Chemical Forums
Chemistry Forums for Students => Organic Chemistry Forum => Organic Chemistry Forum for Graduate Students and Professionals => Topic started by: kriggy on October 27, 2021, 08:51:09 AM
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Hey guys,
its somehow related to my previous question but its probably better to start a new topic.
Anyway, Im strugling to remove my protecting group: phenylfluorenyl. I can get the protected intermediate without much trouble but the removal is pain. Using TFA with various scavengers (EtSiH, TIPS, MeOH...) I clearly see the starting material gone (by TLC) but afterwards, I dont see the mass of the product in my LCMS analyssis or only traces to be more specific and the spectrum further shows various unknown impurities. Furthermore, only very weak oftentimes none ninhydrin reaction is observed with TLC analysis.
I tried to deprotect the PhSe intermediate hoping the double bond is the cause of the problems with similar results observing some kind of PhSe transfer to get double selenylated product. Hydrogenation of the PhSe at 2.5 atm did not show any traces of reaction.
Furthermore, I recently started the same sequence with trityl protected aminoacid (trt is more labile to acid) but during first step (KHMDS enolate then PhSe quench) im observing poor reactivity and cleavage of the trityl group. Anyone observed this happening? I did quench the reaction with acetic acid but I dont think trt is that labile.
Im glad for any tips you can give me. Esp. some tricks for LCMS detection of aminoacids are wellcome (i detect by SIR but from that its hard to extimate the amounf of the aminoacid in my sample
thank you
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Is it necessary to use TFA? The trityl goes with 80%HOAc, maybe you can try that, maybe some heat? Maybe scavenger is bad in this case, I would try without.
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Pf is about 6000x times more stable so I think TFA is needed. I might try HCl in ether and maybe the hydrochloride crashes out. I think I did try without the scavenger and the results are fairly the same. I get rid of the Pf group (TLC) but then im unable to isolate the product.
I think I might be actually getting the pyroglutamic derivative or some kind of michael addition.
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I work a good deal with unsaturated amino acids, but I doubt that I can be of much help in this instance. KMnO4 dipping has worked well for visualizing our TLC plates. The amino group should not be performing a Michael addition, unless one is under basic conditions; therefore, I am puzzled. Regarding MS, if I could spare a small portion of a product with a free amino group, I would at least consider using heptafluorobutyric anhydride to make a derivative.
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OK, maybe you can run a deprotection in a NMR-tube to see what goes wrong, if the double-bond is affected this should be easy to spot.
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Protect the seleno compound from oxygen. It is stable to acid. Seleno compounds can oxidize during column chromatography... Why not just deprotect then do the elimination step???
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The amino group can be oxidized if its unprotected I think, I guess you use hydrogen peroxide to get the selen oxide before elimination? Also the unsaturated product is a Michael addition substrate so a free amino group can add to the double bond. Its also a di ester so it can ringclose or polymerize via aminolysis.
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The amino group can be oxidized if its unprotected I think, I guess you use hydrogen peroxide to get the selen oxide before elimination? Also the unsaturated product is a Michael addition substrate so a free amino group can add to the double bond. Its also a di ester so it can ringclose or polymerize via aminolysis.
Yes MCPBA may be the wrong reagent. I would try a chlorine source under aqueous and buffered conditions.
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Protect the seleno compound from oxygen. It is stable to acid. Seleno compounds can oxidize during column chromatography... Why not just deprotect then do the elimination step???
Im able to purify on column without any issues - except the starting material spot is very cloes to the product as much as they are almost coeluting but its doable.
The amino group can be oxidized if its unprotected I think, I guess you use hydrogen peroxide to get the selen oxide before elimination? Also the unsaturated product is a Michael addition substrate so a free amino group can add to the double bond. Its also a di ester so it can ringclose or polymerize via aminolysis.
Yes MCPBA may be the wrong reagent. I would try a chlorine source under aqueous and buffered conditions.
The original procedure for the elimination on this cpd used mcpba, I find NaIO4 bit better and easier to do (just mix and stir) but the deprotection is then not described because they do ester reduction, dihydroxylation and deprotection later.
Protect the seleno compound from oxygen. It is stable to acid. Seleno compounds can oxidize during column chromatography... Why not just deprotect then do the elimination step???
I did attempt that as well (one experiment) and while I observed the product in LCMS only traces were there and mostly it was some uknown which was assigned from m/z to compound having 2 SePh groups so some kind of rearrangement or transfer probably happened.
I work a good deal with unsaturated amino acids, but I doubt that I can be of much help in this instance. KMnO4 dipping has worked well for visualizing our TLC plates. The amino group should not be performing a Michael addition, unless one is under basic conditions; therefore, I am puzzled. Regarding MS, if I could spare a small portion of a product with a free amino group, I would at least consider using heptafluorobutyric anhydride to make a derivative.
Thank you. Im used to using CAM which I thought it should work but maybe KMnO4 is better in this case.
OK, maybe you can run a deprotection in a NMR-tube to see what goes wrong, if the double-bond is affected this should be easy to spot.
That is actually great idea. Dont know why I did not think of that already.
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The seleno compound will also stain with KMnO4, it will oxidize.
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I repeat also, this compound with the amino group as free base is not stable.
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I repeat also, this compound with the amino group as free base is not stable.
Why do you think so? Im fine with it being whatever kind of salt that allows subsequent amide bond formation or even deprotecting it in-situ before the coupling
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I like to think of the proton as a poor man's protecting group.
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The amino group can be oxidized if its unprotected I think, I guess you use hydrogen peroxide to get the selen oxide before elimination? Also the unsaturated product is a Michael addition substrate so a free amino group can add to the double bond. Its also a di ester so it can ringclose or polymerize via aminolysis.
As I wrote here, the free base is problematic but it depends on the timescale, it could be stable for short time.
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So I did run the NMR experiment and after addition of TFA the double bond signal shifted towards lower ppm. Im not sure if its because of the N protonation or because of the deprotection is happening but after about hour or so, the double bond signals dissapeared completely. I dont have much idea whats going on but seems I need to do the deprotection quick.
Im running the same sequence with trityl and for some reason, the Phenylselenyl derivative is decomposing to something which is neither alkene nor selenoxide and for some reason, it does not get oxidized using same condition as the Pf derivative..
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We deprotect our amino acids using TFA, and we purify them over Dowex-50 using water first, then HCl. We have not seen loss of the double bond.
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I guess you see coupling pattern on the double bond, if this is intact you just have to run the deprotection for shorter time as you suggest. Whats happening with the trityl is hard to say but it seems like a dead end.
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We deprotect our amino acids using TFA, and we purify them over Dowex-50 using water first, then HCl. We have not seen loss of the double bond.
Hm I dont think we have dowex but im going to look around if I can get some. Also, how do you know how much water / HCl to use? I suppose you cant realy TLC it to see how it moves on dowex column?
I guess you see coupling pattern on the double bond, if this is intact you just have to run the deprotection for shorter time as you suggest. Whats happening with the trityl is hard to say but it seems like a dead end.
I guess. Im worried that the double bond moved around but thats not something I can figure out now without having the purified amino acid.
Thanks for help guys
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TFA will not make a double bond move, if there is a double bond I think its OK.
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For Dowex-50 we use step gradients, such as water then 4 M HCl. We make large pools, comprising several column volumes. We often load the amino acid in 0.5 column volumes of water, then use 1.5 column volumes of water as a wash. Then we use 2-3 column volumes of HCl. If an amino acid elutes slowly (and when there is a hydrophobic side-chain, it has in our hands), then the product still make show up in more than one pool. We take several pools, rotovap/vacuum, and check masses.
Dowex-50 is an old-school technique for purifying amino acids. Often people used water followed by 2 M ammonia , and I have used this many times myself in the past. Although I have seen people back the ammonia down to 0.5 M once or twice (presumably for sensitive side chains), I decided to avoid any possibility of its attacking the amino acid, and that is why I chose to elute with HCl. 4 M HCl is probably overkill, but people have used it before, so I just decided to try it. We only occasionally use TLC on amino acids, but Stahl's venerable TLC book gives five common solvents. 70:30 ethanol/water is one of them. There are both acidic and basic conditions.
We occasionally use Dowex-1 in the acetate form, but it does not always work in our hands for reasons that are not known to me. The idea is for the carboxylate group of the amino acid to displace an acetate ion on the resin. Acetate is less strongly bound to the resin than chloride; the conversion of the resin from the chloride form to the acetate form is lengthy but not difficult. If you need any references or protocols, just let me know.
This brings me to a question. Someone once asked me if I knew of any papers on the purification of amino acids by ion-exchange. I said that I did not, but that I could almost write one. Since that time my students and I worked out one additional method that is applicable when the side chain carries a charge. Would any journal or serial publication (by which I mean something like Methods in Enzymology) be interested in such a work?
EDT
Just FYI, there are some reviews (I have not read them in a while) of unsaturated amino acids that deals a little with isotope labeling:
Kaiser J et al., Org. Biomol. Chem. 2005 3:3435-3467. (a review which covers 2000-2005)
DOI: 10.1039/b001538p (a review which covers 1990-2000 IIRC)
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You could use some MeOH in the eluent to push out the product from the resin if you have lipophilic groups in the product. 4M sounds really strong, I would try 0,5M for sensitive compounds. But if it works good, there is no reason to change anything offcourse.
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@Babcock_Hall:
Thank you very much for your reply and the references, it was very helpful.
I have few question towards the work with dowex. At first, I need to wash with HCl to make sure its in the H+ cycle, then with water untill the elution is neutral. Then add my aminoacid in water (free base not salt) and elute with water and then with HCl. ?
If my reaction mixture contains various salts (NaIO4) I suppose they are converted into HIO4 and still elute while the Na+ is retained on the ion exchange?
Thank you.
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When converting from a more tightly bound ion to a less tightly bound ion, one has to use a greater volume of the new ion. "As a general rule, use 1 bed volume of 1 M counterion solution for each unit difference in relative selectivity. For example, converting AG 50W-X8 resin from the K+ form (relative selectivity 2.5) to the H+ form (relative selectivity 1.0) would require 2-3 bed volumes of 1 M HCl. The conversion is complete when all the K+ ions are displaced by the H+ ions." This information is useful when one reuses Dowex-50 that is in the ammonium form (see below).
Regarding the rinse after the conversion (for example) from H+ to Na+, BioRad recommends 4 bed volumes of water and checking the pH. The protons on Dowex-50 should protonate your amino acid. The sodium ions will also stick. Therefore when you calculate how much resin you need, you will need to sum the moles of amino acid and the moles of NaIO4. I typically use 20-fold capacity of resin, relative to this sum, but a lower ratio may also work. You can elute with acid or with ammonia. In the presence of a sensitive group, I have seen a few authors elute with a lower concentration of ammonia than is typical (0.5 M vs. 2.0 M), but if HCl is compatible with your functional group, then it should work fine. We have used HCl in the presence of either an aldehyde or a Michael acceptor.
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http://www.jbc.org/content/213/1/39.citation
Black and Wright, J. Biological Chemistry (1955) 213:39-50 converted allylglycine into aspartate beta-semialdehyde and then purified over Dowex, using 4 M HCl. There are probably other examples of acid elution in the literature, but my paper copies of articles on this subject are in arrears (I would start by looking up papers by Stein and Moore, who did a great deal of analytics biochemistry on amino acids and proteins many years ago). I think that 4 M is overkill and 1 M would probably work fine, but we used 4 M twice, both in the presence of a Michael acceptor. The alternative to HCl elution is elution with ammonia, which deprotonates the bound amino acid.
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Yes, if it works its fine. I guess you have to rotavape a lot of acid, othervise it does nor matter.