September 14, 2005
102 (39) 13767-13772
Conotoxins (CTXs), with their exquisite specificity and potency, have recently created much excitement as drug leads. However, like most peptides, their beneficial activities may potentially be undermined by susceptibility to proteolysis in vivo. By cyclizing the α-CTX MII by using a range of linkers, we have engineered peptides that preserve their full activity but have greatly improved resistance to proteolytic degradation. The cyclic MII analogue containing a seven-residue linker joining the N and C termini was as active and selective as the native peptide for native and recombinant neuronal nicotinic acetylcholine receptor subtypes present in bovine chromaffin cells and expressed in Xenopus oocytes, respectively. Furthermore, its resistance to proteolysis against a specific protease and in human plasma was significantly improved. More generally, to our knowledge, this report is the first on the cyclization of disulfide-rich toxins. Cyclization strategies represent an approach for stabilizing bioactive peptides while keeping their full potencies and should boost applications of peptide-based drugs in human medicine.
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Venoms from marine snails of the Conus genus comprise a myriad of peptides called conotoxins (CTXs) for the rapid immobilization of prey (1, 2). These 12- to 30-aa peptides target membrane receptors with exquisite selectivity and potency and have become invaluable neurophysiological probes and drug leads. Recently, the CTX ziconotide (MVIIA) was approved for use in the treatment of severe chronic pain by the FDA, and other CTXs have entered clinical trials as treatments for pain (3, 4). In addition, CTXs have played a critical role in dissecting the molecular mechanisms of ion channel and transporter functions in the nervous system (2). One family of CTXs, the α-CTXs, consists of members that antagonize the nicotinic acetylcholine receptors (nAChRs). Ranging in size from 12 to 19 residues, α-CTXs are the smallest of all of the CTXs, yet this family is the most widely distributed among Conus venoms (5).
Despite their exciting applications, many peptide toxins are susceptible to enzymatic degradation by proteases. This characteristic may limit the therapeutic applications of CTXs, and, hence, methods that provide improvements in biological half-life are valuable. Cyclization has been used in the past as a strategy in the pharmaceutical industry for stabilizing and locking the conformation of small peptides (6). Similarly, microorganisms are known to produce cyclized peptides, such as cyclosporin A, which is now in widespread use as an immunosuppressant. Such a strategy has not been applied in the past to disulfide-rich proteins, but with the recent discovery of the cyclotide family of macrocyclic miniproteins (7), it is clear that the approach can be applied to disulfide-rich toxins to produce additional stabilization with the potential to dramatically increase the therapeutic potential of these molecules when limited by poor in vivo stability.
This study focuses on the cyclization of MII, a 16-residue α-CTX isolated from Conus magus (8). The 3D structure of MII consists of a central segment of α-helix with β-turns at the N and C termini (9, 10) and is stabilized by two disulfide bonds in a CysI-CysIII and CysII-CysIV configuration that is common to most members of the α-CTX family. In addition, the N and C termini of the peptide are in close proximity to each other, making MII a good candidate for studying the principles of backbone cyclization. MII is a potent inhibitor of the nAChR that is specific for the α3β2 subtype (8) and is also implicated in binding to the α6 nAChR, ligands of which are potentially important for Parkinson's disease therapy (11). There are currently a number of patents describing the use of MII in therapeutic applications.
To illustrate the advantage of cyclization of linear proteins, we designed and synthesized three cyclic MII analogues by adding a linker segment between the N and C termini. Structural studies of the analogues were undertaken, and activity and stability assays were performed. To our knowledge, this is the first study on the cyclization of CTXs. We also discuss the potential for backbone cyclization to enhance the therapeutic potential of peptide toxins.
Peptide design was based on an analysis of homology models generated by using the structural coordinate file of MII (Protein Data Bank ID Code 1MII), available from the PDB (www.rcsb.org/pdb), and the modeler module within insight ii (Accelrys, Inc., San Diego). Energy-minimized linkers of varying sizes were built into the linear MII molecule, and the resulting cyclic analogue models were evaluated.
All peptides were assembled on phenylacetamidomethyl resin by manual solid-phase peptide synthesis using the in situ neutralization/HBTU [2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexaf luorophosphate] protocol for Boc (tert-butoxycarbonyl) chemistry (12). The peptide was attached to the resin by means of a linker that generates a C-terminal thioester on hydrogen fluoride (HF) cleavage, which facilitates the intramolecular native chemical ligation reaction used to cyclize the peptide (13). Cleavage of the peptide from the resin was achieved by using HF with p-cresol and p-thiocresol as scavengers [9:0.8:0.2 (vol/vol) HF:p-cresol:p-thiocresol] at –5 to 0°C for 1.5 h. After cleavage, the peptide was precipitated with ether and then dissolved in 50% acetonitrile containing 0.05% trifluoroacetic acid (TFA) and lyophilized. The crude peptide was purified by RP-HPLC on a C 18 column using a gradient of 0–80% B (A, H 2 O/0.05% TFA; B, 90% CH 3 CN/10% H 2 0/0.045% TFA) in 80 min. Analytical RP-HPLC and electrospray MS confirmed the purity and molecular mass of the synthesized peptide.
The linear reduced peptides were cyclized and oxidized by incubating in 50/50 0.1 M NH 4 HCO 3, pH 8.2/isopropanol (0.3 mg/ml) overnight at room temperature. The reaction mixture was purified by RP-HPLC to yield the cyclic/oxidized peptides. Analytical RP-HPLC and electrospray MS confirmed the purity of the final products, and 1 H NMR was used to determine whether the cyclic peptides were folded.
The disulfide connectivity of the cyclic analogues was determined by using a reduction/alkylation procedure followed by MS/MS analysis. Partial reduction was achieved by incubating the peptide (50 μg) in 0.2 M citrate buffer (pH 3, 50 μl) with 50 equivalents of Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) at 37°C. The reaction was left for 3 min, after which it was quenched by injection onto the RP-HPLC. The collected HPLC fractions were analyzed by MS to identify the one-disulfide (1SS) species. An equal volume of 60 mM N-ethylmaleimide in 0.2 M citrate buffer (pH 3) was then added to the HPLC fraction containing the 1SS peptide and left at 37°C for 1 h to alkylate the free Cys residues. The peptide was purified by RP-HPLC and then fully reduced by incubating with 10 equivalents of DTT in 0.1 M NH 4 HCO 3 (pH 8.2) for 3 h at 37°C. The reduced alkylated peptide was isolated by RP-HPLC, lyophilized, and cleaved with EndoGluC by incubating for 3 h in 0.1 M NH 4 HCO 3 with 2 equivalents of TCEP at 37°C. The peptide was purified by RP-HPLC, and the linear product, containing alkyl groups on one pair of cysteines, was analyzed by MS/MS, which enabled the identification of the two alkylated cysteines and hence the disulfide connectivity.
The structures of the cyclic MII analogues were derived by using NMR spectroscopy for samples dissolved in 90% H 2 O and 10% D 2 O. Avance 600 and DMX 750 MHz spectrometers (Bruker, Billerica, MA) were used in the acquisition of data. 2D NMR experiments included double quantum filtered (DQF)-COSY, exclusive (E)-COSY, TOCSY (total correlation spectroscopy), and NOESY, with mixing times of 150 and 300 ms. All spectra were recorded at 298 K. 1 H spectra acquired immediately after dissolution of the protonated peptide in D 2 O at pH 4.7 were used to detect slow-exchanging amide protons. The peak areas in these spectra were measured and normalized to a nonexchangeable proton, and the exchange rates were calculated by fitting the volume of the decaying signals over time to the equation I = I o exp(–k ex× t) + I(∞) (14).
Distance information was obtained from the NOESY spectrum with a mixing time of 300 ms. Backbone dihedral restraints were derived from 3 J HN-Hα coupling constants obtained from a 2D DQF-COSY spectrum or from a 1D 1 H NMR spectrum. The φ angle was restrained to 120 ± 30° for 3 J HN-Hα> 8 Hz and –60 ± 30° for 3 J HN-Hα< 5.8 Hz. Intraresidue nuclear Overhauser effect (NOE) and 3 J Hα-H β coupling patterns, determined from an E-COSY spectrum, were used in assigning the χ 1 angle conformations of side chains.
Initial structures were generated by using dyana software (15), and final structures were calculated in explicit water with cns (16) as described in ref. 17. Fifty structures were calculated, and the 20 structures with the lowest overall energies were retained for analysis. Structures were analyzed with promotif (18) and procheck _ nmr (19).
The biological activity of the cyclic analogues was assessed by recording membrane currents using the dialyzed whole-cell patch clamp recording technique and by monitoring catecholamine secretion from adrenal chromaffin cells.
Chromaffin cells were prepared from bovine adrenal glands and maintained on glass coverslips as described in ref. 20. Glass electrodes were pulled, fire-polished (–2 to 3 MΩ) and filled with intracellular solution (in mM: 140 CsCl/2 CaCl 2/11 EGTA/2 MgATP/10 Hepes-KOH, pH 7.2). Agonists were diluted in bath solution (in mM: 140 NaCl/3 KCl/1.2 MgCl 2/2.5 CaCl 2/7.7 glucose/10 Hepes-NaOH, pH 7.35) and applied to cells by a 10-ms pressure ejection (15 psi, Picospritzer II, General Valve, Fairfield, NJ) from an extracellular pipette positioned ≈50 μm from the cell to evoke maximal responses to agonists (21). CTXs were bath-applied. Membrane currents evoked by agonist application were amplified and low-pass filtered (10 kHz) by using a MultiClamp 700B amplifier (Axon Instruments, Union City, CA), and voltage steps were generated by using pclamp 9.2 and a Digidata 1322A interface (Axon Instruments). All experiments were carried out at 22°C.
RNA preparation, oocyte preparation, and expression of nAChR subunits in Xenopus oocytes were performed as described in ref. 22. cDNA encoding the rat α2-7 and β2-4 nAChR subunits were provided by J. Patrick (Baylor College of Medicine, Houston). Oocytes were injected with 2.5 ng of cRNA and kept at 18°C in ND96 buffer (96 mM NaCl/2 mM KCl/1 mM CaCl 2/1 mM MgCl 2/5 mM Hepes, pH 7.4) supplemented with 50 mg/liter gentamycin and 5 mM pyruvic acid 2–5 days before recording.
Membrane currents were recorded from Xenopus oocytes by using an OpusXpress 6000A workstation (Axon Instruments). Electrodes were filled with 3 M KCl (–0.3 to 1.5 MΩ). During recordings, the oocytes were perfused with ND96 buffer at 22°C continuously at a rate of 1.5 ml/min, with 200-s incubation times. Acetylcholine (100 μM) was applied for 2 s at 5 ml/min, with 600-s washout periods. Cells were held at –80 mV with data sampled at 500 Hz and filtered at
