Palladium–peptide oxidative addition complexes for bioconjugation

The synthesis of palladium oxidative addition complexes derived from unprotected peptides is described. Incorporation of 4-halophenylalanine into a peptide during solid phase peptide synthesis allows for subsequent oxidative addition at this position upon treatment with a palladium precursor and suitable ligand. The resulting palladium–peptide complexes are solid, storable, water-soluble, and easily purified via high-performance liquid chromatography. These complexes react with thiols in aqueous buffer, offering an efficient method for bioconjugation. Using this strategy, peptides can be functionalized with small molecules to prepare modified aryl thioether side-chains at low micromolar concentrations. Additionally, peptide–peptide and peptide–protein ligations are demonstrated under dilute aqueous conditions.


Determination of Bioconjugation Conversions
Data were processed using Agilent MassHunter software package. All reported yields were determined by integrating total ion current (TIC) spectra. First, the peak areas for all relevant peptide-containing species on the chromatogram were integrated using Agilent MassHunter software package. Since no peptide-based side products were generated in the experiments, the yields shown in Schemes 1, 2, and 3 were determined as follows: %yield = Spr/Stotal where Spr is the peak area of the product and Stotal is the peak area of combined peptide-containing species (product and starting material). For protein bioconjugation, deconvoluted masses of proteins were obtained using maximum entropy algorithm. LC-MS data shown were acquired using Method A, unless otherwise noted. Mass spectrum insets correspond to the integration of the TIC peak unless otherwise noted. In order to provide clarity, an example of how bioconjugation conversion is detailed below:

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The crude reaction mixture is depicted below: This mixture was analyzed by LC-MS using Agilent MassHunter software package. The peptidecontaining peaks that correspond to the starting material and product were identified by mass as can be seen from the inlets depicted below.
Finally, using the calculation described above, the Agilent MassHunter software package was used to integrate below the starting material and product peaks to determine the percent of conversion to the product.

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In some cases, a small non-peptidic impurity near the starting material was found, resulting in an additional peak. This peak was omitted from conversion calculations because it does not correspond to total peptide-containing species.
A longer, 26-minute LC-MS method was also performed to confirm the lack of byproducts produced from the crude reaction.

Linear Peptide Synthesis
General Peptide Synthesis Procedure: All peptides were synthesized on a 0.2 mmol scale using automated Fmoc-SPPS chemistry under flow as previously described. 2 Specifically, ChemMatrix Rink Amide HYR resin (200 mg) was loaded into a reactor maintained at 90 ºC. All reagents were flowed at 80 mL/min with HPLC pumps through a stainless-steel loop maintained at 90 ºC before introduction into the reactor. For each coupling, 10 mL of a solution containing 0.2 M amino acid and 0.17 M HATU in DMF were mixed with diisopropylethylamine (200 μL) and delivered to the reactor. Fmoc removal was accomplished using 20% (v/v) piperidine (10.4 mL). Between each step, DMF (15 mL) was used to wash out the reactor. Special coupling conditions were used for arginine, in which the flow rate was reduced to 40 mL/min and 10 mL of a solution containing 0.2 M Fmoc-L-Arg(Pbf)-OH and 0.17 M PyAOP in DMF were mixed with diisopropylethylamine (200 μL) and delivered to the reactor.
After completion of the stepwise, automated SPPS, the resin was washed thoroughly with CH2Cl2 and dried under vacuum. The peptide was simultaneously cleaved from the resin and the side-chains were deprotected by treatment with 5mL of the cleavage cocktail containing 82.5% TFA, 5% phenol, 5% thioanisole, 5% water, and 2.5% EDT (v/v) at 60 °C for 8 min. The resulting solution was then triturated and washed with cold diethyl ether (3x). The obtained solid was dissolved in a solution of 50% H2O and 50% acetonitrile containing 0.1% TFA and lyophilized. The following peptides were synthesized following this procedure:

Peptide Purification
Peptides were dissolved in water and acetonitrile containing 0.1% TFA, filtered through a 0.22 μm nylon filter and purified by mass-directed semi-preparative reversed-phase HPLC (RP-HPLC). Solvent A was water with 0.1% TFA additive and Solvent B was acetonitrile with 0.1% TFA additive. A linear gradient from 5-45% B changing a rate of 0.5% B/min was used. The peptides were purified on an Agilent Zorbax SB C3 column: 9.4 x 250 mm, 5 μm. Using the mass data collected for each fraction from the instrument, only pure fractions were combined and S9 lyophilized. The purity of the fraction pool was confirmed by LC-MS. Total-ion current chromatograms for each peptide is shown below.

Synthesis of Pd-Peptide Oxidative Addition Complexes
Synthesis of [(1,5-COD)Pd(CH 2 TMS) 2 ]. This compound was synthesized according to the procedure reported in the literature. 1 The 1 H and 13 C NMR spectra of the obtained material are identical to those reported in the literature. 1 The title compound was stored in a freezer at -20 °C.
General Procedure for the Synthesis of Palladium-Peptide Oxidative Addition Complexes.
In a scintillation vial (10 mL) open to the air and equipped with a magnetic stir bar, the vial was charged with sSPhos ligand (2.5 equiv), peptide-Ar-X (1 equiv), and DMSO (1 mL). Solid (1,5-COD)Pd(CH2SiMe3)2 (2.5 equiv) was added rapidly in one portion and the resulting solution was stirred for 1 h at rt.* After this time, the solution was frozen by placing the capped vial into a liquid nitrogen bath and subsequently lyophilized to remove the DMSO. The Pd-Peptide oxidative addition complexes were purified via mass-directed preparative RP-HPLC.

Palladium-Peptide Oxidative Addition Complex Purification
Although the purified palladium-peptide oxidative addition complexes are readily soluble in aqueous solutions, we found that the crude palladium-peptide oxidative complexes exhibited limited solubility, likely due to the excess ligand. Therefore, after lyophilization, the palladiumpeptide oxidative addition complexes were dissolved in 2 mL of a 50:50 (v/v) mix of water and acetonitrile containing 0.1% TFA. The solution was diluted with 1 mL of water containing 0.1% TFA, filtered through a 0.22 μm nylon filter, and purified by mass-directed semi-preparative reversed-phase HPLC (RP-HPLC). Solvent A was water with 0.1% TFA additive and Solvent B was acetonitrile with 0.1% TFA additive. A linear gradient from 15-50% B changing at a rate of 0.5% B/min was used. The peptides were purified on an Agilent Zorbax SB C3 column: 9.4 x 250 mm, 5 μm.
Using mass data from the instrument about each fraction, pure fractions were pooled and lyophilized. Despite different retention times, peptide OA complexes and ligand often co-elute, which can reduce overall yield (particularly for 1C). The purity of the fraction pool was confirmed by LC-MS. Total-ion current chromatograms for each complex is shown below. With electrospray ionization, we observe ions corresponding to each palladium-peptide oxidative addition complex without sodium and halide. An additional characterization of OA complex 1B using MALDI-TOF-MS is also presented, and we also observe the loss of sodium and halide.

1A:
The LC-MS chromatogram of OA complex 1A has a peak at 8.4 min that corresponds to free ligand. HPLC purification fails to fully resolve the complex from the ligand, and techniques to improve the purification have been unsuccessful but the subsequent reactivity of the complex is identical to that of the other Pd-peptide complexes.

1E:
For the LC-MS chromatogram of OA complex 1E, there are two peaks observed at 8 min. These peaks both have the same mass, and we hypothesize they arise due to two distinct conformations of the ligand when the bromophenylalanine is in the middle of the peptide (i.e. rotamers).

MALDI-TOF-MS Analysis
For purified OA complex 1B, the compound was also analyzed using MALDI-TOF-MS. Shown below is the spectrum obtained after preparing the sample in -cyano-4-hydroxycinnamic acid matrix. The spectrum was acquired in positive reflector mode, with PIE delay set to 240 ns, laser repetition rate set to 60 Hz, reflector detector voltage set to 1.854 kV, ion source voltage 1 set to 19 kV, ion source voltage 2 set to 15.9 kV, ion source lens voltage set to 9.1 kV, and the number of shots set to 400. Mass calculated C89H121N19O20PPdS + [M-NaBr+H] +1 : 1944.75. Mass observed: 1944.19. The additional peak at 1350.9 corresponds to M -Na, -Br, -Pd, -ligand, +H.

PALLADIUM-PEPTIDE REACTIONS WITH SMALL MOLECULE THIOLS
General Bioconjugation Procedure A solution of the palladium-peptide reagent 1B (20 µM) in water was added to an equal volume of a solution containing the small molecule thiol (100 µM) in 0.1 M TRIS buffer at pH 7.5. The palladium reagents were sonicated for 10 s to ensure the reagents were fully dissolved. Final conditions: [Pd] = 10 µM; [R-SH] = 50 µM; After 1 h at 37 °C, of 3-mercaptopropionic acid (3 equiv) dissolved in H2O was added to the reaction mixture to quench the remaining palladium species. The reaction was allowed to stand for 5 min and subsequently characterized by LC-MS.
Following the general bioconjugation procedure with the N-acetylated palladium-peptide depicted (10 µM) 4-(trifluoromethyl)benzenethiol (50 uM) resulted in a 99% conversion to the desired product as determined by LC-MS analysis.

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Following the general bioconjugation procedure with the depicted palladium-peptide with free Nterminus (10 µM) and 4-mercaptophenylboronic acid (50 uM) resulted in a 98% conversion to the desired product as determined by LC-MS analysis.
Following the general bioconjugation procedure with the depicted N-acetylated palladium-peptide (10 µM) and 4-aminothiophenol (50 uM) resulted in a 99% conversion to the desired product as determined by LC-MS analysis.

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Following the general bioconjugation procedure with the depicted N-acetylated palladium-peptide (10 µM) and 4-mercaptophenol (50 uM) resulted in a 98% conversion to the desired product as determined by LC-MS analysis.

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Following the general bioconjugation procedure with the depicted N-acetylated palladium-peptide (10 µM) and 2-fluorothiophenol (50 uM) resulted in a 98% conversion to the desired product as determined by LC-MS analysis.
Following the general bioconjugation procedure with the depicted N-acetylated palladium-peptide (10 µM) and 4-mercaptotetrahydropyran (50 uM) resulted in a 99% conversion to the desired product as determined by LC-MS analysis.

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Following the general bioconjugation procedure with the depicted palladium-peptide with free Nterminus (10 µM) and thiocholesterol (50 uM) resulted in a 99% conversion to the desired product as determined by LC-MS analysis.
Following the general bioconjugation procedure with the depicted palladium-peptide with free Nterminus (10 µM) and 4-phenylthiazole-2-thiol (50 uM) resulted in a 98% conversion to the desired product as determined by LC-MS analysis.

General pH Study Procedure -(Buffer Experiments)
A solution of the depicted N-acetylated palladium-peptide reagent (20 µM) in water was added to a equal volume of a solution containing biphenyl-4-thiol (100 µM) in buffer (0.1 M, Citrate buffer = pH 3.0-5.0, Phosphate buffer = pH 5.5-7.0, TRIS buffer = pH 7.5-9.0). The palladium reagents were sonicated for 10 s to ensure the reagents were fully dissolved.

General Halide Study Procedure -(Halide Effect Experiments)
A

General Peptide-Peptide Ligation Procedure
A solution of the palladium-peptide reagent (1D) in water (40 M) was added to a solution of a peptide containing a single cysteine (20 M) in TRIS buffer (0.1 M, pH 7.5). Note: if the palladium reagent was not readily soluble in H2O, the slurry was sonicated for 10 s to facilitate this process and become fully solubilized. Final conditions: [Pd] = 20 M; [R-SH] = 10 M; After 6 h at 37 °C, 3-mercaptopropionic acid (3 equiv to the palladium complex solution in 6 L of H2O) was added to the reaction mixture to quench the remaining palladium species. The reaction was allowed to stand for 5 min and subsequently characterized by LC-MS.
Note: In the following pages, the full chromatogram is depicted alongside the extracted MS chromatogram from the associated peaks at the indicated time interval. All data was processed with Agilent MassHunter Qualitative Analysis 10.0. Conversion was assessed by integrating the peak on the chromatogram of the desired product divided by the sum total of the integration of the peaks of the starting thiol containing peptide + any observed products derived from the starting thiol containing peptide. Calculations are illustrated. All compounds were assigned based on their agreement with calculated masses generated using ChemDraw 20.0. Peak numbers are indicated above the peaks, and mass spectra follow for each peak number. Non-product peaks derived from peptide P1 are indicated in blue, product indicated in red 997.0262 Peak numbers are indicated above the peaks, and mass spectra follow for each peak number. Non-product peaks derived from peptide P1 are indicated in blue, product indicated in red

Protein Expression and Purification
pD444 PAC-CO plasmids were constructed as previously reported. 3 Cysteine mutations were introduced by site-directed mutagenesis using QuickChange Lightning Single Site-directed Mutagenesis Kit (Agilent) following the manufacturer's instructions. Sequences of the generated protein construct are summarized in Table S2.
PA [K653C] was expressed in the periplasm of E. coli BL21(DE3) cells. Cells were grown in LB medium containing ampicillin (100 μg/mL) at 37 °C to an A600 of 0.8. Expression was induced by addition of IPTG to final concentration of 0.4 mM, and the culture was grown overnight at 18 °C. The protein was purified by anion exchange chromatography followed by size exclusion chromatography.
The protein was analyzed by LC-MS confirming sample purity and molecular weight. Protein aliquots were flash frozen and stored in -80 °C freezer.

Protein Labeling Experiments
To a 1. Note: The palladium-peptide 1B did not fully dissolve in DMSO and created a slightly foggy solution. Efforts to perform this reaction in the absence of organic solvent led to variable conversion, which we attributed to the low solubility of the palladium-peptide. As a result, we chose to dissolve the peptide in DMSO.
Reactions were analyzed using Agilent BioConfirm 10.0 and deconvoluted spectra were obtained using the maximum entropy algorithm with: a mass range of 20,000 Da to 100,000 Da; mass step of 0.5; utilizing a limited m/z range from 950.00-3000.00 Da; and with baseline subtraction of 7.0.
Chromatograms and mass spectra of PA-cys and the crude reaction mixture are on the following two pages.

PA-Cys Starting Material
A background chromatogram is overlayed and depicted in red. The protein peak from 7.5-13 min was extracted to give the following chromatogram: Deconvolution of this spectrum gave the following mass for PA-Cys:

Crude Reaction Mixture
A background chromatogram is overlayed and depicted in red. The protein peak from 7.0-12.6 min was extracted to give the following mass spectrum: Deconvolution as described above to give the following mass spectrum, with relative peak area of the region highlighted in grey written below the indicated mass.

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Note 1: When attempting the reaction without His buffer or without the use of DMSO, the reaction typically gave lower conversions and/or gave byproducts that were difficult to remove (a product whose mass corresponded to the product + Pd + sSPhos was observed. The use of DMSO was found to give reliable conversions and the use of His buffer was found to minimize the observation of this Pd + sSPhos adduct.

Note 2:
During initial attempts using His buffer for the conjugation of peptide 1B with PA-cys, a 500 mM, pH 6.1 buffer was used instead of the described 200 mM buffer used in the described protocol. In addition, 25 equivalents of the peptide 1B were used. During this reaction, conversion was high (>90%). However, the protein was observed to fragment over the course of time to give a 22.6 kDa and a 60.5 kDa fragment. Decreasing the buffer concentration to 100-200 mM and increasing the pH to 7.0 was found to avoid this issue. As a result of this observed cleavage, SDS-PAGE analysis of the reaction mixtures were obtained to corroborate LC-MS data. The following is a summary of conditions and observations, with reaction B being the above procedure:

OXIDATIVE ADDITION COMPLEX STABILITY
When stored as a solid or kept frozen in solution, the palladium-peptide oxidative addition complexes display remarkable stability for upwards of 12 months and show no diminished reactivity. This is confirmed via a set of experiments wherein the oxidative addition complexes were stored as either a frozen solution or solid and then, after 12 months, allowed to react with a small molecule thiol. The reaction conversions are comparable to that of a freshly prepared palladium-peptide oxidative addition complex.
A solution of the palladium-peptide reagent 1B (20 µM) in water was added to a equal volume of a solution containing biphenyl-4-thiol (100 µM) in TRIS buffer (0.1 M, pH 7.5). The palladium reagents were sonicated for 10 s to ensure the reagents were fully dissolved.

Stability in Solution
The three palladium-peptide oxidative addition complexes (1A, 1B, and 1C) were dissolved in 50% water / 50% acetonitrile containing 0.1% TFA at a concentration of 300 nanograms / µL. The solutions were analysed by LC-MS using method A after 1, 2, and 3 days .  For compounds 1A, 1B, and 1C, the peaks ~5.6 minutes have mass 1349.7 Da and 1374.7 Da, which corresponds with replacement of Pd+X+Ligand with either a hydrogen (-H) or nitrile group (-CN). For compound 1C, the peak ~5.7 minutes corresponds to peptide P6.

COMPLEX 1B MS/MS CHARACTERIZATION
To confirm that the palladium is located at the N-terminal 4-bromophenylalanine residue, and not elsewhere on the peptide, we subjected oxidative addition complex 1B to LC-MS/MS analysis. Method A was modified to acquire secondary MS spectra targeting specific ions. Ions with z = +2 and m/z = 973.87 or 973.37 Da were selected for secondary MS. Shown below is the secondary MS spectrum at 7.95 minutes for the 973.37 Da ion, with CID at 37.3.
The data were analysed using mMass software. The presence of palladium makes comprehensive analysis of the MS/MS data challenging due its isotopic abundance (deisotoping algorithms do not account for the multiple Pd isotopes). Therefore, a few selected ions are shown below. Critically, no y-ions exhibited an isotope pattern suggestive of palladium, as would be expected for a palladium complex on the N-terminal residue. Additionally, the b-ions exhibited the characteristic palladium isotope pattern.

General Experimental
NMR spectra were measured on a Bruker AVANCE Neo 600 MHz spectrometer equipped with a 5 mm QCI-F cryoprobe. Unless otherwise noted spectra were recorded at 25 °C. Standard COSY, TOCSY, 13 C-HSQC, 13 C-HMBC, and multiplicity-edited 13 C-HSQC spectra were recorded for resonance assignment. ROESY spectra were recorded with a mixing time of 400 ms for H-Pro-Pro-Phe(4-Br)-NH2 and 200 ms for H-Pro-Pro-Phe(4-Br)-NH2-OAC.

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The ROESY cross-peaks integration was performed with MestReNova 7.1.2, The volumes of crosspeaks were converted into corresponding distances using Eq. (1)  The distances between H  and H  located on the same side of pyrrolidine rings of proline residues were assumed to correspond to 2.4 Å. 6 The distance between two geminal protons was assumed to be equal to 1.8 Å. For H-Pro-Pro-Phe(4-Br)-NH 2 (2A) and H-Pro-Pro-Phe(4-Br)-NH 2 -OAC (2B) the averaged P 1  -P 1  1 and P 2  -P 1  1 peaks as well as P 2  1 -P 2  2 were used for calibration. If the crosspeaks from both sides of the diagonal could be integrated, they were separately converted into distances and then averaged. The volume integrals used to estimate distances scale with the sixth power of distance. As such, the error estimates are expected to be small.

General Computational Considerations
All reported DFT calculations were performed using the ORCA software 4 or Gaussian 03. 5 Images of the 3D structures were rendered using CYLView. 6 The geometry of all reactants and transition states were optimized using the B3LYP 7,8 functional in the gas phase. In these geometry optimizations, a mixed basis set of SDD 9,10 for Pd and 6-31G(d) 11,12,13 for all other atoms was used. Ground state geometries were validated by vibrational analysis at the same level. These frequencies were also used to calculate zero-point and thermal energies at 298 K using the standard rigid-rotor and harmonic approximations in Gaussian 03. Higher-level single point energies were obtained for each stationary point using the B3LYP functional with a mixed basis set of SDD for Pd and 6-311+G(d,p) for all other atoms. In these calculations, the SMD implicit solvation model 14 with water as solvent ( = 80.4) was applied. The estimated Gibbs free energies of solvation are reported as a composite of this higherlevel electronic energy (SDD/6-311+G(d,p), SMD) plus all other contributions from the lower level (SDD/6-31G(d), gas phase).

Generation of Candidate Conformations
Using the Confab utility, 15 conformations of the peptide 2A were systematically generated by a torsion driving approach. This procedure sampled ~2 million geometries, which were reduced to a set of 438 unique candidates with energies within 20 kcal/mol of the minimum energy conformer, as assessed using the MMFF94 forcefield. 16i Next, gas phase DFT optimization of this set (B3LYP/6-31G(d)/SDD(Pd)) led to a further reduction to 48 unique conformations. Assessment of the solutionphase Gibbs free energies (B3LYP/6-311+G(d,p)/SDD(Pd)/SMD(water)) revealed Conf 61 to be lowest in energy, with the remaining members spanning a roughly 12 kcal/mol range of energies above Conf 61. Notably, many of the bond distances in Conf 61 agree with ROESY measurements for the peptide, shown below.

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Based on these preliminary results, we set out to perform a similar conformational search for the palladium oxidative-addition complex derived from this peptide (2B, above). We inserted SPhosPd into each of the 48 candidate conformations of peptide 2A with several different geometries for the Pd-containing fragment. Following the results in a previous theoretical study, it was important to assess both the "O-bound" and "C-bound" conformations of SPhosPd, which were similar in energy for several model compounds (within 1 kcal/mol). 17ii Furthermore, the -PCy 2 group could display several similar conformations, differing by rotation about the P-C bonds. Finally, rotation around the C(aryl)-Pd bond needed to be considered for each complex. The entire set of Pd-containing candidates was subjected to gas phase DFT optimization (B3LYP/6-31G(d)/SDD(Pd)) then ranking by approximate solution phase Gibbs free energy (B3LYP/6-311+G(d,p)/SDD(Pd)/SMD(water)). The energies of the 100 most stable conformations are displayed in Table S9.

Conf 162D
The first three conformers (61A, 1A, 6A) differ primarily by rotation of the C-terminal amide group and are very similar in energy. Thus, we assigned Conf 61A as the most probable ground state conformation, with contributions from amide rotamers Conf 6A and Conf 1A. The consistency of this assignment with ROESY data is supported by the values in Table S10.