Studies of Coupling Kinetics and Correlation of Reaction Conversions

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Studies of Coupling Kinetics and Correlation of Reaction Conversions to Color Tests for Solid Phase Peptide Synthesis of AMG 416 by NMR Tsang-Lin Hwang, Krishnakumar Ranganathan, Yuan-Qing Fang, Richard D. Crockett, Steve Osgood, and Sheng Cui Org. Process Res. Dev., Just Accepted Manuscript • DOI: 10.1021/acs.oprd.8b00177 • Publication Date (Web): 20 Jul 2018 Downloaded from http://pubs.acs.org on July 20, 2018

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Organic Process Research & Development

Studies of Coupling Kinetics and Correlation of Reaction Conversions to Color Tests for Solid Phase Peptide Synthesis of AMG 416 by NMR Tsang-Lin Hwang1,*, Krishnakumar Ranganathan2,†, Yuan-Qing Fang2,#, Richard D. Crockett2, Steve Osgood1, Sheng Cui2 1

Attribute Sciences, and 2 Drug Substance Technologies, Amgen Inc., One Amgen Center Drive, Thousand Oaks, CA 91320

ABSTRACT: By using the solution NMR technique and the “cleave and analyze” approach, we have modeled the kinetics of coupling reactions between activated Fmoc-Arg(Pbf)-OH and NH2-Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM resin, determined the reaction rates, and correlated the extent of conversions to the Kaiser and TNBS color test results. Since NMR spectroscopy is a highly specific and quantitative tool with structural elucidation capability, the NMR assay for conversion determination of reaction in solid phase peptide synthesis (SPPS) can be developed by comparing the peaks from the reactant peptide containing the free amino groups with the product peptide containing the newly coupled Fmoc-amino acid moiety in the 1D 1 H NMR spectrum. We utilized the NMR assays to measure conversion for each cycle in SPPS to assemble Ac-c(C)arrrar-NH2 and provided an in-process testing (IPT) time limit for the full-scale production. The data showed that after 18 hours of reaction, the conversion for all cycles achieved > 99.9%, demonstrating the robustness of SPPS process and ensuring control of certain impurities, such as deletion sequences, below 0.1% in the active pharmaceutical ingredient (API). KEYWORDS: solid phase peptide synthesis, NMR, kinetics modeling, reaction conversion, color test

INTRODUCTION Solid phase peptide synthesis (SPPS) has become a platform technology to manufacture peptide based drug substances ever since the discovery of peptide chain elongation on resin by Merrifield in 19631-4. In SPPS, beads of resin5 serve as macroscopic sized protecting groups for amino acids/peptides by anchoring their C-termini thus preventing chemistry at this site that would otherwise result in mixtures of products like those seen in solution phase peptide chemistry. The given amount of peptide that a resin can anchor (i.e. loading) is on the order of 1 mmol peptide per gram of resin. The resin beads are constructed of polymeric chains functionalized with molecular linkers to which the first amino acid of a peptide sequence is anchored. These polymeric chains are also cross-linked to prevent bead dissolution in organic solvent. An important consequence of this crosslinked structure is that > 99% of the functionalized sites (linkers) are interior to the bead. Another consequence is that bead size, bead size uniformity and bead solvation can affect diffusion of reagents into/out of the resin and can play a key role in reaction rates. Given the sheer number of beads used in a single synthesis, bead size and size uniformity become important when studying reaction rates. In the Fmoc-SPPS strategy, the resin contains NFmoc protected linkers. These linkers are deprotected under basic conditions. The resulting free amine is then coupled with an activated, Fmoc protected amino acid. Once coupled, the

Fmoc amino acid is deprotected and the coupling/deprotection process continues in an iterative fashion (i.e., in cycles) until the desired peptide sequence is obtained. Amino acid sidechain functionalities are protected with acid cleavable protecting groups6. The completion of a coupling reaction in each cycle is usually monitored by the Kaiser7 and/or TNBS8 colorimetric tests in which a “positive” Kaiser test (a dark blue solution) and a “positive” TNBS test (orange/red colored resin beads) indicate the presence of unreacted primary amines. These qualitative color tests are useful and are standard practice, but interpretation of their results is often highly subjective due to the nature of visual inspection, sample size and dilution. In fact, the TNBS test often requires a microscope to more clearly ascertain the presence of faint color in the resin beads. For positive colorimetric test results, either an acylation is performed which effectively shuts down the unreacted amines (capping) or a re-coupling is employed in which the resin is drained, washed several times and treated with fresh reagents to fully maximize yield. Both approaches add cost and processing time to the manufacturing process. Furthermore, capping adds further burden to final purification steps aimed at removing truncated sequences which can be difficult to separate. Therefore, a fundamental understanding of the reaction kinetics is imperative in terms of reaction time, yield, reagent charge, impurity profile, etc. To monitor reaction progress in SPPS, or similarly in the solid phase organic chemistry (SPOC), which also applies

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solid support to carry out chemical reactions9-10, both ‘cleave and analyze’ and ‘on-bead’ analysis methodologies have been

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developed11-13. In the ‘cleave and analyze’ approach, the reaction products are cleaved from the resin, thus simplifying

Scheme 1. Coupling of Fmoc-Arg(Pbf)OH to a peptide sequence by solid phase peptide synthesis.

the analysis of product and starting material by removing the matrix effects caused by the resin14-15. On-bead analysis can be performed by comparing intensities of specific signals from reactants and products. Examples include (1) gel-phase NMR, with particular use of 19F, 31P, or 13C for signal resolution and spectral simplification16; (2) high-resolution magic-angle spinning (HR-MAS) NMR17-19, where magic angle spinning can remove magnetic susceptibility between the solid and liquid interface and other anisotropic magnetic interactions, making the NMR peaks sharper. Nonetheless, the results depend on the characteristics of solid support and solvent used20; (3) IR and related spectroscopy, where change in the functional group of a molecule may result in the alteration of absorption band in the spectrum12; and (4) mass spectrometry to detect specific peptide ions for identity21. AMG 416 is an unnatural octapeptide agonist with the sequence Ac-c(C)arrrar-NH2 and acts as a calcimimetic to treat secondary hyperparathyroidism in patients with chronic kidney disease on hemodialysis22. The synthesis of the AMG 416 backbone containing seven D-amino acids was carried out using the Fmoc strategy on Rink amide-AM resin. During the manufacturing of AMG 416, the coupling rate of the 5th amino acid (4th Arg from the C-terminus, cycle 5) was found to be slower than those of other coupling cycles. We were therefore particularly interested in the kinetic study of this coupling reaction to improve the conversion. Also, the carboxy activated arginine amino acid intermediate is known to form δlactam by-products23. The kinetics of this competing side reaction required understanding to improve the efficiency of the overall coupling process. Initial attempts to analyze supernatant solutions in cycle 5 by HPLC provided incorrect information due to partial hydrolysis of the activated arginine intermediate. Moreover,

the different arginine species could not be distinguished using in-situ process analytical tools such as a react-IR. Similarly, the HR-MAS NMR technique for the on-bead analysis applied to different cycles of SPPS for AMG 416 revealed signals from resin and no peptide signals could be identified in the spectra, probably because the Rink amide-AM resin does not have a flexible arm. Therefore, in this study we employed solution NMR to monitor arginine species present in the supernatant and the traditional “cleave and analyze” approach to observe the peptide buildup on the resin at different time points. The advantages of using solution NMR to monitor reactions include: (1) peaks from different components in the reaction mixture are resolved with high specificity in the spectrum; (2) peak integrals directly correspond to relative molar concentrations of components in the sample allowing easy quantitation; (3) peak identification can be performed by acquiring and elucidating 1D and 2D NMR data; (4) cryoprobes can be used to increase sensitivity and reduce experimental time when compared to room temperature probes and HR-MAS probes24-25; and (5) sample and NMR experimental conditions can be adjusted according to the intended sensitivity requirement and use of the assay. In this paper, we wish to report the application of solution NMR and the “cleave and analyze” approach to study the arginine coupling kinetics, correlate the kinetic data to color test results, and apply the methodology to other coupling cycles of AMG 416 to better define reaction time for each coupling cycle during the production runs. RESULTS and DISCUSSION Scheme 1 shows the generalized coupling of FmocD-Arg(Pbf)OH (abbreviated as Arg-OH) to a free amine. The Arg-OH is first activated by DIC (N,N′-


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Diisopropylcarbodiimide)26 and Oxyma (ethyl 2-cyano-2(hydroxyimino)acetate)27 in the solution (Figure S1), and then the activated Arg-Oxyma intermediate can either form Arglactam (Figure S2) as a side product or couple to free-amino peptide resin to form the desired peptide product.

To better understand the formation rate of Arglactam from Arg-OH with DIC and Oxyma, we first conducted the experiment without resin and without agitation in an NMR tube.

Figure 1. 1D 1H NMR spectra collected from supernatant of Arg-OH coupling reaction with NH2-Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM Resin in SPPS as reaction progresses. Kinetic Simulation versus Experimental Data 0.1 0.09

Concentration of Species (M)

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0.08 0.07 0.06

[Arg-OH] [Arg-oxyma]

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[Lact am] uptake by resin coupling

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[Arg-OH] siculated simulatedcurve curve

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[Arg-oxyma] simulated curve [lactam] simulated curve

0.02 [resin uptake] simulated curve

0.01 0 0

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Time (min)

Figure 2. Kinetic modeling of experimental data for Arg-OH coupling with NH2-Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM Resin in SPPS, including the consumption of Arg-OH and DIC, and formation of Arg-Oxyma, Arg-Lactam and Arg-Oxyma uptake by the resin for product formation40.



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Figure 3. Measurements of conversion for Arg-OH coupling with NH2-Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM Resin in SPPS for the 1 to 6 hours of reaction. After microcleavage, NH2-Arg-Arg-Ala-Arg-NH2 serves as the initial reactant, and Fmoc-Arg-Arg-ArgAla-Arg-NH2 is the final product.

The data showed that 13.4% of Arg-OH converted to ArgLactam in the first hour, and in 24 hours the Arg-OH concertation reached zero (Figures S3 and S4). This result suggested that the maximum reaction time for coupling should not be more than one day for activated arginine. Next, we performed the coupling reaction between Arg-OH and NH2Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM Resin under vigorous agitation to mimic the manufacturing process. To avoid the interference of broad peaks from resin, we only measured different arginine species in the supernatant as a function of reaction time. The δ proton peaks of Arg-OH, ArgOxyma and Arg-lactam can be differentiated at 3.259, 3.282, and 4.203 ppm, respectively, in the 1D 1H proton spectra (Figure 1). Experimentally, Arg-DIC was formed and rapidly consumed to produce Arg-Oxyma, as evident by the correspondence between the decrease of Arg-OH peak and the increase of DIC-Urea methyl peak at 1.067 ppm (data not shown). Since the molar concentrations of different species are proportional to their respectively normalized NMR peak integrals, the Arg-Oxyma uptake by resin was calculated by taking the difference between the initial Arg-OH concentration (0.094M) and all Arg species still present in solution (Figure 2). We then modeled the data using reaction progress kinetic analysis28. The results showed that (1) the consumption of Arg-OH is a second order decay with respect to the concentrations of Arg-OH and DIC (Figure S5); (2) the concentration of Arg-Oxyma plateaued around 2 hours, and gradually decreased over the reaction time; (3) the rate of ArgLactam formation has a first order dependence on the concentration of Arg-Oxyma (The rate constant was extrapolated from initial Arg-OH to Arg-Lactam kinetics experiment in the absence of resin); and (4) the on-resin coupling rate could be approximately described as a function of available peptide NH2 groups, independent of Arg-Oxyma concentration (Figures S6 and S7). The coupling rate depends on many factors, such as the reactivity of available active sites, the mass transfer and diffusion of reactants to the active sites within the resin beads29. Further kinetic analysis showed that for the first 3.5 hours, the rate of Arg-Oxyma uptake by resin was greater than the rate of Arg-Lactam formation. After this time point, the rate of Arg-Lactam formation dominated.

Therefore, the key to understand reaction completion for desired product was to study the tail kinetics of the reaction, which requires the analysis of product formation on resin. The conversion of resin-bound free-amine peptide to product peptide at different time points can be monitored by NMR18, 30-32 coupled with the “cleave and analyze” approach for the peptides on resin14, 33. To help perform conversion studies, NMR reference standards of the “4-mer” peptide NH2Arg-Arg-Ala-Arg-NH2 (starting material) and the 5-mer, Fmoc-Arg-Arg-Arg-Ala-Arg-NH2 (final product) were obtained by cleaving small portions (micro-cleavage) of previously prepared resins. Figure 3 shows the spectra of the deprotected peptide NH2-Arg(4)-Arg(5)-Ala(6)-Arg-NH2 with its backbone protons (Arg534: 8.553 ppm, Ala6: 8.192 ppm) and Hα proton of Arg4 (3.852 ppm) as unique compared to the Hα proton from Arg3 (4.027 ppm) of the Fmoc-protected counterpart (Fmoc-Arg(3)-Arg-Arg-Ala-Arg-NH2). The micro-cleaved peptide sample obtained from 1hour reaction time contained a mixture of NH2-Arg-Arg-AlaArg-NH2 and Fmoc-Arg-Arg-Arg-Ala-Arg-NH2, and their relative amounts were determined by comparing integrals of the aforementioned unique proton peaks. Although we were concerned that the intensity of backbone amide proton peak may change due to exchange with residual water in the sample dissolved in DMSO-d6, the ratios of the peptides turned out to be the same by using integrals either from the backbone amide proton peaks or from the Hα peaks. Therefore, we could safely use the integrals of backbone amide proton peaks for ratio calculations. The 1D 1H spectra of micro-cleaved peptide samples showed that from 1 hour to 6 hours, the percentage of un-reacted free amino peptide dropped from 25.8% to 0.6%. By considering the resin loading, mass balance of Arg-OH was achieved by adding concentrations of Arg related species in supernatant and in the product together. As shown in this case, the NMR assay for conversion measurement can be developed by comparing integrals from the unique peaks of reactant and product peptides. Similar quantitation for free amino reactant peptide and Fmoc product peptide would be less straight forward by using an UV or MS detector in the HPLC analysis because the extinction coefficients and ionization efficiencies for these two peptides are quite different 35-36.


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Based on these data, we have performed other experiments to verify the kinetic modeling. All these experiments contained 1 equiv. of NH2-Arg(Pbf)-Arg(Pbf)Ala-Arg(Pbf)-Rink amide-AM Resin and 2.3 equiv. of Oxyma, with other various conditions, including (1) with 1.5 equiv. of Arg-OH present, initially charging 1 equiv. of DIC followed by portion wise addition of 0.25 equiv. of DIC each at 3.4, 4.1, 4.8, and 5.4 hours, and 0.5 equiv. of DIC each at 6.7 and 7.3 hours; (2) with 2. equiv. of DIC present, initially charging 1 equiv. of Arg-OH followed by portion wise addition of up to 0.5 equiv. of Arg-OH (0.1 equiv. each at 3.2, 3.9, 4.6, 5.3 and 6.0 hours), and 1 equiv. of DIC at 0.5 hour; and (3) with 1.5 equiv. of Arg-OH and 2 equiv. of DIC initially present, charging 1 additional equiv. DIC at 0.5 hour, and another 1 equiv. of excess DIC at 3 hours. The first two approaches for slow addition of DIC or Arg-OH were tried to control the Arg-Lactam formation which depends on the concentration of Arg-Oxyma in solution, while Arg-Oxyma, in turn, forms after the initial reaction of Arg-OH and DIC. The delayed addition of DIC or Arg-OH not only slowed down the formation of activated Arg-Oxyma, but also the product formation. The third approach did not have the effect of quickly generating Arg-Oxyma from the remaining small amount of Arg-OH. Therefore, the original reagent charges used in the manufacturing process seemed optimal, i.e., 1.5 equiv. Arg-OH, 2.3 equiv. Oxyma and 2.0 equiv. DIC followed by an additional 1.0 equiv. DIC after 30 minutes of reaction. Additional experimental results demonstrated that

vigorous agitation is needed to generate fast enough coupling rate to have the coupling reaction completed in an overnight run. After determining the percentage of un-reacted free amino peptide for the reaction shown in Figure 3, we performed the Kaiser and TNBS colorimetric tests on the resin beads at the same time points as the microcleavages had been performed. Figure 4 shows that the Kaiser color test was positive (dark blue solution) for conversions up to 92.3% at 2 hours. As the reaction progressed toward 99.4% conversion at 6 hours, the solution changed from blue to yellowish to colorless. Similarly, for the TNBS test, the amount of orange color in the resin beads decreased when the percentage of primary un-reacted amino groups was reduced. Figure 5 shows the orange color of the beads was largely gone at 99.4% conversion. The kinetic data presented for the Arg coupling step were obtained from the lab-scale experiments. We were interested to understand how this data would translate to fullscale production runs. Figure 6 shows 1D 1H spectra of peptide samples obtained from microcleavage of 4- and 18hour samples from the same SPPS step (cycle 5 coupling) in the full-scale production. In the 4-hour sample, the two amide peaks at 8.553 and 8.192 ppm in free amino reactant peptide (top trace) shifted to 8.536 and 8.188 ppm, respectively, and diminished in the 18-hour spectrum. This indicated that the reaction was

Figure 4. Correlation of conversions from 1 to 6 hours of reaction shown in Figure 3 to Kaiser colorimetric tests.



Figure 5. Correlation of conversions from 1 to 6 hours of reaction shown in Figure 3 to TNBS colorimetric tests.

completed within 18 hours. Furthermore, in the spectrum of the 4-hour sample, the peak at 8.536 ppm had an integral value of 0.77 with respect to the normalized peak integral of single Arg proton as 100. Therefore, the reaction conversion was 99.2%. The signal to noise (S/N) calculation for this peak is 40, indicating that the limit of quantitation (LOQ) for unreacted free peptide in the Fmoc-Arg-Arg-Arg-Ala-ArgNH2 product is 0.19 % (S/N=10). In the lab-scale data, the conversion of same reaction was 98.6 % in 4 hours and 99.1 % in 5 hours. Comparison of these data suggests that (1) agitation in the full-scale production is at least as efficient as the lab-scale experiment, and (2) the lab-scale kinetic modeling is applicable to the full-scale production in this case study. Kaiser color tests show that for the 4-hour resin sample, some blue beads were observed, and for the 18-hour resin sample, the beads did not have the blue color (Figure S8). We further considered the sensitivity of the 1D 1H NMR assay for conversion determination by examining the NMR spectra of peptides obtained from the last cysteine addition cycle of SPPS (Cycle 7) for AMG 416. When making the same concentration of samples from each cycle (~ 30

Figure 6. Measurements of conversion of Arg-OH coupling reaction at 4 and 18 hours in cycle 5 for the full-scale production of SPPS for AMG 416.


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Figure 7. Measurements of conversion of last cysteine coupling reaction at 3 and 18 hours for the full-scale production of SPPS for AMG 416.

mg/ml in DMSO-d6), the micro-cleaved peptides in the last cycle would have the least amount of material in terms of molarity due to the highest molecular weight of peptides in the last SPPS cycle when compared to those in the previous cycles. Less molar material in the sample leads to lower sensitivity for NMR signals. Figure 7 shows the 1D 1H NMR spectra of free amino NH2-Ala-Arg-Arg-Arg-Ala-Arg-NH2 and Fmoc-Cys-Ala-Arg-Arg-Arg-Ala-Arg-NH2 at reaction time 3 and 18 hours from the production run. The amide peak at 8.509 ppm in free amino peptide (top trace) shifted to 8.537 ppm in the spectrum otained from the 3-hour reaction sample of Fmoc-Cys-Ala-Arg-Arg-Arg-Ala-Arg-NH2, and diminished in the corresponding 18-hour spectrum, indicating that the reaction was completed within 18 hours. The shift of amide proton peak in the 3-hour spectrum was confirmed by 2D data analysis. 2D 1H-15N HSQC spectra, considered as fingerprinting for peptides and proteins37, showed the amide peaks at 8.509 ppm of NH2-Ala-Arg-Arg-Arg-Ala-Arg-NH2 and at 8.537 ppm of Fmoc-Cys-Ala-Arg-Arg-Arg-Ala-ArgNH2 have close nitrogen chemical shift at 118.6 and 118.4 ppm, respectively (Figure S9). In addition, 2D 1H-1H TOCSY spectra showed both amide peaks connect to Hα, Hβ, Hγ and Hδ for the arginine spin system (Figure S10). Further structural elucidation indicated this amide peak belonged to Arg3 in NH2-Ala-Arg(3)-Arg-Arg-Ala-Arg-NH2. Although detailed analysis of Fmoc-Cys-Ala-Arg-Arg-Arg-Ala-ArgNH2 peptide showed that about half of the Cys still has the trityl (Trt) protection group on the peptide after the microcleavage process, the peaks from Fmoc-Cys-Ala-ArgArg-Arg-Ala-Arg-NH2 do not overlap with the amide proton peak of Arg3 in NH2-Ala-Arg-Arg-Arg-Ala-Arg-NH2, thus not affecting the conversion analysis. The amide peak at 8.537 ppm has the peak integral about 0.69 with respect to the normalized peak integral of single Arg proton as 100. Therefore, the conversion was at 99.3% in 3 hours. The S/N calculation for the peak at 8.537 ppm is 20. The quantitation of unreacted free peptide in the Fmoc-peptide product sample for cycle 7 is about 0.35% as LOQ (S/N=10) and 0.1% for limit of detection (LOD, S/N=3). For simplicity, we define reaction completion as > 99.9 % for all cycles of SPPS for AMG 416 using the NMR assay.

We then applied this approach to analyze conversion for all SPPS cycles to produce AMG 416 at the full-scale production. The data indicated that all cycles have > 99.2% conversion after 4 hours of reaction, and > 99.9% conversion after 18 hours (overnight) of reaction. As widely known in the industry, the quality of resin is important for peptide production in SPPS. In one set of micro-cleaved samples obtained after 18 hours of reaction for all the cycles, no peak from free amino peptide and other amine related impurities was observed. However, in another set of data, spectra of some cycles show small amine, amide or aromatic peaks appeared near the unique amide proton peak of the free amino peptide in the 1D 1H peptide spectrum, and required further analysis on 2D NMR data to differentiate whether these peaks were from free amino peptides as starting material or from impurities. For this set of samples, additional purification efforts were needed in the process to meet the in-process control criteria, indicating that the resin quality could vary and affect the impurity profiles. Nonetheless, we can apply three NMR techniques to differentiate if these small peaks were originated from impurities: (1) in 2D 1H-15N HSQC data, the proton to nitrogen correlation will show different nitrogen chemical shift than that from the reactant free amino peptide; (2) in 2D 1H-1H TOCSY data, the proton spin system will be different than that from the reactant free amino peptide; and/or (3) NMR diffusion experiment can distinguish whether these peaks are originated from small molecules or not, where peaks from small molecules diminish in the diffusion spectra. We found all these small amine, amide or aromatic peaks are from small-molecule impurities, which do not possess amino-acid type of spin systems. These cycles of peptides that had amine related impurities were also subject to the capping procedure due to positive color test results. Therefore, corroborating NMR data can also be obtained from the acetylated peaks of capped peptides. That is, if free amino peptide as starting material did exist in the sample, the acetylation procedure should generate the peptides that have the N-terminal acetyl methyl peak around 1.867 ppm. Since the methyl peak is a sharp singlet from three protons (compared to broader amide peak as doublet from one proton), 0.01 molar% difference in the peak area with respect to the final product can be detected in the 1D 1H spectrum. After microcleavage of resin-bound peptides that have gone through the capping procedure, we performed the NMR assay and looked for peaks in the 1D 1H spectra that were around 1.867 ppm for signal intensity increase compared to those spectra obtained from peptides before the capping procedure. The increases of methyl signal intensities were about 0.03 molar% for peak at 1.857 ppm for cycle 2, 0.01 molar% for peak at 1.867 ppm for cycle 3, and no signal intensity increase for peak at 1.867 ppm for cycles 4 and 6. Therefore, we can infer that if free amino reactant peptide was indeed in the sample, the amount was less than 0.1 % in the SPPS cycles, consistent with the results using the amide proton peaks for the conversion study. Overall, all these data from the conversion study give us confidence on the coupling efficiencies (> 99.9% conversion) for different cycles of SPPS in the full-scale AMG 416 production, and ensure that certain impurities, such as deletion sequences38, is less than 0.1% in API. Robust manufacturing process to control impurities is necessary to warrant the quality of API for patient safety, which is a regulatory requirement39.



CONCLUSION In this study, we have shown that the combination of solution NMR and the “cleave and analyze” approach is a powerful method for studying the kinetics and reaction conversions in SPPS. Since NMR spectroscopy is a highly specific and quantitative tool with structural elucidation capability, the NMR assay for conversion determination can be developed by comparing the unique peaks of reactant peptide with free amino group and the product peptide containing newly coupled Fmoc-amino acid in the 1D 1H NMR spectrum. In contrast, color test result is semiquantitative and subjective for determination of reaction completion. Furthermore, NMR data collected for reaction monitoring can be directly imported for kinetics modeling. For the case study of Arg-OH coupling to NH2-Arg(Pbf)Arg(Pbf)-Ala-Arg(Pbf)-Rink Amide-AM Resin in SPPS, the lab-scale conversion data were consistent with those for the full-scale production data, establishing the scale-independent modeling. The approach demonstrated here can be widely utilized to augment the understanding of the kinetics of peptide coupling reactions using the SPPS platform, and thus to better define the manufacturing strategy. EXPERIMENTAL SECTION 99.96% deuterated DMF-d7 and DMSO-d6 were obtained from CIL (Cambridge Isotope Laboratories, Tewksbury, MA, USA). All the samples were placed into 5 mm NMR tubes for analysis. The NMR spectra were acquired on a Bruker AVANCE III 600 MHz NMR spectrometer equipped with a 5 mm TCI cryoprobe at 20°C (Bruker BioSpin Corporation, Billerica, MA, USA). The 1D 1H NMR assay applies 90 degree pulse with 3-second pre-saturation on the water signal, 30 seconds for the interpulse delay, and 16 scans with 4 dummy scans for data acquisition. Relevant 1D and 2D NMR experiments for structural elucidation purposes in this study are in the Bruker TopSpin software. 1D NMR spectra for kinetic modeling were analyzed by using the MestReNova software (Mesrelab Research, S.L., Spain). Different cycles of AMG 416 peptides (intermediates with different peptide lengths produced by solid phase peptide synthesis) on Rink amide-AM resins were obtained from Bachem (Bubendorf, Switzerland). The side chains of FmocCys and Fmoc-Arg were protected by the Trt and 2,2,4,6,7pentamethyldihydrobenzofuran-5-sulfonyl (Pbf) groups, respectively. DIC, triisopropylsilane (TIPS), piperidine, trifluoroacetic acid (TFA) and Oxyma were procured from Sigma-Aldrich (St. Louis, MO, USA). 4.1 Deprotection of the Fmoc group from Fmoc-Arg(Pbf)Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM resin. 50 grams of Fmoc-Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM resin were washed with 2 x 20% piperidine/DMF (100 mL, 60 min) to deprotect the Fmoc group. The resin was then washed with DMF (100 mL, 30 min), IPA (100 mL, 15 min), and DMF (75 mL, 15min; 75 mL, 30 min, 100 mL, 30 min). The resin was dried under N2 overnight. 4.2 Kinetic modeling and colorimetric tests for synthesis of Fmoc-Arg(Pbf)-Arg(Pbf)-Arg(Pbf)-Ala-Arg(Pbf)-Rink amide-AM resin. The first sample was made by mixing Fmoc-Arg-Pbf-OH (0.789 mmol, 1.0 equiv.), and Oxyma (1.5 equiv.) in DMF-d7 (3.7 mL), and DIC (2 equiv.) was

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subsequently added to the solution. 0.7 mL of the mixed solution was transferred to a 5 mm NMR tube for analysis. 1D 1 H spectra were taken at every 15 minutes for the first hour, and then every 1 hour for the next 17 hours (for Figure S3). In another experiment, resin from 4.1 (1.32 mmol, 1.0 equiv.), Fmoc-Arg-Pbf-OH (1.5 equiv.) and Oxyma (2.3 equiv.) in DMF-d7 (18 mL) was mixed first in a reaction vessel, and then DIC (2 equiv. + 1 equiv. at 30 minutes later) was added to the solution. The reaction vessel was agitated vigorously. 0.6 mL of the supernatant was taken out of the solution for NMR measurement at different reaction times for the first 7.5 hours. Before taking the supernatant solution, the reaction vessel was stopped. After taking supernatant into an NMR tube, the previous supernatant, finished with the NMR measurement was put back into the reaction vessel to keep the volume constant, and agitation of reaction vessel restarted (for Figures 1 and 2). The third experiment used resin from 4.1 (3.44 mmol, 1 equiv.), Fmoc-Arg-Pbf-OH (1.5 equiv.), Oxyma (2.3 equiv.) and DIC (2 equiv. + 1 equiv. at 30 minutes later) in DMF (32.8 mL). Small portion of resin were taken out for micro-cleavage and color tests every hour for the first 6 hours (for Figures 3 to 5). 4.3 Micro-cleavage of peptide from resin along with side chain deprotection for the conversion study. Resins produced from different SPPS cycles of AMG 416 were treated by two different procedures: 4.3.1. free-amino peptide as standard: About 0.5 g of resin from different cycles was washed with 8.6 mL of cleavage cocktail (20% piperidine/DMF by volume) for 30 min twice. The resin was washed with 1 x 8.6 mL DMF and 2 x 10 mL acetonitrile then dried under N2 for 5 mins. Added 4 mL of cleavage cocktail and put on RotoShake Genie overnight. The solution was then filtered and the resin washed with acetonitrile. Concentrate the filtrate. The residue was treated with IPE and aged for 1h and filtered through a fine frit funnel. The precipitate dried for 30 min under N2. 4.3.2. Fmoc-peptide as standard and at different reaction time points: About 0.3 g of resin from different cycles and 4 mL of cleavage cocktail (96.9% TFA, 2.6% water and 0.5% TIPS by volume) were shaked for 18 hours at room temperature. The solution was then filtered and the resin washed with 5.0 mL TFA. The filtrate was subjected to concentration under reduced pressure. The residue was treated with 20 mL diethyl ether and aged for 2h and filtered through a fine frit funnel. The precipitate was filtered and washed with diethyl ether and dried under vacuum for 12h. Free-amino peptides from 4.3.1 or Fmoc-peptides from 4.3.2 were used to make approximately 30 mg/ml samples in DMSO-d6 for the conversion study by NMR. ASSOCIATED CONTENT Supporting Information Additional NMR spectra and fitting for the kinetic modeling. The Supporting Information is available free of charge on the ACS Publications website. AUTHOR INFORMATION Corresponding Author * [email protected] ORCID


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Tsang-Lin Hwang: 0000-0001-9897-5751 Yuan-Qing Fang: 0000-0002-8952-1018 Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT The authors thank helpful discussions with Drs. Aleksander Swietlow, Peter Zhou, and Asher Lower at Amgen, and Amy Freund at Bruker. Present Addresses † Celgene Inc., Summit, NJ 07901 # Snapdragon Chemistry Inc., Cambridge, MA 02140

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(35) Einarsson, S.; Josefsson, B.; Lagerkvist, S. Determination of Amino Acids with 9-Fluorenylmethyl Chloroformate and ReversedPhase High-Performance Liquid Chromatography, J. Chromatogr. 1983, 282, 609-618. (36) Gartenmann, K.; Kochhar, S. Short-chain Peptide Analysis by High-Performance Liquid Chromatography Coupled to Electrospray Ionization Mass Spectrometer After Derivatization with 9Fluorenylmethyl Chloroformate, J. Agric. Food Chem. 1999, 47, 5068-5071. (37) Cavanagh, J.; Fairbrother, W. J.; Palmer, A.G., III.; Skelton, N. J.; Rance, M., Protein NMR spectroscopy: Principles and Practice. 2nd ed.; Elsevier: San Diego, 2007 (38) Eggen, I.; Gregg, B.; Verlander, M.; Swietlow, A.; Rode, H.; Szajek, A. Control Strategies for Synthetic Therapeutic Peptide APIs Part III: Manufacturing Process Considerations, Pharm. Technol. 2014, 38(5). (39) International Conference on Harmonisation of Technical Requirements for Registration of Pharmaceuticals for Human Use, ICH Harmonized Tripartite Guideline, Impurities in New Drug Substances, Q3A(R2), Geneva, Switzerland, 2006. (40) For the kinetic modeling, we assumed the solution volume to be 21 mL for the initial Arg-OH concentration at 0.094 M. This model fits well under the condition [Arg-Oxyma] ≥ 0.01 M.

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Studies of Coupling Kinetics and Correlation of Reaction Conversions

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