobic residues (e.g., Phe) to minimize the effect of Bpa on the binding interface. Without structural information, a library of constructs can be generated by sampling alternative staple and Bpa positions along the length of the peptide template, selecting those constructs with successful a-helical induction (as determined by circular dichroism analysis; Bird, Bernal, Pitter, & Walensky, 2008) and preserved binding activity for experimental application. Of note, to simplify interpretation of the MS2-based identification of Bpa-crosslinked peptides, it is also preferable to locate Bpa residues in regions of the peptide such that trypsinization will yield short cleavage products. If necessary, a lysine or arginine residue can be strategically substituted into the peptide, ideally on the noninteracting surface or as a conserved substitution observed in other species (Leshchiner, Braun, Bird, & Walensky, 2013), to accomplish this goal. Once designed, stapled peptides are synthesized as previously described in detail (Bird et al., 2008; Bird, Crannell, & Walensky, 2011) using Fmoc chemistry, stapling amino acids installed at (i, i + 4) or (i, i + 7) positions (to bridge one or two turns of the a-helix, respectively), and ruthenium-catalyzed olefin metathesis with Grubbs generation I catalyst (Fig. 2.5). Of note, methionine residues in the template sequence are typically replaced by norleucines, as the unprotected sulfur can decrease the efficiency of the Grubbs catalyst. Following peptide deprotection and cleavage, pSAHs are purified by reverse phase high performance liquid chromatography–mass spectrometry and quantified by amino acid analysis (AAA). Pure peptide is stored as a lyophilized powder at 20 C until use. In addition to the experimental conditions summarized in Fig. 2.5 and the step-by-step synthetic protocol with reagent list detailed in our recently updated methodologic review (Bird et al., 2011), our approach to stapled peptide synthesis incorporates the following general principles:
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R Fmoc
1. 20% piperidine in NMP
H N
N H
HCTU/DIEA in NMP
O
n
O
H
2.
Fmoc N
OH Bp
1. 20% piperidine in NMP HCTU/DIEA in NMP
2. Fmoc
H N
O
O
(S)
OH
H Fmoc N
R N H
Bp O
H N
Fmoc
R N H
R
O
H N
N H
O
HCTU/DIEA in NMP
N H
1. 20% piperidine in NMP
O
R N H
O
HCTU/DIEA in NMP
R
2.
H N
n
O
1. 20% piperidine in NMP
Fmoc
H N O
3
n
R
N H
O
n⬘
H N
R
O
H N
Fmoc
Fmoc
OH
N H
OH
N H
O
O O
R
H N
Fmoc
O
N H
H N O
n⬙
n
3
R
2.
R N H
N H
O
Bp
R
O
H N
H N
O
O
n⬘
R N H
H N O
n
3
1. 20% piperidine in NMP 2. Fmoc
H N
O (S)
1. 20% piperidine in NMP
OH
HCTU/DIEA in NMP
Fmoc
O
H N
2. Ac2O, DIEA, NMP
R N H
O
H N
R
H N
N H
O
n
3
O
R Ac
O
N H
H N O
R N H
O
H N O
Bp
R N H
O
R N H
O
n⬘
n⬙
H N
H N O
n
3
1. 20% piperidine in NMP 2.
HCTU/DIEA in NMP
1.
R Fmoc
PCy3
Cl
OH
N H
, DCE
Ru Cl
O
PCy3
2. TFA:TIS:H2O (95:2.5:2.5) R Fmoc
N H
H N O
n⬘
R
O N H
H N O
3
R
O N H
H N O
n
O
R Ac
N H
H N O
n⬙
R N H
Bp
H N O
n⬘
R
O N H
H N O
O
R N H
NH 2 O
n
Figure 2.5 Synthetic steps for the automated production of pSAHs by Fmoc-based solid-phase peptide synthesis and ruthenium-catalyzed RCM. Alternatives to N-terminal acetylation include capping with FITC or biotin, depending on the desired application. Bp, 4-benzoylphenyl.
a. Create an automated synthesis program by employing standard double Fmoc deprotections, vigorous washes with DMF, and subsequent single 10-fold excess or double fivefold excess amino acid couplings. The precise timing and coupling excesses are optimized based on the specific
Photoreactive Stapled Peptides
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coupling reagents, bases, and synthesizer employed. Particular attention is devoted to reactions involving the a,a-disubstituted amino acids, as the amine is linked to a quaternary carbon center, and therefore, can be recalcitrant to Fmoc removal and the subsequent acylation steps. For example, couplings that involve beta-branched amino acids and arginine can be especially challenging. Typical protocol modifications include multiple extended deprotections and acylations, and sometimes capping at these difficult steps with acetic acid (AcOH) or even decanoic acid in order to capture and terminate unreacted amine. In this manner, the production of full-length peptide species can be maximized, while also avoiding the challenge of chromatographic separation of truncated byproducts. b. Prepare the peptide synthesis reagents and solutions by weighing out the resin and dissolving the calculated amount of each amino acid in anhydrous NMP. Although more expensive than DMF, NMP has greater stability and we have seen no deterioration in peptide quality using reagent solutions for up to 2 weeks. A secondary amine is used to remove the Fmoc protective group; appropriate choices include piperidine, piperazine, or 1,8-diazabicycloundec-7-ene. Likewise, there are multiple coupling reagents to select from. HCTU/DIEA and HOBt (or HOAt)/DIC nicely balance reactivity with solution stability, and more reactive coupling reagents range from uronium based (HATU) to the more exotic tris-pyrrolidinophosphonium derivatives. c. Evaluate synthetic success by subjecting a small sample of beads to standard trifluoroacetic acid (TFA) cleavage (95:2.5:2.5 TFA:water: triisopropylsilane) and analysis by LC/MS. If the full-length species is identified in good yield, the next step is acylation with acetic anhydride or Fmoc-b-alanine, followed by stapling with Grubbs I catalyst (bis (tricyclohexylphosphine) benzylidine ruthenium (IV) dichloride) dissolved in dichloroethane. For most sequences and staple locations, a 2 2 h incubation using several milliliters of a 4 mg/mL solution will yield a completely metathesized product. If there is incomplete metathesis, the reaction can be repeated multiple times and with longer incubation periods. d. After confirmation of successful stapling, the Fmoc group of Fmocb-alanine is removed and the N-terminus derivatized with biotin or fluorescein isothiocyanate (FITC), depending on the application (see later). The final product is cleaved off the resin in large scale, precipitated with ether:hexanes, and purified by HPLC or, if available, by LC/MS with mass-triggered fraction collection. The pure fractions are pooled,
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lyophilized, and peptide powder resuspended in a known volume, followed by submission of several dilutions for AAA. Although we typically store stapled peptides as dried powder (see above), the material can also be solubilized in 100% DMSO for storage at 20 C.
4. PHOTOAFFINITY LABELING AND RETRIEVAL With a panel of photoreactive stapled peptides in hand, the next step is to isolate pure protein-of-interest for photoaffinity labeling. We have typically expressed glutathione S-transferase, chitin-binding domain, or histidine-tagged fusion proteins followed by affinity purification, cleavage of the tag, and size exclusion chromatography. Protein purity and identity are confirmed by SDS-PAGE, protein staining (e.g., coomassie, silver), protein-specific western blot, and mass spectrometry analyses. Protein concentrations are determined using the Bradford assay (Biorad). Photoreactive stapled peptide and pure recombinant protein (e.g., 10 mM) are typically mixed at a 1:1 ratio, but higher pSAH concentrations can be used if necessary to increase the quantity of crosslinked product. The peptide/protein mixture is vortexed, incubated for 20 min in the dark, and then irradiated (365 nm) on ice for 1.5 h by use of a Spectroline handheld UV lamp (Model En280L, Spectronics Corporation). The duration of UV exposure can also be adjusted based on the efficiency of crosslinking for a particular peptide/protein complex. The mixtures are then diluted with 4 LDS loading buffer (Invitrogen), electrophoresed (e.g., 4–12% Bis–Tris gels [Invitrogen]), and analyzed by coomassie staining (SimplyBlue Safestain, Invitrogen). The appearance of a slower migrating band just above the protein starting material reflects the pSAH-crosslinked species, which is excised for MS analysis (see below). Several supplementary experiments can be especially useful when piloting this approach. For example, to confirm that insertion of staples and Bpa residues at discrete sites within the peptide template do not disrupt target engagement, we recommend generating fluorescently labeled (FITC) pSAH derivatives for comparative fluorescence polarization (FP) binding analyses with a positive control ligand. In the case of FITC-BAD pSAHBs, the substitution of Bpa residues at four positions within the BAD BH3 sequence had no disruptive effect on binding to the antiapoptotic protein BCL-XLDC (Braun et al., 2010; Fig. 2.6A). The FITC-derivatized pSAHs can also be used for photoaffinity labeling, with conversion of recombinant protein into a fluorescent species that can readily be detected by fluorescence
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Photoreactive Stapled Peptides
imaging (Gel Doc XR Molecular Imager and Quantity One software, BioRad), providing further confirmation of effective crosslinking (Braun et al., 2010; Fig. 2.6B). Biotinylated pSAHs can also be used for high sensitivity screening of effective crosslinking, with electrophoresed protein analyzed by anti-biotin western blot (see below). Binding specificity experiments using bovine serum albumin (BSA), for example, are also advised. Crosslinking is performed as above, except that BSA is added in molar excess Coomassie
Compound
BAD SAHB BAD pSAHB-1 BAD pSAHB-2 BAD pSAHB-3 BAD pSAHB-4
C
60.5 (51.9–70.6) 46.7 (44.2–49.4) 38.9 (35.9–42.1) 38.5 (36.5–40.6) 55.6 (53.1–58.1)
BCL-XLΔC Fluorescence BSA Crosslinked BCL-XLΔC BCL-XLΔC
100
60
0 10 10
D
20
10–9
10–8
10–7
10–6
10–5
[BCL-XLΔC], M
0 0.5 1
Fluorescence
Ve hi BA cle D BA pS D AH BA pS B-1 D AH BA pS B-2 D AH p B Ve SA -3 H hi BA cle B-4 D p BA S D AH BA pS B-1 D AH BA pS B-2 D AH pS BAH 3 B4
Coomassie
10 10 10
30 10 10
Fluorescence
Crosslinked BCL-XLΔC BCL-XLΔC
B
.5 10 10
Coomassie
Crosslinked BCL-XLΔC BCL-XLΔC
40
Crosslinked BCL-XLΔC BCL-XLΔC
10 0 0
BSA (mM) BCL-XLΔC (mM) BAD pSAHB-2 (mM)
80
0 10–10
BSA Crosslinked BCL-XLΔC
BAD pSAHB-2 crosslinking yield (%)
Normalized polarization (%)
A
EC50 95% CI
2
5 10 20 30 40 50 60 80 100 120 Time (min)
40
30
20
10
0 0
20
40
60
80
100
120
Time (min)
Figure 2.6 pSAHs bind and selectively crosslink to their protein targets, as exemplified by the photocrosslinking of BAD pSAHBs to antiapoptotic BCL-XLDC (Braun et al., 2010). (A) A series of FITC-BAD pSAHBs retain nanomolar binding affinity to BCL-XLDC as measured by FP analysis. (B) FITC-BAD pSAHBs exhibit a spectrum of BCL-XLDC-crosslinking efficiency upon UV exposure, with BAD pSAHB-2 demonstrating the greatest reactivity toward BCL-XLDC (left, coomassie stain; right, fluorescence scan). (C) The selectivity of BAD pSAHB-2 is reflected by the absence of crosslinking to BSA and the lack of effect of added BSA on the BCL-XLDC-crosslinking efficiency of FITC-BAD pSAHB-2. (D) Time course for photocrosslinking of FITC-BAD pSAHB-2 to BCL-XLDC as monitored by coomassie stain (top) and fluorescence scan (middle). A plot of crosslinking yield, calculated based on densitometry, demonstrates the time-dependent production of crosslinked BCL-XLDC (bottom).
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prior to addition of pSAH. In the case of FITC-BAD pSAHB-2/BCL-XLDC crosslinking, the addition of molar excess BSA had no effect on the yield of pSAHB-crosslinked BCL-XLDC (Braun et al., 2010; Fig. 2.6C). Tracking the production of pSAH-crosslinked protein over time is useful for determining just how long to extend the reaction time to produce maximal reaction product for the MS analyses (Fig. 2.6D). If the efficiency of pSAH crosslinking is suboptimal after increasing the peptide/protein ratio and extending the reaction time (i.e., crosslinked species not readily visualized by coomassie stain), the capacity to load sufficient material for electrophoresis and band excision can be compromised. Although we typically find that sampling a variety of pSAH constructs reveals lead photoaffinity labeling reagents (Fig. 2.6B), an alternative approach to enrich for crosslinked species entails the synthesis and application of pSAHs capped at the N-terminus with biotin. Photoaffinity labeling with biotinylated pSAHs is performed as above and then unreacted peptide is removed from the irradiated samples by overnight dialysis at 4 C in 50 mM Tris pH 7.4, 200 mM NaCl buffer using 6–8 kDa molecular weight cut-off D-Tube dialyzers (EMD Biosciences). After addition of SDS to a final concentration of 0.2%, biotinylated protein is isolated from unreacted protein by incubation with high capacity streptavidin agarose (50 mL 50% slurry/reaction) for 2 h at room temperature. The streptavidin beads are successively washed at room temperature in 1% SDS in PBS, 1 M NaCl in PBS, and then 10% ethanol in PBS for 3 10 min each. Biotinylated proteins are eluted by boiling for 30 min in a 10% SDS solution (Promega) containing biotin (10 mg/mL), electrophoresed (e.g., 4–12% Bis–Tris gels [Invitrogen]), and then subjected to coomassie staining. Using this enrichment protocol for purifying biotinylated pSAH-crosslinked protein, isolating sufficient material for MS analyses of even low efficiency crosslinked complexes (e.g., weak or transient protein interactions) can typically be achieved (Edwards et al., 2013; Leshchiner et al., 2013).
5. MASS SPECTROMETRY ANALYSIS To prepare pSAH-crosslinked protein for MS analysis, we perform an in-gel digest of the excised protein band using a standard protocol (Braun et al., 2010), mass spectrometry grade reagents, and due caution to avoid keratin contamination. Gels are subjected to coomassie or other MS-compatible stain (e.g., Pierce Silver Stain) and the excised gel slab cut into 1 mm cubes for processing as follows:
Photoreactive Stapled Peptides
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a. Destain: Add 500 mL of 50% acetonitrile (ACN)/50 mM ammonium bicarbonate (AB), pH 8 and incubate at 37 C with shaking for 15 min; remove solution and repeat until the blue color is eliminated from the gel pieces. b. Dehydrate: Add 500 mL of ACN, incubate at room temperature for 5 min, remove solution, and repeat. c. Reduce disulfides: Add 100 mL of 10 mM dithiothreitol in AB, incubate at 56 C for 30 min, and remove solution. d. Alkylate cysteines: Add 100 mL of 50 mM iodoacetamide in AB, incubate at room temperature in the dark for 20 min, and then remove solution. e. Dehydrate: As above. f. Digest: Cover gel pieces with cold 12.5 ng/mL trypsin (Promega) in AB, incubate on ice for 45 min and then overnight at 37 C. Peptides are extracted twice in 50% ACN/5% formic acid (FA) and dried in a speed-vac. Samples are prepared for LC–MS/MS using home-made C18 stage tips or Agilent C18 Omix tips (Braun et al., 2010; Jakobsen, Schroder, Larsen, Lundberg, & Andersen, 2013). The sample preparation procedure below employs 100 mL Omix tips: a. Place 6 mL of elution buffer (80% ACN/0.5% AcOH) in a clean elution tube for each sample. b. Resuspend the dried peptides in 100 mL of 0.1% TFA. c. Wet tips by pipetting 2 100 mL 40% ACN/0.5% AcOH. Be sure to prevent air from being introduced into the tips once wetted. d. Equilibrate tips by pipetting 2 100 mL of 0.1% TFA. e. Bind peptides in sample by carefully pipetting up and down 10 times. f. Wash by pipetting 3 100 mL of 0.1% TFA. g. Elute by carefully pipetting up and down in a prepared elution tube three times. h. Dry the eluate by speed-vac. We use a split flow LC system with in-house packed C4/C18 columns for MS analysis on a Thermo LTQ Orbitrap Discovery mass spectrometer. Samples are reconstituted in 5% ACN/5% FA and loaded onto the LC. We employ 30 cm of 100 mm inner diameter fused silica tips packed with
Photoreactive Stapled Peptides to Identify and Characterize BCL-2 Family Interaction Sites by Mass Spectrometry Susan Lee*,†, Craig R. Braun*,†,{, Gregory H. Bird*,†, Loren D. Walensky*,†,1
*Department of Pediatric Oncology and the Linde Program in Cancer Chemical Biology, Dana-Farber Cancer Institute, Boston, Massachusetts, USA † Department of Pediatrics, Children’s Hospital Boston, Harvard Medical School, Boston, Massachusetts, USA { Department of Cell Biology, Harvard Medical School, Boston, Massachusetts, USA 1 Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. Overview of Photoreactive Stabilized Alpha-Helices Methodology 3. Design and Synthesis of Photoreactive Stapled Peptides 4. Photoaffinity Labeling and Retrieval 5. Mass Spectrometry Analysis 6. Computational Docking 7. Mapping Novel Binding Sites 8. Conclusions and Future Directions Acknowledgments References
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Abstract Protein interactions dictate a myriad of cellular activities that maintain health or cause disease. Dissecting these binding partnerships, and especially their sites of interaction, fuels the discovery of signaling pathways, disease mechanisms, and next-generation therapeutics. We previously applied all-hydrocarbon peptide stapling to chemically restore a-helical shape to bioactive motifs that become unfolded when taken out of context from native signaling proteins. For example, we developed stabilized alphahelices of BCL-2 domains (SAHBs) to dissect and target protein interactions of the BCL-2 family, a critical network that regulates the apoptotic pathway. SAHBs are a-helical surrogates that bind both stable and transient physiologic interactors and have effectively uncovered novel sites of BCL-2 family protein interaction. To leverage stapled peptides for proteomic discovery, we describe our conversion of SAHBs into photoreactive agents that irreversibly capture their protein targets and facilitate rapid
Methods in Enzymology, Volume 544 ISSN 0076-6879 http://dx.doi.org/10.1016/B978-0-12-417158-9.00002-9
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2014 Elsevier Inc. All rights reserved.
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identification of the peptide helix binding sites. We envision that the development of photoreactive stapled peptides will accelerate the discovery of novel and unanticipated protein interactions and how they impact health and disease.
1. INTRODUCTION Alpha-helical interactions are found throughout the cell and govern critical biological processes, such as infection (Weissenhorn, Dessen, Harrison, Skehel, & Wiley, 1997), the immune response (Blum, Stevens, & DeFranco, 1993), apoptosis (Sattler et al., 1997), and transcription (Kussie et al., 1996; Fig. 2.1). The natural complexity of peptide a-helices enables them to engage diverse cellular targets with high affinity and selectivity. Indeed, helical peptides can effectively discriminate among homologous proteins owing to the high fidelity key-in-lock specificity afforded by their amino acid composition. The conserved BCL-2 homology 3 (BH3) domain of BCL-2 family proteins is a critical interaction module that mediates
A. Cell surface
B. Plasma membrane
C. Cytosol and mitochondrion
D. Nucleus
Figure 2.1 The peptide a-helix is a ubiquitous secondary structural motif that mediates a host of biomedically relevant protein interactions. Hydrocarbon-stapled peptides modeled after bioactive helices can be generated to target and modulate protein interactions from the surface to the inner nuclear core of the cell.
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member-specific communication. The canonical complex between a proapoptotic BH3 a-helix and a surface groove on antiapoptotic members reflects the molecular wrestling match between pro- and antiapoptotic signals (Sattler et al., 1997). If antiapoptotic grooves are sufficient in number to bind and sequester the proapoptotic BH3 signals, cell survival prevails. In contrast, if the capacity to withstand proapoptotic assault is breached, cell death ensues (Fig. 2.2). Not only did the discovery of this protein interaction paradigm provide a mechanism for apoptotic regulation, but it also informed the development of drugs to reactivate cell death through targeted inhibition of antiapoptotic BH3-binding pockets (Oltersdorf et al., 2005). Given the critical roles of amphipathic alpha-helices in mediating signal transduction, we advanced all-hydrocarbon stapling to refold bioactive Multi-BH domain antiapoptotic proteins
Multi-BH domain proapoptotic proteins
Heavy cell stress
BAX/BAK activation by BH3-only BAX
Anti-apoptotic inhibition by BH3-only
BH3-only displacement of sequestered BAX/BAK
BH3-only proapoptotic proteins
Weak cell stress
Anti-apoptotic sequestration of BH3-only
Anti-apoptotic sequestration of BAX/BAK
BAX/BAK remain inactive BAX
BAK
BAK
Oligomerization
Cytochrome c release
Cell death
Cell survival
Figure 2.2 BCL-2 family proteins regulate the life and death decision of stressed cells. If mitochondrial antiapoptotic proteins (orange) can effectively harness their C-terminal binding pockets to trap and sequester the BH3-signaling helices of proapoptotic proteins, cell survival prevails. However, with increased cellular stress, proapoptotic signals overwhelm the antiapoptotic reserve. The BH3 domain helices of BH3-only proteins (blue) can directly activate the essential executioner proteins BAX and BAK (gray) and also release trapped forms from antiapoptotic inhibition by targeting the antiapoptotic groove.
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alpha-helical peptides for use as research tools and prototype therapeutics (Fig. 2.3A). Depending on their amino acid composition and design, “stapled peptides” have proven to be structurally stable, protease resistant, and cell permeable agents capable of interrogating and modulating protein interactions in vitro and in vivo (Bernal et al., 2010; Kim et al., 2013; LaBelle et al., 2012; Takada et al., 2012; Walensky et al., 2004). Such stapled peptides have revealed unanticipated and functionally relevant helix/target interactions, including engagement of (1) antiapoptotic MCL-1 by its own MCL-1 BH3 helix (Stewart et al., 2010; Fig. 2.3B), (2) a novel “trigger site” on proapoptotic BAX by the BIM BH3 helix (Gavathiotis et al., 2008; Fig. 2.3C), and (3) glucokinase by a BH3-phosphorylated form of the BH3-only protein BAD (Danial et al., 2008). The capacity of these stabilized alpha-helices of BCL-2 domains (SAHBs) to access canonical and noncanonical protein interactors, and reveal sites of interaction by their use in NMR and X-ray crystallography studies, prompted us to consider whether they could be harnessed for higher throughput binding site discovery. Here, we describe our approach to converting stapled peptides into high fidelity photoaffinity reagents and the methodologies developed to rapidly identify their sites of target protein interaction.
2. OVERVIEW OF PHOTOREACTIVE STABILIZED ALPHA-HELICES METHODOLOGY We generate photoreactive stabilized alpha-helices or pSAHs by substituting into the peptide sequence a nonnatural amino acid bearing a benzophenone moiety (4-benzoylphenylalanine, Bpa) (Fig. 2.4A), which covalently crosslinks to protein targets upon exposure to ultraviolet (UV) light (Dorman & Prestwich, 1994; Saghatelian, Jessani, Joseph, Humphrey, & Cravatt, 2004; Vodovozova, 2007). We then insert an (i, i + 4) or (i, i + 7) pair of nonnatural amino acids bearing olefin tethers, followed by ruthenium-catalyzed ring-closing metathesis (RCM), to generate a hydrocarbon-stapled peptide with reinforced a-helical structure (Blackwell et al., 2001; Schafmeister, Po, & Verdine, 2000; Fig. 2.4B). For binding site identification, pSAHs containing alternatively placed Bpa moieties are individually incubated with a protein-of-interest and UV irradiated (Fig. 2.4C). The mixture is subjected to electrophoresis and the crosslinked species excised, trypsinized, and prepared for mass spectrometry analysis, which is designed to identify peptidic fragments bearing Bpa crosslinks and thus the explicit sites of target protein intercalation (Fig. 2.4D).
A
Insert nonnatural amino acids Cy3P CI
Ph
Ru
Chemically “staple” the peptide back into shape
O NH
O O
*
OH CH3
CI Cy3P
[ ]n
Unstructured peptide - Loss of functional shape - Easily degraded - Does not enter cells Hydrophobic Positive charge
Stapled peptide - Stabilized structure - Not degraded - Enters cells Hydrophilic Negative charge
B
C R214 D218
MCL-1 SAHBD BIM BH3 K21 R263 N260 D256
S255 Q32
Staple
R134 E131
MCL-1Δ NΔ C
BAX
Figure 2.3 Synthetic overview of all-hydrocarbon stapling and its applications in developing a-helical peptides for structural studies of protein interactions. (A) Pairs of a,a-disubstituted nonnatural amino acids bearing olefin tethers are substituted into the peptide sequence at discrete locations (e.g., i, i + 4), followed by rutheniumcatalyzed ring-closing metathesis (RCM) to generate “stapled peptides.” (B) Crystal structure of a stapled MCL-1 BH3 helix in complex with antiapoptotic MCL-1 (Stewart, Fire, Keating, & Walensky, 2010). (C) Calculated model structure of a BIM BH3 helix engaging the N-terminal trigger site of full-length BAX, as derived from paramagnetic relaxation enhancement NMR analyses using full-length 15N-BAX- and MTSL-labeled BIM SAHBs (aa 145–164) (Gavathiotis et al., 2008).
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A
B
Cy3P
O O
NH O
CI *
OH CH3
Ph
Ru CI
Cy3P
[ ]n
O
UV light
Trypsinolysis
D
O
O
C
O
O
E
F
Figure 2.4 Design of photoreactive stabilized alpha-helices (pSAHs) for binding site identification by mass spectrometry. A photoreactive Bpa residue (A) and a pair of stapling amino acids (B) are inserted into the peptide template followed by RCM to generate a pSAH. Upon exposure to UV light, the bound pSAH covalently crosslinks to the target protein (C) and, following electrophoresis of the mixture, crosslinked protein is excised from the gel and subjected to in-gel digestion with trypsin (D). LC–MS/MS analysis identifies the explicit sites of covalent modification, which when mapped on to the protein structure reveal the region of pSAH interaction (E). Top scoring crosslinks from a series of experiments employing sequentially placed Bpa residues can provide interaction restraints for calculating model structures (F).
By mapping the crosslink sites of pSAHs bearing alternatively placed Bpa moieties onto the protein structure (if known), the location of the interaction site can be determined (Fig. 2.4E). Furthermore, if the structure of the protein target has been solved and a series of pSAHs with sequentially localized Bpa residues map sites of intercalation with sufficient resolution,
Photoreactive Stapled Peptides
31
computational docking can be applied to calculate a model structure of the complex for biochemical and functional validation (Fig. 2.4F).
3. DESIGN AND SYNTHESIS OF PHOTOREACTIVE STAPLED PEPTIDES The design of photoreactive stapled peptides is ideally based on a structure involving the template a-helix, so that the structurally reinforcing staples can be placed on the noninteracting surface and Bpa residues located at or near the putative interaction surface to facilitate UV crosslinking. Ideally, Bpa residues are substituted at positions of homologous, bulky, and hydrophobic residues (e.g., Phe) to minimize the effect of Bpa on the binding interface. Without structural information, a library of constructs can be generated by sampling alternative staple and Bpa positions along the length of the peptide template, selecting those constructs with successful a-helical induction (as determined by circular dichroism analysis; Bird, Bernal, Pitter, & Walensky, 2008) and preserved binding activity for experimental application. Of note, to simplify interpretation of the MS2-based identification of Bpa-crosslinked peptides, it is also preferable to locate Bpa residues in regions of the peptide such that trypsinization will yield short cleavage products. If necessary, a lysine or arginine residue can be strategically substituted into the peptide, ideally on the noninteracting surface or as a conserved substitution observed in other species (Leshchiner, Braun, Bird, & Walensky, 2013), to accomplish this goal. Once designed, stapled peptides are synthesized as previously described in detail (Bird et al., 2008; Bird, Crannell, & Walensky, 2011) using Fmoc chemistry, stapling amino acids installed at (i, i + 4) or (i, i + 7) positions (to bridge one or two turns of the a-helix, respectively), and ruthenium-catalyzed olefin metathesis with Grubbs generation I catalyst (Fig. 2.5). Of note, methionine residues in the template sequence are typically replaced by norleucines, as the unprotected sulfur can decrease the efficiency of the Grubbs catalyst. Following peptide deprotection and cleavage, pSAHs are purified by reverse phase high performance liquid chromatography–mass spectrometry and quantified by amino acid analysis (AAA). Pure peptide is stored as a lyophilized powder at 20 C until use. In addition to the experimental conditions summarized in Fig. 2.5 and the step-by-step synthetic protocol with reagent list detailed in our recently updated methodologic review (Bird et al., 2011), our approach to stapled peptide synthesis incorporates the following general principles:
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R Fmoc
1. 20% piperidine in NMP
H N
N H
HCTU/DIEA in NMP
O
n
O
H
2.
Fmoc N
OH Bp
1. 20% piperidine in NMP HCTU/DIEA in NMP
2. Fmoc
H N
O
O
(S)
OH
H Fmoc N
R N H
Bp O
H N
Fmoc
R N H
R
O
H N
N H
O
HCTU/DIEA in NMP
N H
1. 20% piperidine in NMP
O
R N H
O
HCTU/DIEA in NMP
R
2.
H N
n
O
1. 20% piperidine in NMP
Fmoc
H N O
3
n
R
N H
O
n⬘
H N
R
O
H N
Fmoc
Fmoc
OH
N H
OH
N H
O
O O
R
H N
Fmoc
O
N H
H N O
n⬙
n
3
R
2.
R N H
N H
O
Bp
R
O
H N
H N
O
O
n⬘
R N H
H N O
n
3
1. 20% piperidine in NMP 2. Fmoc
H N
O (S)
1. 20% piperidine in NMP
OH
HCTU/DIEA in NMP
Fmoc
O
H N
2. Ac2O, DIEA, NMP
R N H
O
H N
R
H N
N H
O
n
3
O
R Ac
O
N H
H N O
R N H
O
H N O
Bp
R N H
O
R N H
O
n⬘
n⬙
H N
H N O
n
3
1. 20% piperidine in NMP 2.
HCTU/DIEA in NMP
1.
R Fmoc
PCy3
Cl
OH
N H
, DCE
Ru Cl
O
PCy3
2. TFA:TIS:H2O (95:2.5:2.5) R Fmoc
N H
H N O
n⬘
R
O N H
H N O
3
R
O N H
H N O
n
O
R Ac
N H
H N O
n⬙
R N H
Bp
H N O
n⬘
R
O N H
H N O
O
R N H
NH 2 O
n
Figure 2.5 Synthetic steps for the automated production of pSAHs by Fmoc-based solid-phase peptide synthesis and ruthenium-catalyzed RCM. Alternatives to N-terminal acetylation include capping with FITC or biotin, depending on the desired application. Bp, 4-benzoylphenyl.
a. Create an automated synthesis program by employing standard double Fmoc deprotections, vigorous washes with DMF, and subsequent single 10-fold excess or double fivefold excess amino acid couplings. The precise timing and coupling excesses are optimized based on the specific
Photoreactive Stapled Peptides
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coupling reagents, bases, and synthesizer employed. Particular attention is devoted to reactions involving the a,a-disubstituted amino acids, as the amine is linked to a quaternary carbon center, and therefore, can be recalcitrant to Fmoc removal and the subsequent acylation steps. For example, couplings that involve beta-branched amino acids and arginine can be especially challenging. Typical protocol modifications include multiple extended deprotections and acylations, and sometimes capping at these difficult steps with acetic acid (AcOH) or even decanoic acid in order to capture and terminate unreacted amine. In this manner, the production of full-length peptide species can be maximized, while also avoiding the challenge of chromatographic separation of truncated byproducts. b. Prepare the peptide synthesis reagents and solutions by weighing out the resin and dissolving the calculated amount of each amino acid in anhydrous NMP. Although more expensive than DMF, NMP has greater stability and we have seen no deterioration in peptide quality using reagent solutions for up to 2 weeks. A secondary amine is used to remove the Fmoc protective group; appropriate choices include piperidine, piperazine, or 1,8-diazabicycloundec-7-ene. Likewise, there are multiple coupling reagents to select from. HCTU/DIEA and HOBt (or HOAt)/DIC nicely balance reactivity with solution stability, and more reactive coupling reagents range from uronium based (HATU) to the more exotic tris-pyrrolidinophosphonium derivatives. c. Evaluate synthetic success by subjecting a small sample of beads to standard trifluoroacetic acid (TFA) cleavage (95:2.5:2.5 TFA:water: triisopropylsilane) and analysis by LC/MS. If the full-length species is identified in good yield, the next step is acylation with acetic anhydride or Fmoc-b-alanine, followed by stapling with Grubbs I catalyst (bis (tricyclohexylphosphine) benzylidine ruthenium (IV) dichloride) dissolved in dichloroethane. For most sequences and staple locations, a 2 2 h incubation using several milliliters of a 4 mg/mL solution will yield a completely metathesized product. If there is incomplete metathesis, the reaction can be repeated multiple times and with longer incubation periods. d. After confirmation of successful stapling, the Fmoc group of Fmocb-alanine is removed and the N-terminus derivatized with biotin or fluorescein isothiocyanate (FITC), depending on the application (see later). The final product is cleaved off the resin in large scale, precipitated with ether:hexanes, and purified by HPLC or, if available, by LC/MS with mass-triggered fraction collection. The pure fractions are pooled,
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lyophilized, and peptide powder resuspended in a known volume, followed by submission of several dilutions for AAA. Although we typically store stapled peptides as dried powder (see above), the material can also be solubilized in 100% DMSO for storage at 20 C.
4. PHOTOAFFINITY LABELING AND RETRIEVAL With a panel of photoreactive stapled peptides in hand, the next step is to isolate pure protein-of-interest for photoaffinity labeling. We have typically expressed glutathione S-transferase, chitin-binding domain, or histidine-tagged fusion proteins followed by affinity purification, cleavage of the tag, and size exclusion chromatography. Protein purity and identity are confirmed by SDS-PAGE, protein staining (e.g., coomassie, silver), protein-specific western blot, and mass spectrometry analyses. Protein concentrations are determined using the Bradford assay (Biorad). Photoreactive stapled peptide and pure recombinant protein (e.g., 10 mM) are typically mixed at a 1:1 ratio, but higher pSAH concentrations can be used if necessary to increase the quantity of crosslinked product. The peptide/protein mixture is vortexed, incubated for 20 min in the dark, and then irradiated (365 nm) on ice for 1.5 h by use of a Spectroline handheld UV lamp (Model En280L, Spectronics Corporation). The duration of UV exposure can also be adjusted based on the efficiency of crosslinking for a particular peptide/protein complex. The mixtures are then diluted with 4 LDS loading buffer (Invitrogen), electrophoresed (e.g., 4–12% Bis–Tris gels [Invitrogen]), and analyzed by coomassie staining (SimplyBlue Safestain, Invitrogen). The appearance of a slower migrating band just above the protein starting material reflects the pSAH-crosslinked species, which is excised for MS analysis (see below). Several supplementary experiments can be especially useful when piloting this approach. For example, to confirm that insertion of staples and Bpa residues at discrete sites within the peptide template do not disrupt target engagement, we recommend generating fluorescently labeled (FITC) pSAH derivatives for comparative fluorescence polarization (FP) binding analyses with a positive control ligand. In the case of FITC-BAD pSAHBs, the substitution of Bpa residues at four positions within the BAD BH3 sequence had no disruptive effect on binding to the antiapoptotic protein BCL-XLDC (Braun et al., 2010; Fig. 2.6A). The FITC-derivatized pSAHs can also be used for photoaffinity labeling, with conversion of recombinant protein into a fluorescent species that can readily be detected by fluorescence
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imaging (Gel Doc XR Molecular Imager and Quantity One software, BioRad), providing further confirmation of effective crosslinking (Braun et al., 2010; Fig. 2.6B). Biotinylated pSAHs can also be used for high sensitivity screening of effective crosslinking, with electrophoresed protein analyzed by anti-biotin western blot (see below). Binding specificity experiments using bovine serum albumin (BSA), for example, are also advised. Crosslinking is performed as above, except that BSA is added in molar excess Coomassie
Compound
BAD SAHB BAD pSAHB-1 BAD pSAHB-2 BAD pSAHB-3 BAD pSAHB-4
C
60.5 (51.9–70.6) 46.7 (44.2–49.4) 38.9 (35.9–42.1) 38.5 (36.5–40.6) 55.6 (53.1–58.1)
BCL-XLΔC Fluorescence BSA Crosslinked BCL-XLΔC BCL-XLΔC
100
60
0 10 10
D
20
10–9
10–8
10–7
10–6
10–5
[BCL-XLΔC], M
0 0.5 1
Fluorescence
Ve hi BA cle D BA pS D AH BA pS B-1 D AH BA pS B-2 D AH p B Ve SA -3 H hi BA cle B-4 D p BA S D AH BA pS B-1 D AH BA pS B-2 D AH pS BAH 3 B4
Coomassie
10 10 10
30 10 10
Fluorescence
Crosslinked BCL-XLΔC BCL-XLΔC
B
.5 10 10
Coomassie
Crosslinked BCL-XLΔC BCL-XLΔC
40
Crosslinked BCL-XLΔC BCL-XLΔC
10 0 0
BSA (mM) BCL-XLΔC (mM) BAD pSAHB-2 (mM)
80
0 10–10
BSA Crosslinked BCL-XLΔC
BAD pSAHB-2 crosslinking yield (%)
Normalized polarization (%)
A
EC50 95% CI
2
5 10 20 30 40 50 60 80 100 120 Time (min)
40
30
20
10
0 0
20
40
60
80
100
120
Time (min)
Figure 2.6 pSAHs bind and selectively crosslink to their protein targets, as exemplified by the photocrosslinking of BAD pSAHBs to antiapoptotic BCL-XLDC (Braun et al., 2010). (A) A series of FITC-BAD pSAHBs retain nanomolar binding affinity to BCL-XLDC as measured by FP analysis. (B) FITC-BAD pSAHBs exhibit a spectrum of BCL-XLDC-crosslinking efficiency upon UV exposure, with BAD pSAHB-2 demonstrating the greatest reactivity toward BCL-XLDC (left, coomassie stain; right, fluorescence scan). (C) The selectivity of BAD pSAHB-2 is reflected by the absence of crosslinking to BSA and the lack of effect of added BSA on the BCL-XLDC-crosslinking efficiency of FITC-BAD pSAHB-2. (D) Time course for photocrosslinking of FITC-BAD pSAHB-2 to BCL-XLDC as monitored by coomassie stain (top) and fluorescence scan (middle). A plot of crosslinking yield, calculated based on densitometry, demonstrates the time-dependent production of crosslinked BCL-XLDC (bottom).
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prior to addition of pSAH. In the case of FITC-BAD pSAHB-2/BCL-XLDC crosslinking, the addition of molar excess BSA had no effect on the yield of pSAHB-crosslinked BCL-XLDC (Braun et al., 2010; Fig. 2.6C). Tracking the production of pSAH-crosslinked protein over time is useful for determining just how long to extend the reaction time to produce maximal reaction product for the MS analyses (Fig. 2.6D). If the efficiency of pSAH crosslinking is suboptimal after increasing the peptide/protein ratio and extending the reaction time (i.e., crosslinked species not readily visualized by coomassie stain), the capacity to load sufficient material for electrophoresis and band excision can be compromised. Although we typically find that sampling a variety of pSAH constructs reveals lead photoaffinity labeling reagents (Fig. 2.6B), an alternative approach to enrich for crosslinked species entails the synthesis and application of pSAHs capped at the N-terminus with biotin. Photoaffinity labeling with biotinylated pSAHs is performed as above and then unreacted peptide is removed from the irradiated samples by overnight dialysis at 4 C in 50 mM Tris pH 7.4, 200 mM NaCl buffer using 6–8 kDa molecular weight cut-off D-Tube dialyzers (EMD Biosciences). After addition of SDS to a final concentration of 0.2%, biotinylated protein is isolated from unreacted protein by incubation with high capacity streptavidin agarose (50 mL 50% slurry/reaction) for 2 h at room temperature. The streptavidin beads are successively washed at room temperature in 1% SDS in PBS, 1 M NaCl in PBS, and then 10% ethanol in PBS for 3 10 min each. Biotinylated proteins are eluted by boiling for 30 min in a 10% SDS solution (Promega) containing biotin (10 mg/mL), electrophoresed (e.g., 4–12% Bis–Tris gels [Invitrogen]), and then subjected to coomassie staining. Using this enrichment protocol for purifying biotinylated pSAH-crosslinked protein, isolating sufficient material for MS analyses of even low efficiency crosslinked complexes (e.g., weak or transient protein interactions) can typically be achieved (Edwards et al., 2013; Leshchiner et al., 2013).
5. MASS SPECTROMETRY ANALYSIS To prepare pSAH-crosslinked protein for MS analysis, we perform an in-gel digest of the excised protein band using a standard protocol (Braun et al., 2010), mass spectrometry grade reagents, and due caution to avoid keratin contamination. Gels are subjected to coomassie or other MS-compatible stain (e.g., Pierce Silver Stain) and the excised gel slab cut into 1 mm cubes for processing as follows:
Photoreactive Stapled Peptides
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a. Destain: Add 500 mL of 50% acetonitrile (ACN)/50 mM ammonium bicarbonate (AB), pH 8 and incubate at 37 C with shaking for 15 min; remove solution and repeat until the blue color is eliminated from the gel pieces. b. Dehydrate: Add 500 mL of ACN, incubate at room temperature for 5 min, remove solution, and repeat. c. Reduce disulfides: Add 100 mL of 10 mM dithiothreitol in AB, incubate at 56 C for 30 min, and remove solution. d. Alkylate cysteines: Add 100 mL of 50 mM iodoacetamide in AB, incubate at room temperature in the dark for 20 min, and then remove solution. e. Dehydrate: As above. f. Digest: Cover gel pieces with cold 12.5 ng/mL trypsin (Promega) in AB, incubate on ice for 45 min and then overnight at 37 C. Peptides are extracted twice in 50% ACN/5% formic acid (FA) and dried in a speed-vac. Samples are prepared for LC–MS/MS using home-made C18 stage tips or Agilent C18 Omix tips (Braun et al., 2010; Jakobsen, Schroder, Larsen, Lundberg, & Andersen, 2013). The sample preparation procedure below employs 100 mL Omix tips: a. Place 6 mL of elution buffer (80% ACN/0.5% AcOH) in a clean elution tube for each sample. b. Resuspend the dried peptides in 100 mL of 0.1% TFA. c. Wet tips by pipetting 2 100 mL 40% ACN/0.5% AcOH. Be sure to prevent air from being introduced into the tips once wetted. d. Equilibrate tips by pipetting 2 100 mL of 0.1% TFA. e. Bind peptides in sample by carefully pipetting up and down 10 times. f. Wash by pipetting 3 100 mL of 0.1% TFA. g. Elute by carefully pipetting up and down in a prepared elution tube three times. h. Dry the eluate by speed-vac. We use a split flow LC system with in-house packed C4/C18 columns for MS analysis on a Thermo LTQ Orbitrap Discovery mass spectrometer. Samples are reconstituted in 5% ACN/5% FA and loaded onto the LC. We employ 30 cm of 100 mm inner diameter fused silica tips packed with
