In a previous post, I evaluated how flow rate can impact my purification efficiency using flash chromatography. I noticed though, that my peptide eluted significantly later with high mobile phase flow rates. I hypothesized that the increased pressure (caused by higher flow rates) 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?
Covalent stapling strategies that stabilize a particular secondary structure have garnered much attention as interest in peptide therapeutics continues to grow. One such strategy - using olefin-bearing unnatural amino acids covalently bonded using ring-closing metathesis - has been exploited to the greatest extent thusfar.
In today's post, I'll discuss some strategies to overcome DMF poisoning of the Grubbs catalyst used during the metathesis reaction towards fully automating the synthesis and secondary chemistry required for stapled peptides.
Hydrocarbon stapling as a strategy to stabilize secondary structures of peptides, while introduced by Miller, Blackwell and Grubbs in the mid 1990s, really grew to the forefront with seminal work by Schaffmeister and Verdine in early 2000s. Protocols have been developed that enable this post-synthesis modification while the peptide is still on resin, but often these metathesis reactions are performed manually, and at room temperature.
In today's post, I'll compare several different sets of reaction conditions using microwave heating with the goal of expediting the olefin metathesis reaction, without compromising reaction efficiency, and towards automating the entire synthesis.
There are many techniques available to analyze and identify synthetic compounds that we are taught in the first few years of our chemistry education. While tools like NMR spectroscopy, IR spectroscopy, and others, are extremely useful for determining or confirming the structure of synthetic small molecules, these strategies are not as well suited for quick characterization of peptides. As a result, peptide chemists rely heavily on peak shape observed during a liquid chromatography step and mass spectrometry for mass confirmation.
In today's post, I'll discuss several of the mass spectrometry techniques that are used for analyzing crude or purified peptide samples.
More and more we are seeing groups that would historically undertake only traditional organic chemistry or possibly biochemistry/biology, incorporate peptides into their research programs. While this is good for expanding the application scope and diversity in the peptide space, bringing synthesis in house can be a daunting undertaking.
In today's post, I'll talk about what it takes to get a peptide synthesis operation up and running, with a few considerations along the way.
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.
Resins for solid phase peptide synthesis can vary significantly in both functionalization and composition, leading to mixed results at the end of a synthesis. Previously, I demonstrated how the resin loading level affects the success or failure of your peptide synthesis.
In today’s post, I’ll highlight how both the hydrophilicity and swelling capacity of your resin can influence your peptide synthesis.
Mass-directed purification, whether with a preparative HPLC or a bench-top flash system, is quickly gaining interest in the peptide purification space. The simple fact is that using a specific mass, rather that UV absorbance, to trigger fraction collection allows for greater confidence in the identity of the collected fraction. Importantly though, this technique can also reduce your time required for purification, by significantly reducing or even eliminating the need for secondary mass analysis of each collected fraction.
When it comes to synthesizing a peptide, the first thing that comes to mind is the number of stoichiometric equivalents to use. Sometimes that number is as few as 1.5, sometimes it’s as high as 20!
But have you ever thought about the liquid volume that contains those molecules and how that might affect the success of your coupling reaction? In this post I will discuss the impact of amino acid concentration in the overall success of solid phase peptide synthesis.
It used to be easy with only polystyrene based resin types, but nowadays there is a broad choice of types to choose from, including everything from the C-terminal functionality (Rink vs Wang) to the polymer from which the resin itself is synthesized.
All resins have one thing in common, and that’s the reactive site loading level. In this post, I will share my experiences with how this important factor impacts the success of peptide synthesis.
Reversed phase flash chromatography is increasingly being utilized by peptide chemists to decrease purification time and efforts. The larger particles used in flash columns enable large crude sample loads and can lead to highly pure peptide samples despite lower resolution when compared to traditional HPLC methods. However, there are some situations where the purity achieved isn't sufficient. Then what can you do?
In today's post, I'll describe using a focused gradient to achieve higher purity peptides than is possible with a more traditional linear gradient.
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.
In my role as a peptide application scientist, I have had the pleasure of working with many groups that are venturing into the world of peptides for the first time. Although it seems rather straightforward to experienced synthetic chemists, producing acceptable yield and purity certainly comes with unique challenges in solid phase peptide synthesis .
In this post I would like to present some of the tips and tricks that I have picked up along the way.
Purification by reversed-phase chromatography relies primarily on a hydrophobic interaction of the molecule with the alkyl chains bonded to the stationary phase for column retention and elution through a partitioning mechanism. While this is certainly true for purification of peptides, surface area accessibility and media particle size also play critical roles in the resolving power of a particular stationary phase. The particle size influences the loading capacity, however pore size greatly influences molecular accessibility and therefore resolving power.
In today’s post, I will demonstrate how pore size can impact your peptide purification using flash column chromatography.
In the past, when I have 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 recommended 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.
More and more groups are exploring the utility of peptides with an ever widening variety of applications. And although peptides are getting cheaper to purchase outright, many groups are continuing to bring peptide synthesis in house. As more groups join the peptide community, I frequently encounter questions about the basics of peptide synthesis.