I use hydrochloric acid to release a steroidal sapogenin from its saponin, then neutralise it with sodium bicarbonate. I am confident the hydrolysis works because the mixture no longer produces a persistent foam.

I know, from the literature, that this sapogenin is poorly soluble in water but very soluble in chloroform, so I phase extract the aqueous mixture with chloroform. I send the chloroform layer to my collaborator who evaporates it and resuspends in deuterated chloroform to do NMR. I expect several milligrams of the sapogenin but he has been unable to detect any. I'm losing my sapogenin. What am I doing wrong?

Some details and observations:

1) I add 2mls of concentrated (10N) hydrochloric acid to 8mls of the saponin which is in 45% ethanol (it's a “tincture”) so gets diluted to 36% ethanol when the hydrochloric acid is it added.

2) This (10ml) 2N reaction is incubated for 4 hours at 80C in a sealed polypropylene tube.

3) After incubation, the entire 10mls is poured into a flask containing 3 grams of sodium biocarbionate and mixed. Once the foaming (caused by release of carbon dioxide) has stopped, I pour the material into a new polypropylene tube. A lot of salt (NaCl) is produced from the neutralisation step. This salt and unreacted sodium bicarbonate settle to the bottom. I assume the solution is saturated with sodium ions.

4) Copious amounts of a dark red precipitate forms – even under conditions using much less acid, such as 0.01N reactions. When the solution is spun in a table top centrifuge, it produces a pellet that is about 1.5ml (on top of the salt formed by neutralisation) from 10mls of solution. The supernatant (from this 0.01N reactions) still produces a persistent foam. The red pellet, suspended in water, does not.

Not surprisingly, reactions at 2N also produce this precipitation but it is much darker – like a dark purple approaching black – and there is no persistent foam.

I don't know what this stuff is but I'm guessing it's polytannins.

5) I add 4 mls of chloroform to the (hydrolysed and neutralised) solution. Seal the (polypropylene) tube and shake vigorously for 30 seconds. I then spin the tube in a table top centrifuge at 4000RPM for 5 minutes. Next I use a graduated plastic (polypropylene) transfer pipette to penetrate the aqueous layer, bubbled out the tip (which has a “plug” of “polytannins” and some aqueous) and then collect the chloroform from the bottom, while avoiding the salt pellet. I transfer this into a glass (scintillation) vial. Then I add another 4mls of chloroform and extract again. I end up with more than 8mls in the vial because some of the ethanol moves into the chloroform. (They're miscible.)

Attached is a pdf with some photos and experimental flow chart.

Thanking you in advance for your suggestions.

Best regards,

Jamie