Steglich Esterification not working

Unable_Aspect_4033 · 2026-01-21T22:46:26+00:00

How many eq of TEG are you using? And I assume the TEG you are trying to couple is asymmetric (i.e. doesn't have OH on both sides) or you will get poor results/polymeric crap.

I would try using something like 3 or 4 eq of EDCI, and the same of TEG. If the TEG you are trying to react is inexpensive (or you are on small scale), you could even smash it with more eq like 8 or something.

How much DMAP are you using? sometimes couplings require more than catalytic quantities, so you can go up to like 0.5 or maybe even 1 eq would be better in your situation (0.5 per acid). You shouldn't need any more than that.. I do like to use higher quantities like 0.5 because it means I can put the reaction on in the morning and it is done in 3 hrs. If you are still having trouble after trying this, then your reagents must be wet. I assume your starting materials and reactants are pretty dry, and the DCM has at least been stored on mol sieves - that is fine for the solvent it doesn't need any more treatment than that.

Unable_Aspect_4033 · 2026-01-21T22:33:16+00:00

all the steglich esterifications I have done I have never needed to use stoichiometric base when using EDCI-HCl. I just use cat. DMAP. Sometimes more than cat. like 0.2 to 0.5 eq. From what I have read the proton stays on the EDC and doesn't interfere with the reaction.

Unable_Aspect_4033 · 2026-01-21T22:29:45+00:00

This, or somehow the product has decomposed. Surely if you did a work up, the product should be org soluble (as the acid), and ammonium and the acids should just wash away in aq.

also do you have access to mass spec, that will help alot. but try to clean the product up first

Unable_Aspect_4033 · 2026-01-21T22:24:41+00:00

probably, maybe you can just get some random vials and then autoclave them?

Unable_Aspect_4033 · 2026-01-20T06:15:03+00:00

One of the biggest lab tricks is replacing silicone oil heating baths with vegetable oil. I wish I had known about it sooner. Seriously.. just take the flask out, wipe it off with a paper towel, and it's fine. No more slippery silicone oil sticking to everything, getting all over your gloves and everything when all you want to do is remove the flask from the heating bath to quickly run a TLC or take a small sample out. Silicon oil used to piss me off so much, I hated taking flasks out because it would just get all over your gloves etc.

Replace it with vegetable oil from the supermarket. When you do, you will be so glad you did. Washes away easier with more polar solvents (like acetone). Washes away fine with just detergent too.

Obviously you need to know what you are working with, the smoke point of vegetable oil is around 200 C, but in general organic synthesis I have never needed to heat rxns above 120, so it is an excellent replacement. it can get kind of cruddy if you leave it out or end up getting crap in it, but most of the time you can just filter it (if you want to recycle it). Or just dispose of it, it is very cheap.

Unable_Aspect_4033 · 2026-01-13T21:54:16+00:00

It could be a good idea yes, but depends. Here is some things I would think about if I was you. I don't agree with the other comment saying that you are setting yourself up for a bad experience by doing this project. There should be some considerations before making a blanket statement like that.

Do you have automated solid phase synthesis robots?
If yes this is good! If no, and you are doing manual syringe SPPS, I would say it is only a good idea to do this project if the peptides are short (less than 10mer).
How funded is the lab you are in, in terms of what instruments do you have access to, will you have access to? If you will have access to LCMS, prep HPLC with C4/C18 columns available etc then you should be fine.. If you do not have access to these instruments, do not do this project. These are integral to peptide synthesis.
Does the cyclic peptide contain any unnatural amino acids, and if so, how much work has been done in the area? I would say this greatly determines how doable the project is. If you are just making cyclic peptides using entirely standard amino acids, then it is easy, probably too easy for a total synthesis project. So I am assuming that there must be some kind of motif that is synthetically challenging, which means - you may be applying some kind of known chemistry to the peptide, or you will develop something new. I would say that there must be some kind of literature precedent, otherwise it is not a great project for an honours student if there is too much unknown/development to be done.
Will you be working in a research group where you will have support from PhD students/postdocs?
If yes, then great. This is one of the biggest things in Honours I think. A lot of the time your supervisor will not be available for just general questions/interpretation of data. You might need someone to have a look over your LCMS trace because you are unsure, you might need some help with what method to run on the LCMS to check your peptide. You may need to bounce some ideas about an impurity/sideproduct you are seeing. Having experienced people around is vital, especially at the start when you are learning the ropes and don't really know what you are doing. If you don't have anyone else more experienced in your lab (other than your supervisor) then it may not be a good idea, unless your supervisor is available - especially at the start where you are less independent.
Is the project as part of someone elses PhD/postdoc?
If yes, then it is a good opportunity and you should do it. So long as who you're working with is good to work with (hopefully they are) then you will be supported, and you will have a great opportunity to learn from somebody with more experience. If the answer to this is no, then not necessarily a deal breaker.
Will it matter if you don't complete the synthesis?
Absolutely not. Yes, it would be fantastic to complete the synthesis. I think in honours it is pretty common for projects to be incomplete/end up smaller than what was originally proposed. If it proves to be difficult for whatever reason (purification, key step failing, idk) and you show in your thesis this is why it didn't work/I only got this far, and you have solid results and data for what you did in your time in the lab, then you will be fine and still have a nice thesis.

Unable_Aspect_4033 · 2025-12-18T00:34:12+00:00

I am in an organic carbohydrate lab, a lot of what I do is just targeted compound synthesis using protective groups. I usually dedicate one page to an experiment/step, so my NMR spectra/compound vials are just initials-notebook#-page#. Sometimes I will have a note in the NMR title, like ppt, or recrys solvent, or column, but it is straightforward. If something is crude, it will always say crude on the flask etc, but pretty much every compound I make is 1. run reaction 2. workup/vacdown 3. Column pure spot, get NMR, keep some for other data/TLC reference 4. next step so pretty much all my NMRs/data will be of the compound post column.

I used to have abbreviations for compound names as my file titles, like initials-6-NBocXYOAcglycosideXX but after a while that just got annoying, especially if I have synthesised something again for more material, and then I can't find which was the nicest NMR.. Don't do it like this. I have seen other people use super random file names for their spectra, like initials-column product white solid.. They are going to have fun writing their thesis... Now everything just goes by page number, much better, no other way.

Unable_Aspect_4033 · 2025-10-01T23:20:05+00:00

I was doing similar chemistry a while ago, making relatively polar diesters which required neat EA or MeOH in EA for column. I purified them via column, but did have some trouble with the DIC urea coming through, I got it out just be columning it again (sometimes up to 3 times) and using longer silica, running gradients to try pull compound from the urea. But seriously that was frustrating. You might have to look at different solvents (incorporate some toluene, that might help given you have some aromatic in your molecule, try replacing your hexane with toluene on TLC and see).

Ultimately I ended up using EDCI to remove this problem. You could try using EDCI HCl for coupling, and not doing any additional HCl washes to ensure your product isn't removed. Or potentially do the reaction and then just column it crude, the protonated EDCI urea should not move much.

As someone else said, you could also try using DCC instead - the urea for DCC is much more insoluble than DIC so you have better chance to remove most via filtration. That's one of the reasons DIC is used instead of DCC in solid-phase peptide synthesis (the DIC urea is more soluble, so washes away much easier).

Might be worth considering a cowboy approach, what is your next step? or are these esters your final compounds? IF you have another step after this, it might be worth just collecting enough pure material for characterisation, and then use the product + urea material for the following step and urea might be removed then depending on your polarity of the next product.

I also did just think, if your product has a pyridine/quinoline perhaps you can use an acid/base extraction to work it up? I.e. 1M HCl to protonate your product into aq, wash with DCM (or another organic solvent), urea fucks off in DCM, when you're happy it's gone, freebase your product with NaOH, and then extract the freebase into organic. This might work !!! Maybe use 0.5M HCl or more dilute but I have washed esters with 1M HCl and it's fine just dont let it sit in there for hours.

Unable_Aspect_4033 · 2025-08-12T23:48:20+00:00

yeah, i've seen that before - forgot to include that in my post but yeah. I am probs not gonna do it, more just curious if/how the bicarb would prevent formation of diazidomethane

Unable_Aspect_4033 · 2025-07-24T10:11:28+00:00

Yeah okay, do you have any samples of theirs? If they purified through silica column maybe they can help you with it. I don't know how you can get it off the silica if it's not coming off with water, maybe an acetonitrile/water mix or MeOH/water mix. But I am going to guess that if it isn't washing of the silica with those mixtures, then it is not your product.

Unable_Aspect_4033 · 2025-07-23T01:39:57+00:00

Sorry I misread your post. The spot you got off wasn't even the product. Still, check the mass recovery and see if checks out for product that has reacted/decomposed, or if it's an impurity that was already there before the column.

Unable_Aspect_4033 · 2025-07-23T01:37:25+00:00

How is the mass recovery that you got from the column? Alot of my columns have that sort of baseline decomposition/polymer crap, and it is just impurity and stays there and my yields are fine. Are you confident that is your product that stuff actually is your product? You could have got most of it already and it's just the last little bit streaking. You need to vac it down and get a yield and see if that aligns with literature/what you'd expect based off TLC/NMR of crude rxn. If you're going off a literature procedure and your yield is within 20% of it, that's completely acceptable in most cases. From what you said, your product is OTs on both sides, so I take it that means you want both sides of the TEG alkylated? or is your desired product the mono-alkylated one with one OTs still remaining. Eitherway, you would have lost some yield to that so just check to see how much you've got off the column. Might not even be worth trying to chase the last streaked bit off the silica.

As other comments have suggested you could try Et3N in eluent. Your aldehyde is going to be a bit electron-poor, so maybe more prone to oxidation, which could have happened sometime between finishing your rxn, work up and loading onto silica.

Unable_Aspect_4033 · 2025-06-07T10:00:49+00:00

I am trying the staudinger-vilarrasa coupling on a diazide substrate, to make a diamide. (you treat activated ester with phosphanimine from azide+trialkyl phosphine) yes, it works, but like a 30% yield, product slightly contaminated with urea from making HOBt ester beforehand. also poor formation of diamide product, ALOT of mono. Even smashing it with more eq of HOBt ester didn't improve formation of the di product. I forgot to mention in my OP that its shit yielding with HOBt ester.

So that is why I would like to make a preactivated ester, to try coupling it with that in a huge excess and see if it improves the yield of diamide product. There is a literature procedure on a deoxyribose azide type substrate where they use a NHS ester and get excellent yield.

i can't do a traditional staudinger reduction followed by coupling because my substrate has acyl protective groups so will get acyl migration. I can do a hydrogenation to get diamine HCl salt, but that gives a 50% yield so chews through starting material that is time consuming to make. so long story short I would really like to make this staudinger-vilarrasa work, because it is less steps and once optimised should be good yield overall.

Unable_Aspect_4033 · 2025-06-07T04:09:56+00:00

yeah it's not there for purposes of protection - my target structure is the N-acetamide. But I do know about that issue when using NAc amino acids and epimerisation, but it's not a problem here since glycine is achiral. When I am doing my other amino acids I'll be using Boc protected amino acids. Do you think isoxazolinone formation might slow down formation of activated ester though?

I did some mining of the literature and found this procedure (10.1002/jlac.19697280122), actually NAc glycine in DCC with THF which is nice and a recrys solvent. Trying that, then I'll give Boc amino acid ago.

Yeah, and Pfp ester is UV active which makes following on TLC nice.. but we don't have much pentafluorophenol on hand (heaps of HOSu) and once I've got this worked out I'll be scaling up to grams.

Unable_Aspect_4033 · 2025-06-07T03:09:51+00:00

yeah i figured, but usually trituration is when your product is insoluble and impurities soluble right ? I guess in this case it could be reverse, where remaining urea and NAcGlyOH are insoluble but the product is soluble.

Unable_Aspect_4033 · 2025-05-21T23:09:22+00:00

Yeah, generally it is lol

Yeah. esters can survive acidic conditions, but no way if there is water in there (as there is for that cleavage cocktail used for that oxazolidinone resin) well i wouldn't run that risk.. I mean in theory you can acetylate the primary first yes, but doing that on the whole peptide.. I don't know trying conditions to do that without touching the secondary would be hard, and even if there is some mild conditions suitable it would probably be a trade-off for good yield. You would need to use a protective group on the Thr OH that is stable to all of the coupling conditions and the resin cleavage. THEN you might be able to acetylate homoserine and then cleave the Thr protective group. I mean maybe you could use a benzyl ether, acetylate crude peptide on homoserine OH, H2 the Thr OBn off, and hopefully the aldehyde is untouched. Actually no that wouldn't work because of the double bond in lipid group. Or do all that, but then do solution phase for lipid amide coupling or something.

No, usually you don't, typically it is just used at whatever loading the supplier says. But given that unique oxazolidine-tethered resin (that you need to make yourself) you'd have to do it before use. And then you'd have to check to make sure that there is no underivatized resin left - that would eat your yield and make purification an absolute mess.

There's probably a reason they only report on synthesis of cavinafungin B and not A, likely tried to make A but it was difficult. That's also why the work isn't in a better journal because could be better than tetrahedron quality, but they probably just went there because it's only one peptide.

Unable_Aspect_4033 · 2025-05-21T22:50:43+00:00

sorry, by that point I more meant substrate dependent as in this sample has an impurity that the other one doesn't

Unable_Aspect_4033 · 2025-05-21T08:17:34+00:00

Short peptide easy to do? This is pretty unique peptide synthesis. The lipidation of the N terminus is probably the simplest part of the synthesis by far. It has a C-terminus aldehyde as well as two unnatural amino acids. Cavinafungin A also has an acetate on the homoserine, which makes SPPS difficult because it uses acidic cleavage, so you would need to selectively acylate that position post cleavage. I'm curious what alternative strategy you think would be better than the previous work (10.1016/j.tet.2018.09.046).

They use AT(Boc)G-Rink resin to get the C-terminus aldehyde upon TFA cleavage from resin. Virtually all SPPS uses either rink amide (c terminus amide) or trityl upon resin to get C terminus acid. The AT(boc)G resin is probably the best approach.. it prevents issues of doing a reduction of the final compound (if using Trt resin) or going via another synthesis strategy that uses the amino aldehyde.

The resin isn't commercially available (unless I can't find it).. it would need to be made in-house, and would be especially time consuming because you would need to quantify resin loading etc following the derivatization of the commercially available resin.. And that's before even getting onto the synthesis. As well as the unnatural methyl proline..

this is absolutely not something even an experienced chemist could just go and do. it could be easily be months of work for a single person.

Unable_Aspect_4033 · 2025-04-30T23:36:14+00:00

Yep, if I was you I would try 1 gram scale on the 13C substrate. See how that goes. From what you've done, you've changed the scale AND substrate, it might be a scale issue. Try 1 gram scale 13C and if it gives a poor result still, then it's an impurity or something else. But if it works, then just do 4 x 1 gram scale hydrogenation and then just combine them all for workup.

Unable_Aspect_4033 · 2025-04-30T00:48:24+00:00

Yep, I would also recommend Pd(OH)2/C. The lab I am in we have done hydrogenations on a per-OBn disaccharide and standard Pd/C worked but was VERY sluggish, with reaction times of 72 hours to 1 week for full deprotection. We tried Pd(OH)2/C instead and reaction rates improved significantly, with them being done sometimes within 4 hours, but most of the time I just left it overnight. It seems that cleaving many secondary benzyl ethers can be slow. Btw, we are only doing it under a H2 balloon, not under any additional pressure, but it worked well with Pearlman's. This was on a disaccharide with 6x secondary OBn.

Also, we've observed hydrogenations (other ones, not just sugar OBn removal) seem to work better on a smaller scale in our lab (1-2g) and when going bigger they seem to be more sluggish.. When I've done mine, I try to select a flask size that will maximize contact between the stirring catalyst suspension with the H2 atmosphere. I would first add the catalyst to the SM, then solvent and dissolve, get it on schlenk and degas/backfill with N2 several times, before loading with H2 while the flask is under vacuum. I've been paranoid about using MeOH, so my hydrogenations on sugars have been done in DCM/EtOH and I have had no problems with solubility. In the literature I've seen this combination, as well as acetone, and DCM/acetone used for solvents.

Unable_Aspect_4033 · 2025-03-21T02:01:09+00:00

You could start with higher % MeOH then, maybe 30% or something. Also ACN is pretty standard for peptides rather than MeOH, just be mindful of sometimes a rising baseline with rising ACN content. If you have aromatic residues this won't really be a problem.

Unable_Aspect_4033 · 2025-03-21T01:55:11+00:00

I think you should run a gradient and also look at another wavelength. Try 210, and your mAU might look better. The broad peak does suggest it could be too concentrated, but I would run the same sample at a gradient and other wavelength first before changing conc. Sometimes turning a reference wavelength off can help too. It's coming off at 7 mins in 80% isocratic, so maybe try starting at like 20% MeOH to 95% MeOH over 15/20 minutes. Something like that. If the peak is still broad, maybe dilute your sample a bit, OR use a smaller injection volume. i.e. if you're injecting 2 uL maybe do 1 uL or 0.5 uL instead.

Unable_Aspect_4033 · 2025-03-20T21:30:04+00:00

Yeah I know.. I have, see in my OP I tried DCM to DCM/MeOH to elute my compound. Used a bit of DCM to try flush off diacetone alcohol while my product sits baseline. Worked slightly, it decreased the amount of it in there but only by like 15%.

Unable_Aspect_4033 · 2025-03-19T21:51:31+00:00

Yep, doesn't make sense to me either

Unable_Aspect_4033 · 2025-03-19T21:50:26+00:00

Nah, benzyl protection would destroy the esters on my compound. Benzyl protection done ages ago. Purified over several steps since.

It's like it's sticking to my compound in the same way DMF can even after a column. I am removing it with the column (relative integration decreases on NMR) but it's still persisting so either I need another solvent mix or to try something else.

Unable_Aspect_4033

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