Disulfide rich peptides – in which order should the disulfide bonds be formed during on-resin oxidation?

Disulfide rich peptides are being identified in species of both plants and animals at increasing rates. As new molecules are discovered and disulfide bonding patterns characterized, the need for simplified chemical synthesis strategies is also increasing.

I have previously written about optimizing removal of several orthogonal side chain protecting groups including allyl, alloc, ivDde and acetamidomethyl (Acm) groups. The question that I’ll address today, though, is does the order in which the disulfide bonds are formed matter for cleaning up reactions to produce chemically synthesized disulfide rich peptides?

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Optimizing the removal of an Acm protecting group

Disulfide rich peptides have gained significant attention recently due to their incredible biological stability and tolerance to epitope grafting.  This class of peptides is often folded in solution, assuming the desired disulfide bond pattern correlates with the most thermodynamically stable structure.  Sometimes though, especially for chemically synthesized cysteine rich peptides, this is not the case.  The result is a complex mixture of peptides with varying disulfide bonding patterns and identical mass.

Using pairs of cysteine residues with matched orthogonal side chain protecting groups during chemical synthesis allows for precise regioselective control of the disulfide bond pattern on-resin, simplifying final purification steps.  In today’s post, I’ll explore conditions for removing acetamidomethyl (Acm) protecting groups with simultaneous disulfide bond formation.

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How to choose the right resin functionality for solid phase peptide synthesis

As a chemist new to the peptide community, there are many choices that have to be made.  Which coupling reagents to use? Heat or no heat to promote chemistry? And most importantly, which resin?  I have talked previously about resin choices, from loading levels to swelling capacity and how they affect the synthesis outcome.  But I haven’t addressed yet a fundamental feature of commercially available resins, and that’s the functional handle to which the peptide chain is conjugated.

In today’s post, I’ll describe some, and I mean only some, of the most commonly used chemical functionalities for Fmoc-based solid phase peptide synthesis and some scenarios in which you would choose one resin type over another.

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How to use a scouting column for your peptide purification

Whenever I synthesized a new peptide, I always ran a “scout run” – a small scale injection, usually with an analytical HPLC column – to both get an idea of the crude purity and also to identify a shorter, more optimal gradient for the actual purification.  This strategy is still probably fine when you want to use reversed phase flash chromatography for your purification strategy, but is there a better way?

In today’s post, I’ll discuss using a scouting column to screen gradient conditions prior to peptide purification with reversed phase flash chromatography.

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How to load the first amino acid onto Wang resin

While resins loaded with the natural 20 amino acids are commercially available these days, there may be times when loading the first amino acid onto the resin in house may be necessary.  And unlike loading the first amino acid onto amide-leaving resins, the first coupling reaction for C-terminal acids can be chemically more challenging.

There are several protocols published both in the literature as well as in technical notes from many peptide reagent and instrument suppliers, but they typically occur at room temperature over extended periods of time (3-24 hours and repeated).  In today’s post, I’ll evaluate several conditions suitable for efficiently loading the first amino acid onto Wang-type resin. Continue reading How to load the first amino acid onto Wang resin

How to choose an ion pairing agent to improve your peptide purification

Ion pairing agents are used in a variety of strategies to improve overall purification efficiency. In a previous post, I utilized ion pairing agents to increase the peptide’s hydrophobicity, improving retention by the stationary phase and enabling purification.  But what other strategies can be improved by using ion pairing agents?

In this post, I’ll utilize ion pairing agents to enable rapid peptide purification by flash chromatography.  The use of ion pairing agents can in fact alter the peptide’s apparent hydrophobicity sufficiently that the desired peptide and it’s closely eluting impurities can be resolved.  The question is, which one to choose?

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How does methanol as a mobile phase solvent impact peptide purification by reversed-phase flash chromatography?

Recently there has been substantial motivation to consider and evaluate alternative, more environmentally friendly solvents.  Some countries have even gone so far as to ban some of the more toxic, yet commonly used solvents.  In addition to general toxicity, additional consideration in the green chemistry movement is the volume of solvent used in any particular application.  In this regard, purification solvent selection is closely monitored as they are often used in large quantities.

One alternative that is growing in popularity is the use of methanol in place of acetonitrile for reversed phase purification of small molecules.  Methanol is certainly less expensive, but is also a more environmentally-friendly solvent for use in purification applications.  But it’s use for peptide purification has not been widely adopted to date.  In today’s post, I’ll compare the purification efficiency of methanol when compared to acetonitrile for peptide purification by reversed phase flash chromatography.

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How to purify peptides using a step gradient in flash column chromatography

Flash chromatography can be a challenging technique for peptide purification due to the lower resolution achieved with large particles.  While some may see this as a disadvantage, the significantly greater loading capacity gives me reason to make this work. So how can I achieve the high purity levels often accessed using traditional reversed-phase HPLC methods?

In this post, I’ll discuss using a step gradient for peptide purification.  Step gradients are commonly used in normal-phase small molecule purification and typically improve the purification efficiency while reducing the overall purification time. Continue reading How to purify peptides using a step gradient in flash column chromatography

How long should I let my cleavage reaction stir at room temperature?

As the rules for cell permeability continue to be elucidated, peptides are increasingly being used to deliver either themselves or cargo to the cell’s interior.  One thing is clear, increasing the overall cationic charge of the peptide enhances it’s delivery to not only the cytoplasm, but also the nucleus or other subcellular compartments.  To achieve the positive charge, large numbers of arginine residues are most often incorporated into the peptide sequence.

This begs the question though, should I change my cleavage protocol?  In today’s post, I’ll evaluate several lengths of time used to cleave and fully deprotect an Arg-rich peptide sequence. Continue reading How long should I let my cleavage reaction stir at room temperature?

How does flow rate affect my peptide purification efficiency when using a small pore stationary phase?

In a previous post, I evaluated how flow rate can impact my purification efficiency using flash chromatography.  I noticed though, at high flow rates a significantly later elution time for my peptide.  I hypothesized that the increased pressure was driving the compound further into the pores, increasing the overall interaction with the stationary phase and causing the increased retention.  We know that the particle size and particle pore size impact resolution and purification efficiency, so how does flow rate play a role with a different stationary phase?

In today’s post I’ll evaluate several flow rates using a reversed phase stationary phase material with slightly larger diameter particles that possess significantly smaller pores.  The smaller pores should limit the access of the peptides to the stationary phase and negatively impact the purification.

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