Org Prep Daily

September 27, 2006

TLC Staining solutions

Filed under: procedures — milkshake @ 2:28 am

General Staining:

Cerium-ammonium-molybdate, CAM

40g of ammonium pentamolybdate + 1.6g of cerium(IV) sulfate + 800mL of diluted sulfuric acid (1:9, with water, v/v). On heating, blue-black spots on light background. Slowly fades over several days. Quite universal, often very sensitive. Some amines, amides and oxidation-resistant aromatics do not detect well.

Basic KMnO4

40g of K2CO3 + 6g of KMnO4 in 600mL of water, then 5mL of 10% NaOH added. (KMnO4 takes some time to dissolve completely. Lazy people like me substitute it with NaMnO4 concentrated aq. solution from Aldrich). No heating. Brown spots on pink background. Often very sensitive but staining very disproportionate to quantity, depending on the compound. Fades within hours. Oxidizes anything with diol, C=C, reactive methylene, phenol, thiol, phosphine etc. Particularly useful for detection of tertiary amines.

Phosphomolybdic acid

30-40g of phosphomolybdic acid in 100mL of ethanol (preferably non-denaturated). Good grade of phosphomolybdic acid should provide clear, bright yellow solution. (If there is cloudiness, let it settle and decant.) Light sensitive. On heating, blue-black spots on yellow-green. Good for lipids. Do not overheat or the background goes dark. Usefull for spraying but expensive as dipping-jar solution. (I stopped using PMA, in favor of CAM. One advantage of PMA is that it is compatible with aluminium-backed TLC places whereas most other universal stains containing diluted sulfuric acid, like CAM or anisaldehyde, are not and must be used with glass TLC plates.


40mL of conc. H2SO4 is added (slowly!) into ethanol 800mL, followed by acetic acid 12mL and anisaldehyde 16mL. Light and oxidation sensitive. On heating, colorfull spots on pink background. Color varies on the compound. Good for all things with active methylene, and for distinguishing closely-spaced spots on TLC by their color difference.


Iodine vapor chamber is made from a TLC jar with a good-sealing lid, by adding a dry mix of iodine crystals  (a small spoon) covered with silicagel for chromatography, about a half-inch layer. Put dry TLC in the chamber onto the silica layer face down and watch the brown spots developing. Works on variety compounds but often it is only moderately sensitive. Iodine stained TLC can be developed subsequently with other stains.

Functional group-selective stains


20g of ninhydrin in 600mL of ethanol. (Don’t spill ninhydrin onto your fingers – they would go blue.) Primary amines produce blue spots at R.T., very sensitive detection. Boc-protected primary amines produce spots on heating (as the Boc falls off). Secondary amines sometimes detect but the stain is weak.


3g of dinitrophenylhydrazine in 750mL of 2M HCl. (If htere is some insoluble portion, decant it off.) Aldehydes and ketones produce yellow-orange spots at R.T., quite selective.

Stain dip station: 4 or 6 wide-mouth jars covered with aluminum foil (secured by tape) to protect from light, placed within a tray (to guard against spill).

Spray station: Hand-operated rubber-baloon/glass top sprayers are wonderful but antiquity now – they are hard to get by nowadays. Compressed gas sprayers are much harder to control. Spray advantage: economical on stain solution, the TLC spots do not move as with dipping. Disadvantage: big mess in the hood, a receiver for spraying has to be built (i.e. from a cardboard box). I used to be very partial to spray-staining but was won over by the dip jar convenience.

Update: A procedure for improved Dragendorff stain that keeps well in refrigerator (+4C). This stain is particularly useful for detection of lipophilic amines, basic heterocycles like pyridines but also aryl phosphines, crown ethers and polyethylene glycol polymers. Brown spots on yellow background develop almost instantly at R.T.

A solution of L(+)-tartaric acid 20g in D.I. water 80 mL was added to BiO(NO3) 1.7g and the mixture was sonicated on ultrasonic bath for 15 minutes. A solution of KI 32g in D.I. water 80 mL was added into the mix. Finally, a solution of tartaric acid 175g in D.I. water 950mL was added. The resulting bright orange mixture was stirred for 15 minutes and then placed into a fridge overnight. The solution was decanted off from precipitated crystalline solids (probably K-tartarate), transferred into a wide-mouth dip jar and kept in fridge when not in use. (This Dragendorff reagent gradually darkens over time but the aged reagent still performs quite well even after several months in the fridge.)


  1. Hey, just wanted to let you know that I think this blog is great. I love seeing old-school chemistry, plus the Mosher acid substitute and the azide source look like they could be useful some day.

    I’m a big fan of (fresh) PMA; screw the expense. Black spots on yellow background scans well for those of us too lazy to draw in our TLCs.

    Comment by Peter — September 27, 2006 @ 2:58 am

  2. Thank you, Peter, for the kind words. Btw – I did most of my thesis work on the PMA-stained TLCs. (In Prague, NMR and GC/MS access was a favor one had to beg for in those times, my phospholipids would not crystallize so I had to column pretty much everything.)
    With the work I do now – in medchem, the simplier the better – we don’t do much asym. synthesis if we can help it. We BUY chiral blocks. Most of the times, one ends up making lots of simple pieces, for some subproject that gets finished or killed within few months. Medicinal chemists patent a lot but don’t publish as much, for obvious reasons. Most of the intermediates are no longer usefull for the company after some time. There is no danger in showing these procedures – but they are unpublishable. So I decided to put up scraps of my current and past work – if I though I had a decent procedure and the stuff would be at least mildly interesting.

    Comment by milkshake — September 27, 2006 @ 3:25 am

  3. Check out this pretty comprehensive list of tlc stains, a lot of which are chemoselective.
    TLC Visualization Reagents
    I know i love trying to predict how my compound will behave on silica, and using chem tests to help elucidate results of tlcs. Please feel free to write in reviewing your experiences with some of these.
    I regularly use the ninhydrin dip, for amines and sometimes it also works for amides carbamates although, its less selective for these.
    I have been a big fan of various tests for phenols although would love to see how the chloranil and other tests fair. The hydroxylamine/FeCl3 solution for detection of amides, lactones, carboxylic acid esters and anhydrides is kinda cool. These tests rock when your just doing simplish protection/deprotection steps and just kinda want to confirm it is what it is before you column/nmr/ms etc

    Comment by Q — September 27, 2006 @ 9:19 am

    • yeh for amines we are use ninhydrine,it may be ok.,
      but for debenzylation prouct havimg corboxylli acid in valine ring what we do.
      i tried kmno4 but not getting clearly..?

      Comment by vasanth — June 6, 2011 @ 10:42 am

      • Amine detection with KMnO4: Tertiary amines are oxidized much faster so they will appear right away, whereas with secondary and primary amines the staining takes few minutes at room temperature to develop.

        You can use a Dragendorf reagent stain to reveal all basic compounds (and crown ethers) as orange-brown spots on yellow background

        There is a stain system that is very sensitive for primary, secondary amines and for amides, sulfonamides with free NH:

        First you spray the TLC with a diluted solution of tert-butyl hypochlorite tBuOCl in dichloroethane, then you dry the TLC with a stream of ambient air until the chlorine-like sharp smell of hypochlorite is no longer perceptible by sniffing, then you reveal the presence of N-chloro compounds by a second spray with a diluted solution of tolidine (4,4′-diamino-3,3’dimethylbiphenyl). It shows as black spots on white background.

        Comment by milkshake — June 6, 2011 @ 9:09 pm

  4. Another great stain is Seebach’s (aka “Magic”).

    2.5 g phosphomolybdic acid
    1 g Ce(SO4)2
    6 mL conc H2SO4
    94 ml H2O

    The addition of Ce salts to standard PMS gives a really nice royal blue color, and it is “magic” because nearly everything stains.

    Comment by Jose — September 27, 2006 @ 11:03 am

    • What is the shelf-life of Seebach’s stain?

      Comment by Megan — August 22, 2017 @ 6:41 pm

      • It is very good, year or longer at room temperature if you take care to protect it from light (I would wrap Al foil around the jar) and if you avoid dipping TLCs wet with organic solvents. The stain goes bad by reduction of Ce(IV) to Ce(III) so when the stain is no longer yellow it is time to replace it.

        Comment by milkshake — August 22, 2017 @ 6:44 pm

  5. yes, that should be “PMA,” and not “PMS.”

    Comment by Jose — September 27, 2006 @ 11:05 am

  6. Having used Dragendorff stain dip extensively, I can tell you that the tartaric acid recipe is much more effect. Scott Rychnovsky has a good recipe on his website. Once made, it does tend to discolor over several months (getting darker). However, when stored on the benchtop, it continues to be effective for 2-3 months. The main change over time is that the background stains darker as the dip gets older and the contrast is not as great.

    Comment by Chemist of Sorts — September 27, 2006 @ 11:29 am

  7. During the early days of my research in india, the TLC book by Egon Stahl and the perkin elmer IR machine were the bread and butter of my everyday chemistry life. Paying for NMR was too dear for my then PI, therefore, we almost always tried to confirm the structure based on TLC and IR, followed by UV and in-house massspec and only then did we send it out for NMR. Fact that we were working with coumarins and terpenoids helped.
    Though i like the smell of TLC dipped in anisaldehyde stain (am i a freak?), PMA is what i use most. Thanks for the recipes and i haven’t seen a sprayer in years. Great blog, hope to see a lot of stuff in here.
    Having shifted to a med chem lab recently, i wonder if some deadlines in these labs are just an excuse to do some not-good chemistry.

    Comment by sks — September 27, 2006 @ 1:52 pm

  8. you’ll get far better sensitivity for secondary amines with this ninhydrin recipie:

    -0.75g ninhydrin
    -250 mL 95% EtOH
    -2.5 mL glacial AcOH

    Mix it up, it stores well on the bench sealed. dip your plate in, gently heat. voila! secondary amines appear as orange spots. i used this routinely to analyse reductive amination reactions, and it worked for me.

    Comment by kiwi — September 27, 2006 @ 6:37 pm

  9. Thank you for the Dragendorff, ninhydrin and PMA/Ce(IV) tips, this is most helpfull.
    Also, I remember using extremely unhealthy but powerfull detection system 2 decades ago: tert-Bu-hypochlorite in CCl4 first spray, dry with cold hair-dryer until no more stink of hypochlorite is palpable by sniffing the TLC, then spray with a solution of m,m’-dimethoxybenzidine (anisidine??). Amides and amines which have a free NH give intense black spots on white background (or black background if one forgets to dry the hypochlorite first spray). Very general.

    Comment by milkshake — September 28, 2006 @ 3:23 am

  10. Milkshake: this is an awesome post–printed and saved as a PDF for future reference. A French postdoc in our lab used the Dragendorff stain and I never asked him how to make it before he left. I tried looking for a procedure, but I never found one (probably because I was spelling it wrong). Good stuff.

    Comment by Paul — September 28, 2006 @ 5:51 am

  11. I’m curious why everyone does their KMnO4 stains in water. I’ve always used acetone. Drys much faster and works very effectively.

    Comment by Javaslinger — March 13, 2007 @ 10:26 am

  12. I was doing KMnO4 stain spray in acetone in Prague and it worked – the only problem was that I had to spray (dip would smear the spots), the KMnO4 solution in acetone had to be made fresh and the KMnO4-stained plates did not last nice too long – the background was turning brown quickly.

    Comment by milkshake — March 13, 2007 @ 2:24 pm

  13. I am not having much success with your KMnO4 stain recipe… I get a very smeared and splotchy TLC… My post doc has us mix KMnO4 with Acetone. It seems to work much better but turns to a nasty brown sludge pretty rapidly. Where might I be going wrong with your KMnO4 recipe? I’ve looked around the net and they all seem basically the same, so I’m sure the recipe is fine….

    Also, why don’t people use Ethanol instead of water?



    Comment by Javaslinger — May 7, 2007 @ 2:36 pm

  14. ethanol does not mix well with permanganate as you should know. Even acetone solution of permanganate does not keep well and has to be made fresh every time.

    Permanganate diluted solution fall apart over time, the concentration decreases and MnOx crashes out. What I found helpful was to 1. Use good reagents (organic impurities will make the solution go bad faster). 2. Make a big jar of the reagent dip solution and don’t shake it, let the MnOx muck that slowly forms over time to settle. 3. Wrap the jar in Al foil to keep it from light. 4. Re-make the new KMnO4 stain every two months or so.

    Comment by milkshake — May 7, 2007 @ 7:47 pm

  15. Hi
    could anyone tell me how can i stain specifically for furanocoumarins or if tht is being too specific for coumarins. Im a botany student and dont know much about all this stuff.

    Comment by renuka — June 28, 2007 @ 9:37 am

  16. I would use CAM – but I think permanganate or anisaldehyde should work as well

    Comment by milkshake — June 28, 2007 @ 3:12 pm

  17. Milkshake: the anisaldehyde, should I use only para? As for the “Magic” (#4) solution, how long should I keep it on the bench, before making another fresh batch? ;>

    Comment by taiatuwai — July 11, 2007 @ 1:35 am

  18. The aglycon of cumarins are detectable by UV (Dafeentin is yellow but others are are blue under UV ).
    Furanocoumarins have various colours under UV but their colour is increased by KOH or SbCl3 in CHCl3.

    Comment by Ghannadian — August 28, 2007 @ 5:38 am

  19. why is it that when staining with an iodine bath and using a sulphuric acid spray the results of triolein never show up in thin laer chromatography? i have tried the iodine first and the spray first but they did not manage to show up anything

    Comment by Hazel — November 12, 2007 @ 1:35 pm

  20. maybe you should try CAM or phosphomolybdic acid instead, they work fine for the lipid detection

    Comment by milkshake — November 12, 2007 @ 4:40 pm

  21. Hey
    can anyone tell me wat i can use to stain polyketides(namely pederin.Thanks

    Comment by Asrar — January 3, 2008 @ 6:10 am

  22. CAM should be best for your purpose. Anisaldehyde, phosphomolybdic acid and KMnO4 as second, third and fourth alternative

    Comment by milkshake — January 3, 2008 @ 7:27 am

    • How do you interpret the colours formed on the anisaldehyde and PMA stained plates?

      Comment by CLEMENT — June 7, 2012 @ 6:54 am

      • you dont. PMA stains everything pretty much bluish-black. Anisaldehyde and vaniline stains do produce various colors with different compounds but it is hard to tell without a prior knowledge what to expect

        Comment by milkshake — June 7, 2012 @ 8:46 am

        • do you know what the stains colours with vanillin means???

          Comment by ourvarshee — November 13, 2015 @ 1:45 pm

          • it is hard to tell – some kind of messy condensation reactions

            Comment by milkshake — November 13, 2015 @ 2:53 pm

  23. hi, anybody know the staining reagents for lipids and hydrocarbons detection?

    Comment by Kent — February 3, 2008 @ 3:24 am

  24. Lipids and hydrocarbons are easy to detect: I would recommend CAM or phosphomolybdic acid. But your hydrocarbons can’t be volatile because the detection is done by heating the TLC.

    Comment by milkshake — February 3, 2008 @ 1:22 pm

  25. CAM is the stain which gives me perfect information regarding arganic compounds
    especially about aliphatic compounds with lesser active functional groups such as ethers,
    alcohols etc..

    Comment by Naresh reddy — February 22, 2008 @ 8:12 am

  26. Can anyone tell me what stain to use to visualize sulfur containing lipids (i.e sulfolipids) on a TLC plate.

    Comment by Olivera — February 22, 2008 @ 4:12 pm

  27. I would use CAM or phosphomolybdic acid

    Comment by milkshake — February 22, 2008 @ 4:37 pm

  28. Hi, anyone know why sulfuric acid is used when preparing staining solution? Why sometimes heating is required? What is the role of acetic acid in TLC mobile phase?

    Comment by Kent — February 28, 2008 @ 11:53 am

  29. Do you know if wax esters are stained by FeCl3 (acid solution)?

    Which kind of lipids are stained by anisaldehyde? Which colour would you expect to obtain staining wax esters and sterol esters with anisaldehyde?

    Thanks a lot for your help!

    Comment by Apolo — March 27, 2008 @ 10:06 am

  30. Wax/sterol esters will not stain with FeCl3. They might stain only very poorly with anisaldehyde or not stain at all (especially if they are saturated) because anisaldehyde staining is dependent on acid-catalysed condensation+dehydration reactions, to make conjugated colorful products.

    Use CAM or phosphomolybdic acid. If you ask me I prefer CAM for the purpose (there is discussion of the two at Curly Arrow, see the blogroll)

    Either way, it is pretty easy for you to try and convince yourself. I suppose you have standards of the stuff you want to detect, so make a TLC and see if you can detect the spots/

    Comment by milkshake — March 30, 2008 @ 6:50 am

  31. Just a couple of notes. out of my own experience

    For Phosphatidyl type lipids : H2SO4 or PMA are winners
    For PEG compounds use I2
    For various amino acids and pyrazines/pyrazinones: Use CAM …#

    Hope this of use

    Comment by Jeroen — May 20, 2008 @ 7:19 am

  32. I’m currently dealing with lipid compounds from actinomycetes for antibacterial activity. Just curious whether CAM and PMA have the same colour for lipid compounds (dark green i suppose), and which one is a better one for the detection of lipids. Thanks.

    Comment by Lexxie — October 30, 2008 @ 1:26 am

  33. The spots are usualy bluish-black in both cases (“Mo blue”) but CAM has a weaker background and seems more sensitive. I suggest that you try both and find what works best for you.

    Comment by milkshake — October 30, 2008 @ 2:23 pm

  34. Thanks alot. Milkshake. However, i need to determine which of the two stains give green colour for lipids (from Handbood of Thin layer chromatography, Joseph Sherma). I tried another version of molybdate staining solution by using disodium molybdate (VI) dihydrate, water, H3PO4 and H2SO4, in which im not sure whether it will give the same colouration as CAM and PMA. On the otherhand, the compounds that im dealing with currently are not UV active. Thus, i cannot make use of UV light and HPLC to determine the UV patterns of that unknown compound (coz my HPLC has only deuterium lamp), any suggestions or alternatives on this matter, of how im going to purify my compounds. Thanks alot.

    Comment by Lexxie — October 30, 2008 @ 8:17 pm

  35. I did my thesis work on phospholipids and it was all silica column and TLC. There are evaporative/spray HPLC detectors that are applicable on lipids but they are quite expensive.

    Comment by milkshake — October 31, 2008 @ 4:14 pm

  36. Alrite, thanks alot milkshake. Have a nice day 🙂

    Comment by Lexxie — November 5, 2008 @ 8:24 pm

  37. Hi, I am looking for a good reagent(s) to use in detecting sesquiterpene lactones (STLs), especially the alpha,beta-exocyclic methylene-containing ones. Specifically, the guaianolides, pseudoguaianolides, germacranolides, etc., are of particular interest. Mostly I see references for para-anisaldehyde/H2SO4 but I don’t think this is very specific to these compounds. I have also found studies using phosphomolybdic acid, iodine, Zimmerman and Godin reagents. I am not familiar with the latter two reagents. Any advice for STLs? Thanks if you can help.

    Comment by alchemist9331 — April 22, 2009 @ 1:02 pm

  38. Your best bet would be cerium ammonium molybdate (CAM), on glass-backed TLC plates, but phosphomolybdic acid spray should work well also. In both cases the TLC has to be heated after drying to develop the stain, and the stain reagents are best stored in dark or in aluminum-foil-wrapped jar.

    Basic permanganate should also do excellent job with C=C compounds but CAM stain is more proportional to the quantity of the material and the staining is stable for many days afterwards whereas spots detected by permanganate fade rather quickly

    Comment by milkshake — April 22, 2009 @ 2:05 pm

  39. Thanks for the quick reply!

    Comment by alchemist9331 — April 22, 2009 @ 2:54 pm

  40. Why is it that acetic acid is used in reducing smearing in TLC preparation

    Comment by sam — May 13, 2009 @ 9:35 am

  41. Acetic acid binds strongly on silica, if added to some TLC development systems it forms a gradient on silica as the TLC develops. That can supress tailing of acidic compounds in the separated mix.

    Comment by milkshake — May 13, 2009 @ 7:38 pm

  42. What would be the best stains to use to distinguish between a carboxylic acid and an ester on a prostaglandin intermediate?

    Comment by lunarhog — May 20, 2009 @ 11:39 am

  43. there is this spray with diluted hydroxylamine (free base) solution in ethanol, followed by drying with a heat gun, and then developing with diluted FeCl3.hydrate solution. Non-hindered simple esters get transformed into hydroxamic acids and these provide bright-red complex with Fe(3+).
    Personally I have no experience with this system, and I suppose one needs a real hand-spray to make it work (dip jar will probably smear the spots).

    But during a phospholipid-related project I noticed that simple fatty acids were very easy to separate from esters – when the TLC was first developed in hexane and then in a mixture chloroform-methanol-aqueous conc. ammonia 100:10:1 (volume ratios). Ammonia in the mix retards the free carboxylic acids considerably so they will make a nice sharp spot with low Rf at the bottom whereas the esters will go way up.

    Comment by milkshake — May 20, 2009 @ 3:27 pm

  44. COuld you help me the best staining system for carbohydrate moities.

    Comment by Bala — June 11, 2009 @ 3:15 am

  45. CAM and anisaldehyde

    Comment by milkshake — June 11, 2009 @ 3:28 am

  46. I came back here to look at different stains for my somewhat exotic compounds which CAM stains only faintly.
    Although you don’t list it, I tried vanillin – no good for my stuff. Very good for irritating my nose and my colleagues.

    I ended up settling with simple permanganate. A little trick that we used in our old lab — wash off excess pink with cold water (at least a minute at the bottom of the sink) – this makes the contrast much better. Only works for aluminum plates, because silica is washed off easily from the glass ones.

    Comment by LiqC — July 28, 2009 @ 3:00 am

  47. Has anyone any idea which TLC reagents would be the most sensitive for PEG200 and PEG8000 and what are the lowest amounts detectable of these compounds in TLC, are we talking about 1 ug, 10 ug or 100 ug levels? Thanks!

    Comment by Sami — August 24, 2009 @ 1:49 pm

  48. I don’t know about the quantitation limits – a hard thing to guess – I suppose its easy enough to determine empirically, by doing a dilution series on TLC with a calibrated-volume capillary as a spotter.

    I would try detection with CAM and Dragendroff. (Dragendorff sounds weird but people have been using it for detecting crown ethers so I guess PEG is close enough)

    Comment by milkshake — August 25, 2009 @ 11:53 am

  49. Hi,

    I am a synthetic chemist and looking for good stain which can detect N and S atoms present in the ring. The ring is four membered thiazetidine ring having tertiary N.

    Comment by Ankita — November 11, 2009 @ 11:56 pm

    • I would try the basic permanganate – it is great for tertiary amines and thioethers.

      Comment by milkshake — November 12, 2009 @ 7:30 am

  50. Hi,

    i am working with peptoids which terminates with a secondary amine. Can u please suggest me a suitable staining agent.i tried Kmno4, ninhydrin, iodine nothing worked. thanks

    Comment by nilaya — December 14, 2009 @ 2:01 am

    • On solid phase, for detection of your material attached to the resin, I would use the modern version of chloranil test, 1 drop of 2% CH3CHO in DMF (v/v) followed by one drop of 2% chloranil in DMF. One needs to use good-grade DMF that does not smell like fish (= it is perfectly free of dimethylamine) and store these two solutions separately in freezer because they age at room temperature fast – but they keep good for about a month when refrigerated. Its from a journal called Peptide Reseach, the paper is from mid-1990s and the author name is T. Vojkovsky. Its a modification of a method from an old paper ( I think from 1960s)

      In solution on TLC I would perhaps use spray with diluted tBuOCl in carbon tetrachloride (tert-butyl hypochlorite, it is commercial and there is also easy way to make it from chlorine gas, bleach and tBuOH), the TLC is dried with warm heat-gun (but not too hot) to remove the excess of hypochlorite – until the chlorine smell of hypochlorite is no longer perceptible by sniffing at the TLC. The presence of N-chlorinated products is then visualised by another spraying the TLC, with a diluted solution of o-tolidine (2,2′-dimethyl-4,4’-benzidine). Black spots develop immediately without heating.
      This detects all compounds with free NH and the staining is very sensitive on glass-backed plates (black spots on white-bluish background) but the chronic toxicity of o-tolidine and carbon tetrachloride can be a drawback on repeated use, so the spraying needs to be done in some dedicated “poisoned” fume hood and a good hygiene needs to be observed. Oh, and the solutions have to be stored in fridge because they also age at room temp. Sorry I don’t remember the exact composition, it has been about 15 years since I used it last time, it could have been something like 5% tBuOCl and 2% o-tolidine but you need to look it up.

      Comment by milkshake — December 14, 2009 @ 2:28 pm

  51. thankyoy sir 4 your immediate reply

    Comment by nilaya — December 14, 2009 @ 2:34 pm

  52. Hello there,
    I hope you can help,
    I need to detect amines and amides. I have lauroimidazoline from reacting lauric acid and AEEA. However I need to determne if there are side reactions or residual AEEA present. I was wondering if anyone has any suggestions including TLC or HPLC.
    Thank you in advance.

    Comment by LULU — December 29, 2009 @ 5:35 pm

    • what is AEEA and lauroimidazole? Fatty acids and their derivates detect nicely with CAM or phosphomolybdic acid stains. Primary amines stain with ninhydrine.

      Comment by milkshake — December 29, 2009 @ 9:29 pm

      • Soory !

        AEEA = aminoethylethanolamine
        Lauric acid

        Product = Lauroimdazoline…this is a ring structure and hydrolyses easily and produces ring open structures which have amide and amine functionalities..

        thank for your help

        Comment by lulu — December 30, 2009 @ 6:06 pm


    PRODUCT – Lauroimidazoline is the product

    Possibe intermediates are amide and amine based.

    thanking you in advance

    Comment by lulu — December 30, 2009 @ 5:55 pm

  54. Hi,

    I have been trying to separate oligosaachrides generated as a result of enzymatic hydrolysis of CM cellulose, beechwood xylan and D-glucan on TLC plate using various solvents. My positive controls work fine but my hydrolysis products move as smear a short distance from the origin forming rockets. Is anybody out there to help me solve this mystery. Please e-mail me at:

    Comment by ali — January 23, 2010 @ 11:50 pm

    • since you are probably applying your sample on TLC as an aqueous solution you need to be sure the TLC plate is perfectly dry before developing it – water sticks to silica and messes up the separation. I don’t have much else to suggest because I am not familiar with TLC of oligo-sugars

      Comment by milkshake — January 24, 2010 @ 1:05 am

    • if not all of the ionisable groups are of the same charge, then that can cause streaking. eg. if there are two nitrogens and one is positive and the other is neutral then it will streak cause the two move differently on the tlc. adding an acid or a base to the tlc solvent usually helps to combat this because it causes all of the ionisable groups to become either deprotonated or protonated (depending on the addition of acid or base)
      hope that helps a little

      Comment by Illiah — March 15, 2010 @ 5:14 pm

  55. Could you please lemme know how to remove alcohol impurity in organic synthesis

    Comment by shaists aziz — February 22, 2010 @ 6:54 am

    • it depends which alcohol – and what is you material.

      Comment by milkshake — February 22, 2010 @ 11:46 am

  56. helpful!

    Comment by JuvenilePoet — February 26, 2010 @ 4:36 am

  57. in which staining solution is used for chloro methyl acetate, and chloro acetonitrile.

    Comment by charan — March 6, 2010 @ 2:50 pm

    • these are too volatile compounds for TLC. You need a different analytical method to follow your reaction progress – I would suggest to use proton NMR or GC

      Comment by milkshake — March 6, 2010 @ 5:32 pm

  58. does anyone know what i could use to stain peroxides (something specific not a general stain)? i need something that can stain R-O-O-R’ rather than hydrogen peroxide or a hydroperoxide. I tried N,N-dimethyl-p-phenylenediamine dihydrochloride but it didn’t work 😦
    any help would be much much appreciated.

    Comment by Illiah — March 15, 2010 @ 5:10 pm

    • dialkyl peroxides will be lot less polar than hydroperoxides – they should migrate much faster on TLC, thats how you tell them apart.

      Also, dialkyl peroxides will probably oxidize your stain-producing reagent at higher temperature than hydroperoxides so when you heat your TLC carefully you should be able to tell by the order of how the spots gradually develop.

      Have you tried to use o-tolidine (2,2′-dimethyl-4,4’-benzidine, CAS# 119-93-7) for the detection? Also a starch solution with KI and diluted acid might work as a dip

      Comment by milkshake — March 15, 2010 @ 10:25 pm

      • thanks milkshake for your information. I’m in the process of trying the starch KI solution at the moment so I’ll see how that goes. Why did you suggest the o-tolidine? i looked it up when you suggested it as I hadn’t heard of it before but what i found said it was for visualising gold and free chlorine in water.

        I’m not trying to separate peroxides from hydroperoxides so the migration pattern and the speed of development isn’t really applicable to my work but thank you for the information anyway.

        hopefully the KI solution works (i’m filtering the zinc out now as i write this)

        thanks again

        Comment by Illiah — March 15, 2010 @ 11:13 pm

      • o-tolidine is a derivative of benzidine, I used it before for detection of oxidizing species like N-chloro amides. It produces black benzidine dye upon oxidation. Also, if you are trying iodide/starch system make sure your starch is fully dissolved (10 min boil in water) and your iodide solution is slightly acidified

        Comment by milkshake — March 16, 2010 @ 12:34 am

      • thanks for the info about the o-tolidine. i’ll have to give that a go. the starch was definitely all dissolved and the method i used requires 40mL of acetic acid in the KI solution so definitely acidified enough but it unfortunately didn’t work. i’ll give the o-tolidine a go and see if it works any better.

        thanks again

        Comment by Illiah — March 16, 2010 @ 12:47 am

  59. Hi,
    can anyone suggest a good TLC stain for lactone with a hydrocarbon side chain? I tried permangante, vaniline and anisaldehyde.
    thanks in advance.

    Comment by organic chemist — March 29, 2010 @ 10:42 am

    • I would definitely try CAM. Phosphomolybdic acid might work too. Both stains are developed by heating.

      Comment by milkshake — March 29, 2010 @ 2:05 pm

  60. milkshake – there is this spray with diluted hydroxylamine (free base) solution in ethanol, followed by drying with a heat gun, and then developing with diluted FeCl3.hydrate solution. Non-hindered simple esters get transformed into hydroxamic acids and these provide bright-red complex with Fe(3+).
    Personally I have no experience with this system, and I suppose one needs a real hand-spray to make it work (dip jar will probably smear the spots).

    I’ve tried the dip jar with heating to develop and the spots don’t smear all that much, mind you I am detecting ester derivatives of monosaccharides in small quantity….take it for what its worth. I use a procedure similar to that in Anal. Chem. 47, 8, 1975, pg 1420.

    Comment by RD — March 30, 2010 @ 1:12 pm

  61. Hello there,

    I have a question that hopefully can be answered. I want to plate phospholipids on TLC in ng levels. I am considering HPTLC, but after I want to scrape the plate and extract the phospholipids for further analysis on the GC. Is this possible? What dye is the most sensitive that can detect ng levels and at the same time is non-destructive so that I can scrape the plate and do GC analysis (ie. make fatty acid methyl esters)?


    Comment by AFajardo — May 19, 2010 @ 10:24 pm

    • I did my thesis work on phospholipids and I feel your pain. No TLC stain method for phospholipids is too sensitive, and most are destructive – CAM, phosphomolybdic acid or Sn(2+) with molybdate (sold as Sigma phospholipid molybdate detection reagent). I suppose you are doing prep TLC 20×20 cm plate. I would develop it and then carefully cut off a strip along the side, develop the strip by CAM or phosphomolybdic acid, and use it to determine a position of the zones.

      You can do “nondestructive” detection with brief exposure to iodine vapors, to visualise your zones temporarily – but you risk the iodine reacting with some C=C bonds, especially in alkenyl phospholipids (plasmalogenes) and stuff with arachidonic, linolenic acyls so this is applicable only to perhydrogenated samples…

      Comment by milkshake — May 20, 2010 @ 2:09 am

    • I saw a detection stain method used on greasy stuff – by spraying the plate with a diluted aqueous dye – the greasy zones do not soak up the dye as readily as the rest of the plate, so they stand up as lighter zones on dark background. You would end up with a dye in your sample but that should not affect the GC analysis.

      There were two alternatives: a) 0.1% bromothymol blue solution in 10% aqueous ethanol, that is just made basic by few drops of conc. ammonia
      b) 0.2% aqueous rhodamine B (in this case the detection is done both under visible light and fluorescence is observed under long-wave UV.)

      I have no experience with these dye-based systems

      Comment by milkshake — May 20, 2010 @ 2:31 am

  62. Can you give a best stain for aliphatic azides like ethylazide,cyclopropyl azide………………..

    Comment by rambabu pamarthi — June 8, 2010 @ 1:58 am

    • you can use 1:1 ethanol:propargylic aclohol + catalytic amount of Cu(I)Br, azides show up white against yellow background upon heating

      Comment by Mark Johnson — April 16, 2013 @ 4:40 pm

  63. Hello everybody,
    Can anyone please suggest an idea how to separate betamercaptoethanol(oxidised and reduced forms) by TLC……
    Very very urgent please do consider friends!!!!!!!!!!!

    Comment by durga — June 10, 2010 @ 3:12 am

    • I would use some eluent with dichloromethane and a ethyl acetate (you have to try to find the ratios by yourself) and detect with permanganate stain. The oxidized (dimeric) form should be slower on TLC

      Comment by milkshake — June 10, 2010 @ 7:28 am

  64. Hi…
    Can someone suggest me a staining solution for thiols? I will try with KMNO4, but do you have any other ideas?

    Comment by Nina — June 22, 2010 @ 5:42 am

    • KMnO4 would be my first choice but an oxidizing stain – phosphomolybdic acid, or cerium ammonium nitrate would work fine as well

      Comment by milkshake — June 22, 2010 @ 12:56 pm

  65. thank you! 🙂 But they are not specific for thiols, right?

    Comment by Nina — June 23, 2010 @ 6:00 am

    • no they are not – but you probably want to see all components that are present in your mix anyway.

      Comment by milkshake — June 23, 2010 @ 8:11 am

      • hmmmm, unfortunately not really. I’d like to see only the component with the thiol groups. Do you know any specific stains?

        Comment by Nina — June 24, 2010 @ 8:16 am

        • I don’t know – I never used it but there has to be some heavy-metal based stain that makes colorful thiol complexes.

          Comment by milkshake — June 24, 2010 @ 8:59 am

  66. I will search!! thank you!!

    Comment by nina — June 29, 2010 @ 5:01 am

  67. Hello,
    can anyone suggest any imiscible solvent for beta-mercaptoethanol to extract it from a mixture??????????

    Comment by durga — July 14, 2010 @ 1:07 am

    • dear durga, this is already your fifth (and I am sure it will be the last) comment about mercaptoethanol. I just deleted four preceding comments from you and the replies to them. Please take it easy. I would have to spam-filter you if you don’t stop.

      Comment by milkshake — July 14, 2010 @ 3:33 am

  68. Hi,

    this is rambabu pamarthi woorking in GVK bio. Can you provide best strain list for aliphatic compounds

    Comment by Rambabu pamarthi — July 28, 2010 @ 3:24 am

    • CAM would be my fist choice, phosphomolybdic acid the second

      Comment by milkshake — July 28, 2010 @ 7:51 am

  69. could anyone please suggest me the staining agent for organic sulfides like phenylmethyl sulfide

    Comment by Deepan — July 28, 2010 @ 8:48 am

  70. I’ve used bindone solution in DCM (perhaps a poor choice of solvent, but seemed to work fairly well in my case) to distinguish between N-mono- and N,N’-disubstituted piperazine. Monosubstituted one gave pink to purple stain on very light pink background (fairly weak stain, but clearly visible under UV lamp); disubstituted piperazine didn’t show up. I believe it should work with other secondary, and probably primary amines as well, but I have never tested that.

    Bindone was picked because I had a lot of it laying around, as it was a reagent for some other synthesis. Since it is sometimes used as analytical reagent for amines in solution, I decided to give it a try on TLC as well. No heating necessary in this case, in fact stains disappear on heating.

    Comment by Baltic — August 2, 2010 @ 9:08 pm

  71. Hi,
    could you please suggest me the staining reagents for secondary amine like N-tertbutylaniline?

    Thank you very much

    Comment by Surya — August 5, 2010 @ 10:19 am

  72. Hi,

    Would anisaldehyde be the prefered stain for detecting sugar-1-phosphates. And can it distinguish the sugar-1-phosphate from sugar?
    Thanks in advance.

    Comment by Helena — August 25, 2010 @ 4:51 am

    • it is quite difficult to do chromatography of sugars (on silica) because the polyhydroxy compound bind so strongly (and sugar phosphates will be even worse). I believe there were old paper chromatography methods with very polar solvent mixtures – are you using those?

      I would employ some nonspecific detection to detect everything, like anisaldehyde stain. Then, to distinguish the stuff with phosphates I would use “molybdate reagent” for detection of phosphates – i don’t know the composition of this reagent but Sigma and Fluka are selling the ready-made molybdate reagent. You spray it on the TLC, it develops blue spots after few minutes at room temp (no heating necessary) with stuff containing phosphate esters. I used it for detection of phospholipids. Also, with some nonspecific stains you can re-stain TLCs after you had stained them with molybdate stain, that way you need to do only one TLC.

      Comment by milkshake — August 25, 2010 @ 11:21 am

      • what is mobile phase for sugar

        Comment by Bhaiyyasaheb — July 31, 2017 @ 12:16 am

  73. Question: What spray reagents would you recommend for cyclodextrins on TLC? TIA.-

    Comment by Tapira1 — October 1, 2010 @ 5:10 pm

    • I think CAM or phosphomoybdic acid should work fine, but you can try anisaldehyde as well

      Comment by milkshake — October 2, 2010 @ 2:53 pm

  74. Can you please tell me, how to stain compounds having phosphate groups?
    what is staining agent to detect Phosphate group?

    Comment by Medchem — October 7, 2010 @ 3:48 pm

    • use the ready-made “molybdate reagent” for detection of phosphate esters from Sigma or Fluka. See also my reply to #72 above

      Comment by milkshake — October 7, 2010 @ 4:13 pm

  75. Hello

    I want to stain my compound cantaing N-methyl,N’-chloroprppyl piparezine.Please sugest me proper stain for devolepment of the same.

    Thanks and Regards
    M T Shaikh

    Comment by M T Shaikh — October 11, 2010 @ 5:16 am

    • I would use permanganate – it works very well for staining tertiary amines because they oxidize so easily – it is almost instant. (Primary and secondary amines stain too but it takes often several minutes to develop)

      Comment by milkshake — October 11, 2010 @ 10:53 am

  76. Can anyone think of a stain other than KMnO4 that will work for alkyl halides (mostly bromides)? Reverse phase TLC plates don’t tolerate KMnO4 too well.


    Comment by R Bakus — October 27, 2010 @ 5:56 pm

  77. can you plz tell me a stain to separate thiol(toluene p-thiol) from an aryl thioether(phenyl p-tolyl sulfide)?
    Thank you very much

    Comment by Supuni Duneeshya — November 24, 2010 @ 5:43 am

    • thiophenols extract into aqueous NaOH as thiophenolates. You probably want to deoxygenate your mixture first, by argon sparge before adding NaOH, to limit the air oxidation of thiophenolate

      Comment by milkshake — November 24, 2010 @ 1:01 pm

  78. Hey my work is to prepare triacetin from glycerol using acetic anhydride.. Can any one tell me The appropriate solvent mix in order to monitor the reaction progress towards triacetin

    Comment by jass — January 31, 2011 @ 9:36 am

    • I don’t know because I have not done it myself but I suppose usual mix of ethyl acetate with hexane would work. I would try first maybe hex-EtOAc 4:1 and if this was too slow I would add more EtOAc to it but if it its too fast I would go with more hexane there. You should see the product in front and with monohydroxy trailing behind, followed by dihydroxy.

      This sounds like a typical student lab assignment for practicing TLC monitoring of a reaction – so I am kind of reluctant to help you with this too much because you are supposed to work on it on your own. But I should mention that you probably want to get rid of acetic acid from your sample first (because it would mess up the TLC separation). The way to do this is to take a sample from reaction mix, put it in a vial or a small test tube and blow a stream of nitrogen or air over it for a minute until it stops smelling like vinegar. Then you dilute your sample with few drops of some less polar solvent like dichloromethane or chloroform and spot it on your TLC

      Comment by milkshake — January 31, 2011 @ 8:26 pm

  79. Can some body tell how to stain n-substituted pyrroles.

    Comment by alex joseph — February 16, 2011 @ 5:31 am

    • since pyrroles oxidize so readily, permanganate or CAM should work for detection. But first look at them under UV

      Comment by milkshake — February 16, 2011 @ 6:05 am

  80. very good

    Comment by kalyani — March 27, 2011 @ 1:10 pm

  81. Can anybody tell me any solvent system for getting good resolved phorbol ester on TLC plate

    Comment by Anjali Bose — April 11, 2011 @ 7:13 am

  82. I am working with isolation of bio-active compounds from marine streptomyces strain. I did the general stain with vanilin and anisaldehyde for the purified active compound. On addition to my previous analysis i suspect that there is a sugar derivative attached to the compound.

    How can i visualize such sugar compounds attached to polyketide backbone ??? i will be very happy if i find views about this !!!!?? pls help

    Comment by Niraj Aryal — May 28, 2011 @ 11:24 am

    • I would use CAM stain – it is sensitive and very general; you can also try basic permanganate – KMnO4 should work but CAM is far better

      Comment by milkshake — May 28, 2011 @ 8:25 pm

  83. What stain would you recommend for hydroperoxide compound? 4,5-dihydro-1,3-dioxepin-5-yl hydroperoxide

    Please help!!

    Comment by Yong Jung Shin — June 28, 2011 @ 7:14 pm

    • I have not worked with peroxides much, I only did iodometric titration of tBuOOH solutions. (KI, AcOH, starch indicator, titrated with Na2S2O3 stock solution) so I don’t have the experience – but I wonder if you can dissolve small mount of starch in boiling water, add iodide, cool – and just before use acidify with AcOH. Hydroperoxides should produce I2 from acidified iodide which should stain starch solution deep blue

      Comment by milkshake — June 29, 2011 @ 1:03 pm

  84. I always come back here for any TLC tips and tricks, thanks to milkshake for the advice!

    I noticed an older post about detecting thiols by TLC and would recommend using Ellman’s reagent for detection; it reacts with thiols to give a a bright orange color. Prepare a stock solution in buffer and then dip the plate; this is useful for us because we do alot of peptide work.

    Here are some related references:

    Ellman, G. L. (1959) Arch. Biochem. Biophys. 82, 70-77.
    Bulaj, G.; Kortemme, T.; Goldenberg, D. P. (1998) Biochemistry 37, 8965-8972.

    Comment by RD — July 4, 2011 @ 3:44 pm

  85. Does any one have a good stain for boronic acids which also contains an alkyl azide functionality.

    Comment by Sara Chirayil — July 27, 2011 @ 12:23 pm

  86. Does any one have experience in using TLC for detection of Magnesium L-Threonate? If so please tell me the mobile phase and the staining reagent.

    Comment by Richard Lin — September 11, 2011 @ 3:55 am

  87. What is a proposed prodecure to detect in TLC a poly(methyloxazoline) polymer, that is containing tertiary amide groups in the backbone with the amide oxygen in the side chain? ..CCN(COCH3)CCN(COCH3)CCN(COCH3)CCN(COCH3)..
    Thanks for hints

    Comment by Pat — September 23, 2011 @ 8:16 am

    • this one may be quite hard to stain but I would try iodine vapor chamber – I have had very good results with staining PEG polymers on TLC with it

      Comment by milkshake — September 23, 2011 @ 12:20 pm

  88. Hi!

    Can you propose how to stain aldehydes in TLC by 4-hydrazino-7-nitrobenzofurazane? Can’t find suitable protocol!
    Thank you very much!

    Comment by Ksenia — September 25, 2011 @ 1:55 pm

    • Although I have no experience with using this hydrazone-making reagent my guess is that the stain solution composition is going to be very similar to the dinitrophenylhydrazine stain for aldehydes: you make a diluted solution of the hydrazine in a very diluted HCl (maybe 1%) in water or in ethanol. And you use it without heating the TLC

      Comment by milkshake — September 25, 2011 @ 4:44 pm

  89. Hi, this page is really useful, I’m working on my thesis with N-sustituted thiazolidine-4-carboxylic acid (could be tetriary amides or amines). Can you suggest a stain solution for it?

    Comment by andungtrang — November 28, 2011 @ 12:06 pm

    • since you have this easy-to-oxidize sulfur atom in your molecule, I think any general-purpose oxidizing stain should work well: CAM, permanganate, phosphomolybdic acid…

      Comment by milkshake — November 28, 2011 @ 2:20 pm

  90. Hey !
    I have a really stupid question but I thought I should give it a try 😉
    can anyone explain me what does the chlorine/TDM exactly detects???
    Thanks in advance.

    Comment by neymara — January 16, 2012 @ 4:06 pm

    • No I think it is a good question. Although I have not used it, I believe it is a closely analogous system to one that I used in a peptide group long time ago, for nonselective staining of peptides. Th system I used was a spray with t-butylhypochlorite solution in CCl4 followed by driving off the excess of the chlorination reagent with a hair-drying gun, followed by staining with 3,3′-dimethylbenzidine. You chlorinate all NH groups (amines, amides) and then remove the excess of chlorine absorbed on TLC so as not to have a background. Then the produced chloramides and chloramines (which are not volatile) are revealed on TLC by spraying with some easy-to-oxidize reagent that produces a deeply colored oxidation product with Cl+ source. I think derivatives of benzidine are used for the purpose because they produce deep blue-black quinoid oxidation products.

      Comment by milkshake — January 16, 2012 @ 5:12 pm

      • Thanks a lot for your reply 🙂

        Comment by neymara — January 17, 2012 @ 12:13 pm

  91. i want to know about the proportion to be taken foe phenol saturated in water for the thin layer chromatography . as a mobile phase

    Comment by shaan nak — January 23, 2012 @ 2:50 am

  92. Hi Guys,

    Trying to visualise hydantoins, No UV254, CAM (very faint white shadow then disappears !) Permanganate (no reaction), PMA failed, iodine v. weak spots. Any suggestions ?
    Thanks in advance

    Comment by Andy L — February 10, 2012 @ 12:05 pm

    • If your hydantoins have a free NH your best bet would be a spray with a diluted solutioun of t-butyl hypochlorite in some chlorinated solvent (in old days I used CCl4) followed by drying with cold hair drier (to pass the sniff test – until the TLC no longer smells of chlorine) followed by a spray with 3,3′-dimethylbenzidine (aka “2-tolidine”) – N-chloro compounds generated in situ will produce dark spots on light background. There are version of this detection system that use chlorine gas and TDM, see the comment #91

      Comment by milkshake — February 11, 2012 @ 11:03 pm

  93. Thanks MS, i’ll check if that’s ‘compatible’ with our safety guys and give it a go !

    Comment by Andy L — February 13, 2012 @ 4:56 am

  94. Hey, could someone comment on a good way to stain a very hydrophilic phospocholine (a quarternary amine, 06:0 carbon chains, water soluble)? I’ve tried copper staining the phosphate, Nile-Red to detect the fatty acid, Phosphomolybdic acid, KMnO4 and even conc.H2SO4… the only thing that detects anything at all is iodine. Problem is I’d need to detect ng quantities…

    Comment by fran — February 22, 2012 @ 12:35 pm

    • You may try Dragendorf reagent (the version of stain that uses tartaric acid, the mixed solution is stored in a fridge) to detect the quaternary ammonium and all other amines (R.T., brown spots on pale yellow background). For selective phosphate detection there is “Molybdate Blue” phospholipid stain sold by Sigma that detects phosphate esters (R.T., blue spots on white background) – I dont know what is in this reagent (apart from molybdate salt and diluted sulfuric acid)

      Comment by milkshake — February 22, 2012 @ 2:53 pm

  95. Hi guys,

    someone has an idea of staining epoxides?

    BR, Markus

    Comment by Markus Loeweneck — April 16, 2012 @ 12:03 pm

    • Hi Markus,
      try KCNS. It will open the epoxyde and insert the CNS and give a nice UV-chromophore, should be visible with a fluorescent indicator. It can be used for precolumn derivatization for HPLC (RP). The disadvantage: Two reaction products, however in a reproducible ratio. I never tried it for TLC. However it reacts in aqueous solutions, so it may be used in reversed phase TLC. You can use the peak ratio to discriminate unknown epxides. I used it to characterize the epoxide hydratase reaction with fatty acid epoxides. An alternative are stains with reactive -SH.

      Good luck

      Comment by Klaus — April 27, 2012 @ 7:44 am

      • I’ve seen Picric acid mentioned as a chemoselective TLC stain for epoxides. Probably in J. Chrom. A. Hope this helps

        Comment by Dan — November 16, 2012 @ 3:51 am

  96. I am unable to find exactly how to detect amides in which the amine group is tertiary ?

    Comment by hemender — April 19, 2012 @ 11:19 am

    • I can’t think of any good method. You may try some nonspecific detection, like iodine vapor chamber

      Comment by milkshake — April 19, 2012 @ 1:21 pm

  97. hi can any one tell me the stain for indolizidines like 209d?

    Comment by wish — June 15, 2012 @ 1:15 am

  98. hi,
    this blog is really superb.
    I have three N-methylpyridinium moieties in my compound, can any one suggest me stain for this?

    Comment by Anu — July 9, 2012 @ 6:38 am

    • I would try UV detection and Dragendorff dip (the version with iodobismutate + tartaric acid)

      Comment by milkshake — July 9, 2012 @ 12:55 pm

  99. wow! it’s first that says about ninhydrin reaction of boc-protected amines!!
    Could you say the original reference or source paper about ninhydrin reaction with boc-protected amines?? if you have or remember…:)

    Is there any method to find tertiary amine with ninhydrin? in my experiments(in class), TLC with ninhydrin staining shows unexpected spot, and i think it only can be tertiary amine that i put to make basic condition,…..Unfortunately, i lost my ninhydrin solution’s information… so i can’t say that my ninhydrine was in special condition….
    i couldn’t find any method or mechanism of ninhydrin with tertiary amine, and some websites says that ninhydrin can show tertiary amine on TLC plates(not with exact references, methods)…. such as this one:

    is this just wrong or no-source information? plz say me your opinion or ideas…:)

    Comment by Kim — October 7, 2012 @ 3:49 am

  100. hi.

    i am currently working on separation and purification of hexane (non-polar) fraction of a marine seaweed, of which i’ve dissolved in chloroform (semi-polar) solvent and separated via column chromatography to 42 subfractions (with a 5% increment, from 100% hexane [non-polar] to 100% chloroform [semi-polar], continues to 100% methanol [polar]).

    In this case, assumming that most of the compounds would be non-polar and partially would be semi-polar compounds, what types of dyes i should use for the TLC separations?

    Thanks a lot!


    Comment by Tony Quah — October 30, 2012 @ 8:15 am

    • I would use CAM – it is typically very sensitive and works great for greasy substances. You need to use glass-backed silica TLC plates and heat them quite high though because the detection is based on mineralization of everything organic.

      Comment by milkshake — October 30, 2012 @ 2:28 pm

      • Hi!

        Thanks for your prompt replies. However, is using glass-back plates a must? We have those aluminum ones but no glass ones… 😦



        Comment by Tony Quah — October 31, 2012 @ 4:36 am

        • aluminum-backed TLC plates fall apart if you heat them with a detection-solution that contains H2SO4, like in case of CAM. If you must use Al-backed plates, use phosphomolybdic acid detection. But CAM works better

          Comment by milkshake — October 31, 2012 @ 1:19 pm

          • Thanks a lot! will try to work and see how’s the progress… 🙂

            Comment by Tony Quah — November 1, 2012 @ 1:40 am

  101. A few questions and comments:
    I’ve had no problems using CAM with aluminium backed plates. I do however find that with this (and other aqueous based) stain permeation of the silica layer is poor and I get “patterns/ripples” of blue colour on occasion. I was wondering if there was a way around this, but I guess not as adding anything organic would kill off the stain pretty quick I’d imagine. (I guess a I’m drawing a comparison to the difference between aq. FeCl3 and FeCl3 solution made up in 50% aq. methanol in wetting ability of the plate).

    A former colleague of mine used to make up his vanillin stain using acetic acid in addition to the usually prescribed H2SO4, claiming that it made the stain last longer/affected staining ability (could be either or both I guess). Unfortunately I’m no longer there anymore but wondered if anyone has seen or has experience with a similar recipe? I’m also intrigued as to the real difference in staining ability between vanillin and anisaldehyde, and to a certain extent how these compare with p-dimethylaminobenzaldehyde. All three reagents are electron rich aromatic aldehydes, so I suspect they also stain reasonably similar. I also wonder about the comparitive ability of acidic and basic vanillin – how different are these two reagents (really) and in what sort of situation would you opt for one over the other.

    Finally, I cannot find a recipe for the Dragendorff reagent that is supposedly stable once made up (refrigerator). If no-one has one then would it be prudent to make up a solution as detailed here: under “ready to use stains”? I guess it has to be relatively stable for them to offer it as a ready-made consumable.

    Thanks in advance for any advice


    Comment by Dan — November 16, 2012 @ 4:07 am

  102. Hi,

    I recently sprayed a TLC plate with Dragendorff’s reagent and a spot which is not visible under UV nor to the naked eye turned into dark blue (with a very slight orange) color. What could be the possible type of compound for that spot? The compound was obtained from a plant sample after the acid-base treatment step. It might be an alkaloid but I couldn’t understand why it it stained dark blue with Dragendorff’s reagent. Any comments?

    Comment by Kuan-Hon — August 7, 2013 @ 11:36 am

    • I wonder if the blue-staning spot could be some oligo/polysacharide. There is always some I3(-) present in Dragendorff, from air oxidation of iodide in slightly acidic solution, and iodine (in form of triiodide) famously forms very deep blue complexes with starch – starch is actually used as indicator for iodine

      Comment by milkshake — August 7, 2013 @ 12:06 pm

  103. I want to know if there are any other alternative to Anisaldehyde and Vanillin, that I can use to detect eucalyptol on TLC plate? Also, what is the basic chemical reaction that causes the distinctive coloration by Anisaldehyde on eucalyptol?

    Comment by Shrestha — October 1, 2013 @ 6:35 am

    • you got me – I don’t know. I suppose eucalyptol decomposes upon heating with diluted sulfuric acid (present in anisaldehyde stain) and the produced unsaturated product undergoes some condensation reaction with anisaldehyde. For alternate stain to anisaldehyde – Please have you tried CAM?

      Comment by milkshake — October 1, 2013 @ 9:40 am

  104. What would be a good stain for glucosinolate compounds, both aliphatic and indolic, from a plant extract?

    Comment by Craig — October 24, 2013 @ 12:15 am

    • permanganate or CAM stain or phosphomolybdic acid or anisaldehyde should all work well in this case. I would maybe try CAM first, since it is so sensitive and the staining is reasonably time-stable. Anisaldehyde (or vaniline) stain has the advantage that different glycosides might produce different colors, so complex mix might be easier analysed if the constituents of the mix nearly co-spot. Since your thioglycosides are pretty water-soluble, getting hold of hand-held spray for TLC might be better option than dip-staining with jars of stain which could cause smearing of the developed TLCs. But spraying is little messy – make sure to place a disposable cardboard box in your hood where you do the spraying, to keep the mess within, otherwise the back of your hood will get pretty colorful soon

      Comment by milkshake — October 25, 2013 @ 12:09 pm

  105. please sugest the solvent system for TLC to determine piperazine secondary amines)

    Comment by jay — February 8, 2014 @ 6:08 am

    • if the piperazine is greasy – has few substituents – Dragendorff would be the first choice, it detects all amines except for the very hydrophilic ones.

      Permanganate stain stains tertiary amines fast (they oxidize much faster than secondary amines) but it will detect secondary amines if given enough time.

      Ninhydrine detection of secondary amines is weak. Maybe you can try chloranil test for secondary amines: first spray with 2% acetaldehyde in DMF, then spray with 2% chloranil in DMF (blue spots appear in the presence of secondary amines). The used DMF has to be good quality (free of fishy-smelling amines) and the detection solutions should be kept in fridge only for 1-2 weeks because they decompose easily especially on sunlight.

      Then there is spray system for detection of secondary and primary amines, amides, urethanes, sulfonamides, ureas – anything with free NH – that uses a spray with solution of tBu-hypochlorite in CCl4 (or it can be replaced with chlorine gas chamber) followed by drying with cold air followed by 3,3′-dimethylbenzidine (aka tolidine) spray. You get black spots on white background and the detection is very sensitive

      Comment by milkshake — February 8, 2014 @ 8:27 pm

  106. Does anyone know how Rydon-Smith spray is made. Additionally is there a good replacement for this. I am trying to evaluate Pepstatin A on TLC plates.

    Comment by Yossi Singer — February 27, 2014 @ 3:40 am

  107. what is the best tlc stain for sialic acid derivatives appended with acetates or OMe,etc, I tried PMA, anisaldehyde, its not working well

    Comment by rk — April 30, 2014 @ 10:12 pm

  108. What is the best stain for diphenylmethanol that can be used for quantitation? I’ve tried permanganate, which doesn’t give distinct spots. Iodine gives a faint brown spot, but it disappears quickly. I’ve also tried anisaldehyde and vanillin-sulfuric. Also didn’t work.

    Comment by Pio — May 14, 2014 @ 9:03 am

  109. Will be happy if you can help me with this problem
    I have got a bright pink colored rocket shaped spot on TLC after spraying with anisaldehyde sulphuric acid reagent. this spot is UV negetive but Iodine positive. What class of compound could it be?

    Comment by anupama — September 3, 2014 @ 12:40 pm

    • it could be lots of things, anisaldehyde and iodine are general (non-selective) stains. Since the spot is not UV active it probably does not have any aromatic rings but that is as much as I can tell

      Comment by milkshake — September 3, 2014 @ 12:57 pm

  110. what is the best replacement for anisaldehyde? Our school does not have that reagent. thank you.

    Comment by Francesca Ong — January 28, 2015 @ 10:47 am

  111. If the vanilin dye is made with conc HCl and MeOH, will that work?

    Comment by Kh Tanvir Ahmed — February 4, 2015 @ 6:05 pm

    • Most likely it will not work because HCl is volatile, unlike sulfuric acid which stays on the TLC and helps to carbonize your compound upon heating. You may want to try the original stain recipe before modifying it. But if you want to play, nothing prevents you from making whole lot of TLC alternate stains, dip TLCs into them and see which one works best.

      Comment by milkshake — February 4, 2015 @ 7:48 pm

  112. I always find tour posts immensely helpful in my work.
    While performing TLC separation, I noticed some spots that are UV negetive but stain with iodine. Wonder what they might be?With anisaldehyde sulphuric acid reagent these spots give bluish purplish colour.

    Comment by Anupama — February 10, 2015 @ 4:20 am

  113. I was wondering if it were possible to use the “molybdate reagent” to detect G1P on a PEI-cellulose TLC plate. I switched the stationary phase from silica gel to PEI-cellulose in hopes of having a more robust detection and separation of the sugar nucleotides I’m studying. However, since G1P is not active under UV light, I am having trouble determining an appropriate staining method.

    Comment by gochrisjo — March 25, 2015 @ 1:05 am

    • I think this should work, definitely give this a try. I cannot predict whether polyethylene imine won’t interfere with molybdate, by forming some insoluble salt, but there is a good chance it will be fine, the molybdate stain is pretty selective to phosphate esters

      Comment by milkshake — March 25, 2015 @ 1:53 am

  114. what is the suitable staining agent for 2-(hydroxy(phenyl)methyl)acrylonitrile?

    Comment by Suhasini — April 28, 2015 @ 3:57 am

  115. Hi, anyone knows how to prepare the staining solution of Ellman’s test? Is there any different way aside using buffers (I’ve seen Tris-HCl or phosphate)?

    Comment by PabloQ — May 14, 2015 @ 9:45 am

  116. Hi, could you suggest me any test to confirm methoxy groups?

    Comment by velu — July 27, 2015 @ 4:43 pm

    • Not really. What kind of methoxy groups – ester or ether?

      Comment by milkshake — July 27, 2015 @ 5:04 pm

  117. Hi.
    Can anyone help me with stains for aliphatic bromo compounds..having no other functionality and others having a bromo and amide bond

    Comment by pracs — November 9, 2015 @ 8:31 pm

  118. Hey Milkshake, for the “improved Dragendorff’s” recipe, can I get away with DL-tartaric acid or is L(+)-Tartaric acid truly necessary? I can’t imagine why DL-Tartaric acid wouldn’t suffice, but then again I haven’t the foggiest idea how this reagent works…

    Comment by Ian D. — December 24, 2015 @ 8:45 pm

    • I don’t know, I have not tried it with a racemate. The only differences can be in solubility – racemic compounds often have surprisingly different solubility when compared with enantiopure compounds. I would give it a try, see what happens.

      Dragendorff works by making deep-orange salts of iodobismutate anions with large cations – i.e. quaternary ammonium cations, alkaloids and other greasy amines. It also detects suprisingly well with crown ethers and larger polyethyleneglycols – most likely by precipitating salts with K+ complexed to crown ethers or PEGs.

      I believe tartaric acid is there as a buffer (source of H+) and as a coordination agent that keeps iodobismuthate from decomposing

      Comment by milkshake — December 25, 2015 @ 12:18 pm

      • Thanks, I haven’t used the tartaric acid recipe before, but Dragendorff’s (along with CAM/Hanessian’s and even iodoplatinate) work great for staining my current compounds…mainly PEGylated 2-arylbenzothiazoles and CuAAc “click” conjugates thereof…its surprising how inert some heterocycles are with the more common TLC stains.

        Also…do you have any experience running manual RP-18 flash columns? My compounds are exceedingly difficult to purify by silica once conjugated to anionic/zwitterionic substituents like sulfonated cyanine fluoropores (i.e. sulfo-Cy5-azide). I’ve been using RP-18 preparative TLC as we don’t have access to preparative HPLC at the moment. I’ve got a 25g sample of C18 resin from Silicycle (cat# R33230B) that they recommend for “manual” flash chromatography, but I’ve been hesitant to use it.

        Comment by Ian D. — December 27, 2015 @ 6:08 pm

        • I don’t have experience with doing reverse phase prep LC on CombiFlash/Biotage-like systems, but people who tried it did not like it (compared to prep HPLC), and so they tried it just once and cursed it…

          Comment by milkshake — December 29, 2015 @ 1:32 am

  119. plz suggest a suitable tlc derivatizing agent for amides

    Comment by shilpi — January 7, 2016 @ 1:47 pm

    • amides in general are hard to visualize but if they have free NH (amide, amine) the binary detection system with tBuOCl (or chlorine gas) followed by spraying with o-tolidine would work. This is somewhat nasty and tricky so use this only if there is no other better alternative

      Comment by milkshake — January 7, 2016 @ 2:08 pm

  120. what stain is useful for imine compounds.

    Comment by vibha tiwari — March 25, 2016 @ 12:27 am

    • imines are not very stable on TLC, you would often see their decomposition products on silica, although some benzophenone imines can be TLC-analyzed, especially with TLCs pre-treated with triethylamine.

      Ninhydrin (with a little heating) should work, to detect the starting amine. Any oxidation-based detection should also work (basic KMnO4, CAM). But I think it is not good to monitor imine formation on TLC because of instability, I think you would be better off by using NMR

      Comment by milkshake — March 25, 2016 @ 10:16 am

      • ok thankyou

        Comment by vibha tiwari — March 26, 2016 @ 9:20 am

  121. hi, what visualizing agent I can use to see the Sulphur staining especially used in rubber industy

    Comment by Ranthi — April 6, 2016 @ 2:42 am

    • I would use UV detection (sulfur compounds usually absorb UV very nicely) and then some oxidative stain – CAM, phosphomolybdic acid or basic permanganate

      Comment by milkshake — April 6, 2016 @ 3:16 am

  122. what visualizing agent can be used for 1-bromopentane??.i have tried iodine but is not visible….

    Comment by alia javaid — April 14, 2016 @ 9:11 am

    • 1-bromopentane it is too low boiling for TLC (b.p. 130C), you need to use some other analytical method

      Comment by milkshake — April 14, 2016 @ 9:45 am

  123. Hi,

    Which TLC stain is good for the Silyl or Boryl compounds?

    Comment by Ajit Kale — April 15, 2016 @ 6:42 pm

  124. Hello! I could use some help! I am trying to stain my TLC polar lipid plate with Phosphomolybdic acid and after heating, there is a white streak in the middle of the plate! After I spray though, the entire TLC plate is yellow, and then once I place in the oven, the plate turns white. I am using ethanol to dissolve the crystals, so I don’t understand what the issue is. Any tips?

    Comment by Microbiology — June 27, 2016 @ 2:15 pm

    • I don’t know, I have been doing TLCs of phospholipids with phosphomolybdic acid detection but never had this problem. I wonder though – does your elution solvent system contain ammonia or some other amine? If yes, maybe you should try to pre-heat the TLCs after elution (before the detection), to make sure all amine is gone from the silica before you start spraying it with TLC. If I were to make a guess, there is maybe something in your mobile phase used for TLC elution elution that is killing PMA

      Comment by milkshake — June 27, 2016 @ 5:12 pm

      • Hi! I can give that a try! Thanks for the tip. My solvent system consists of chloroform, methanol, and water for the first dimension, and chloroform, methanol, acetic acid, and water for the second dimension. A second problem that I seem to have developed, and that all of my spots are riding with the solvent line. When I spray my plates, I can see the spots clumped at the top of the plate right with the solvent line. For some reason, the spots do not seem to separate. I have made sure to use fresh reagents and even changed tanks. I have gone from having beautiful spots to now this problem. I would appreciate any advice!

        Comment by Microbiology — June 27, 2016 @ 5:17 pm

        • I have seen with phospholipids (that are very polar and strongly bound to silica) that if there is a large amount of neutral lipids, the phospholipids can ride their front. The way to fix this problem was to pre-develop the TLC in peroxide-free ether first, that just moved the blob of neutral lipids out of the start, let the plate dry for few minutes on air without heating for ether to evaporate – and then run the TLC with the usual elution system for phospholipids

          Comment by milkshake — June 27, 2016 @ 7:17 pm

          • Hi! So, I place the TLC plate in the ether first, prior to loading it with my sample correct? Can I use diethyl either? Also, the funny thing is, I have used the organism before and I got really nice lipid profiles. I even sent the extract giving the weird run to another lab and their profile turned out fine, so something is going wrong with my run. Is there something I could be doing during extraction that gives me inconsistent results between the different runs?

            Comment by Microbiology — June 28, 2016 @ 1:10 pm

          • what I meant was to develop the TLC with diethylether first.

            Also, are the TLC plates the same as before? The mobile phase – was it freshly made, or was it aged in the bottle? Have you checked that the mobile phase did not become biphasic on a warm summer day? (That would really screw things up). Since your method worked fine before and now you have these strange results, there is probably some simple cause for it, one factor that is not under control.

            Comment by milkshake — June 28, 2016 @ 1:22 pm

  125. I have methyl ester protected glutamate-Urea-lysine molecule. I have converted amine group to formamide and i used ninhydrine for confirmation of conversion, but when i carried out conversion of formamide to isocyanide, ninhydrine is not working? which stain would be better for isocyanide. finally i have to deprotect the methyl ester group to make carboxylic acid free.

    Comment by Nadiim — December 1, 2016 @ 5:08 am

  126. Anyone use iodine to stain TLC for a mixture of hydrocarbon oil, polymers, and antioxidant? Any suggestion for polymer blend TLC before Cyclographing. Thank you.

    Comment by John T — February 15, 2017 @ 11:02 am

  127. what is best stain of Polyethylene glycol?

    Comment by Nadiim — April 3, 2017 @ 1:28 am

  128. what stain will give indication for nitrile group containing compound?

    Comment by Bijan Mohon Chaki — June 27, 2017 @ 2:29 am

  129. Dear Sir,
    which mobile phase will be best for polyethylene glycol (PEG 10000)

    Comment by Bhaiyyasaheb — July 31, 2017 @ 12:14 am

    • chloroform-methanol 100:10

      If you need to improve separation of PEGs with carboxy groups or amino groups at the end of the chain, you can add 1% (by volume) of aqueous ammonia or formic acid to the mobile phase, so then it becomes 100:10:1.
      Please can you tell me what are you trying to do? Maybe I could help you because I have made quite a few functionalized PEGs. Obviously you can purify PEGs by chromatography only on a small scale, 1 g or so.

      Comment by milkshake — August 1, 2017 @ 10:40 am

      • Dear Sir,
        we are trying to separate PEG 4000, PEG 6000 and PEG 10000 by TLC method.and which solvent can i used for sample dissolution

        Comment by Bhaiyyasaheb — August 2, 2017 @ 5:21 am

        • unfortunately PEG separation by molecular weight alone by TLC is not going to work – you need to have a difference in functional groups at the end of terminus. From my experience I can tell that smaller Mw PEGs move slightly faster on silica than larger Mw but the difference between PEG 4000 and 6000 is simply not enough. You would need a GPC with gel a column for that

          Comment by milkshake — August 2, 2017 @ 2:07 pm

          • By GPC Different type of PEG molecule can be detected? or average molecular weight detected.

            Comment by Bhaiyyasaheb — August 9, 2017 @ 1:24 am

  130. Hi,

    Anyone knows how to detect secondary aliphatic amines through TLC.?

    Comment by aNSHIKA — March 6, 2018 @ 12:53 am

    • the easiest would be to use Dragendorff stain (the recipe is above) it detects all basic compounds, and also crown ethers and some phosphines

      Comment by milkshake — March 6, 2018 @ 1:02 am

  131. Does anyone have a reference to the statement above? “Boc-protected primary amines produce spots on heating (as the Boc falls off)” I just started as a grad student and boc protecting primary amines is my first project which is giving me trouble.

    Comment by matt m — June 29, 2018 @ 5:42 am

  132. Hi, can anyone suggest a stain for nitrogen heterocycles (triazole, pyrazole, imidazole) susbtituted with large aliphatic groups like adamantane or norbornylene?
    I’ve tried CuCl2 in acetone, vanillin and iodine so far, wasn’t satisfied with the results.

    Comment by dmitry p — January 7, 2019 @ 11:02 am

    • I think imidazole will be basic enough that you should have success with Dragendorff stain with tartaric acid (has to be stored in the fridge, the recipe is above) but I am not sure if basicity of pyrazole or triazole will be sufficient although it might (though for example PPh3 is basic enough, apparently, to be detected). Dragendorff stains anything that forms very large cations, by precipitating dark brown iodobismutate salts from aqueous solution. (Mysteriously, 18-crown-6 and PEGs are are also staining, probably due to their complexation to K(+) present in the stain.)

      Comment by milkshake — January 7, 2019 @ 12:17 pm

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