The Half Decent Pharmaceutical Chemistry Blog

Last night I couldn't sleep, so, I began to think about a remedy for insomnia, in case this problem becomes a recurrent nuisance. If you were thinking that, being a vicious pharmaceutical chemist, benzodiazepines would be my first choice, though, you'd be wrong.

Chamomile tea is well-known, all over the World, as a reliable antiinflammatory for the digestive tract: flowers of Matricaria Recutita, in particular, are full of substances, such as bisabolol and essential oils, with remarkable efficacy in the treatment of severe conditions (even irritable bowel disease).

Weirdly, here in Italy, this plant is massively (and almost only) used as sleep aid, while few people, believe or not, know about its role as spasmolytic for the intestine, like the rest of the World seems to ignore the importance of chamomile tea when it comes to calming your nerves.

So, in case you were wondering which technique I reckon to be the best, here's my hint: instead of doing like anyone else (having a cup of chamomile tea before going to bed), you, chemists, are supposed to extract either the terpenes and the flavonoids from the capitula of M. Recutita you'll certainly find in your kitchen: such a time-consuming procedure, it'll certainly make you fall asleep. If you can't find it, then, go out and get some: hey, I know it's 1:30 am but you're suffering from insomnia, aren't you?!

That's pretty much what we did in our lab course: the extraction of both types of active substances from the flower-heads of M. Recutita, followed by a mere qualitative analysis. No yields, today!

Approximately 1 g of dried capitula of Matricaria Recutita (German Chamomile) were crushed in a mortar. First, we set to extract the lipophilic molecules (terpenes), so,  we extracted them using 20 mL of dichloromethane for each of the two separations.

However, this being a solid-liquid extraction, we didn't need a separatory funnel: what we did, instead, was to stir each time, for 20 minutes, the content of the 100 mL conical flask we were using. Dichloromethane became yellow and, in the end, was entirely collected in a round-bottom flask.

This solution was concentrated, using a rotavap, to 1 mL of a greenish oil. Meanwhile, the mobile phase for the TLC was prepared mixing 9.3 mL of toluene with 0.7 mL of ethyl acetate (93:7).

Once our oil was ready, we quickly loaded it on our silica layer, in order to prevent the terpenes from reacting with air (which would have turned the colour of the oil from green-yellow to blueish).

The layer was therefore spayed with a 5% sulphuric acid (ethanolic) solution (the acid catalyzes the following reaction) and, then, with a 1% vanillin ethanolic solution (predictably the carbonyls react with the carbocations yielded by the acid). We placed the layer in the stove at 100°C and exposed it to UV light ten minutes later.

 

 

I have to say I didn't do any of these operations, since my colleague began this experiment while I was busy getting some IR spectra of products we had yielded on previous days. Still, once she had finished the extractions, she handed me the conical flask with the capitula and I kicked off the second part of the experiment, while she went on with the terpenes.

I put all the capitula in a 100 mL round-bottom flask, added 30 mL of methanol and heated under reflux (80°C) for 15 minutes.

With some cotton in a funnel I transferred all the organic phase in a test tube: 2 mL of this methanolic solution were concentrated at a rotavap, yielding a first sample that should be rich in rutin.

The rest of the test tube was concentrated to 1 mL, as well. 5 mL of water and 1 mL of concentrated HCl were added in the hope to get quercitin only (simply breaking the glycosidic bond of rutin, using a strong acid).

I heated under reflux these reagents for 20 minutes (at 100°C), to increase the yield and added 10 mL of water.

I poured this solution in a test tube, where I also added 2 mL of ethyl acetate: amazingly, this liquid-liquid extraction is accomplished  by simply shaking the tube, allowing the solvents to separate and, at last, decanting the organic phase. This procedure was done twice and the solution of ethyl acetate was dried, in the end, with sodium sulphate.

For the last TLC, given that our analytes are much more hydrophilic, a delicate emulsion had to be quickly prepared: ethyl acetate/acetic acid/water (100/22/27) were, in fact, chosen as the most suitable mobile phase for the job in hand. Terribly reactive, that's for sure.



Finally, I sprayed a 1% natural probe (which contains chlorogenic acid) ethanolic solution over the layer and some PEG (4000), so that yellow stains could be easily detected using fluorescence.

Good night!

My readers have already got used to my typos in many of the reaction schemes I post: that's because I usually prepare the pictures the night before the day I'm likely to write the related post. So, predictably, while the latter are generally correct, the pathways often prove how your concentration levels decrease once midnight has passed.

Still, at least I don't print and hand them to students of any lab course. And if I did it, I'd certainly check them more than once before hitting the "print" button. Apparently, that didn't happen when the procedures for our lab were prepared.

On day 3 of Laboratoire Organique, we set to perform the chiral resolution of one of the most basic compound I've ever handled: alpha-Phenylethanamine. This molecule is, in fact, so reactive that it quickly (really quickly) reacts with the humidity and carbon dioxide in the lab, yielding a white carbonate on the inner surface of the piece of glassware where you placed it.

Our aim was to separate (R)-(+)-alpha-Phenylethanamine by fractional crystallization, using her Majesty (+)-Tartaric Acid. That, at least, was written on the notes we had been given and our professors outlined.

Immediately, though, things looked somehow weird: the picture on our papers actually belonged to (S)-(-)-methylbenylamine, although just below you could read (R)-(+).

Then, as people began to reach the final step of the experiment (namely, the determination of the specific rotation of the pure enantiomer, compared to what reported in literature: +40.1°, which is correct, since it's supposed to be the (+)-enenatiomer) they were rather surprised to get a negative optical rotation!

In the end, when we all had finished and all had got a negative result, the professors looked equally taken aback and admitted they had no idea how this could have happened.

But, of course, I refused to give in and I'm happy to report that I've detected the problem. Here.

You see, the instructions we followed are those to purify the (S)-(-) enantiomer, which, as I'm going to show you, yields lovely prismatic crystals when mixed with (+)-tartaric acid. The (R)-(+) one, on the contrary, remains in solution, although it can be (easily) separated as well. So the picture and the text of the recipe were correct: unfortunately, someone misinterpreted the structure and named it incorrectly...

2.75 mL of a racemic mixture of phenylethanamine were accurately put in a small beaker with a graduated pipette. 14 mL of methanol were therefore added.

To make sure that this really was a racemic mixture, we checked the optical rotation of this solution. Terrifyingly, because you'll then have to go on working with this quantity of phenylethanamine, you can't lose anything, not even spilling a drop. So, everything must be done carefully, slowly and with a Pasteur pipette.

While I was doing this delicate and crucial operation, my colleague weighed 6.20 g of (+)-tartaric acid, dissolved them in a 250 mL Erlenmeyer flask, using 85 mL of methanol. Then, to speed things up a little bit, we heated this solution under reflux, with the magnetic stirrer on: in fact, tartaric acid is much more soluble in hot methanol than at room temperature.

Once the solution looked clear and wasn't hot any more, we added the first amount of methylbenzylamine (drop by drop) and another 3.15 mL of pure racemic mixture. We had to wait until this stage because, if we had begun with all the 5.9 mL of phenylethanamine, an enormous volume of methanol would have been required to dissolve it.

We closed the conical flask, stored it in a safe place and went home. As it turned out, it took three days for the crystallization to occur: finally, the (-)-amine tartrate salt had completely precipitated, in the form of white, tiny, prismatic crystals.

Then, we filtered, using a Büchner funnel, and washed them with a very small volume of methanol.

To yield the purified amine, we dissolved in water (approximately four times the weight of our product) and added exactly 2.97 mL of a 14 M NaOH solution, we prepared at the moment (5.6 g in 10 mL of water).

You use a little more NaOH than that actually needed (looking at the stoichiometry of the reaction), just to be sure you get back as much amine as possible.

This basic solution was poured in a separatory funnel, where four extractions with ethyl ether (15 mL each time) were carried out.

We washed the organic solution with Brine and got rid of water adding sodium sulphate (which we washed with ether, in the end).

We put a bit of cotton in a funnel and poured the solution in a 50 mL round-bottom flask, whose weight had already been calculated (64.2565 g) and evaporated all the ether utilizing a rotavap: we ended up with an oil with a constant weight.
The weight of the flask (65.4904 g) meant we had 1.234 g of (S)-(-)-phenylethanamine.

With a pipette, we transferred all the oil into a 10 mL volumetric flask, which we filled with methanol: from the optical rotation ( -3.85°), we derived the specific rotation of the product ( -31.20°).

Now, theoretically, the specific rotation should be - 40.1°, but the most important thing was to get anything below - 30° and, moreover, when we analyzed all the results we had got, we realised ours was in line with the mean specific rotation.

 

For what concerns the person who got confused with the chirality, I think we should blamed tartaric acid for this inconvenient: as you mention its name, you can't avoid having a glass (or two) of red wine, can you?

 

Three years ago I spent a week in Dublin. In those days, I still used to smoke (although I was going to start my third year at the university), like most of nerd teenagers who want to look older, cooler and tougher and, so, begin to smoke cigarettes in the bathrooms of their high school.

I remember that, due to terrifying price of cigarettes in Ireland, on that very trip I began to ask myself how hard giving up smoking could have been: as it turned out, it was terribly easy.

Nevertheless, even though I was considering quitting, I was particularly attracted by a posh cigar shop in the center of the city. So attracted, that you can realise how much I liked that place. Especially its witty name...

 

 

I'd like to stress I didn't take the picture displayed above: I stole it from another website, downloaded it on my hard disk and edited it, since the original was so bad, that made me feel sick.

That said, let's go back to shop: you see, although I gave up smocking two years ago and I've never suffered from (real, vicious) craving,  I'll always be ready to celebrate any (massive, outstanding, insert-a-big-adjective) achievement (my graduation, my admission at a prestigious university for my PhD) with a sophisticated and outrageously expensive, Caribbean cigar.

During my lab course, though, instead of a decent cigar emporium, those who were in charge to buy the cigars should have looked for a much cheaper place: still, it's not that stupid to use the cheapest cigars that money could buy, for extracting nicotine and determining its concentration.




Mind you, at least we became, for an afternoon, the one and only Half Decent Cigar Emporium in the entire World!

Each of us was given approximately 2 g of a rubbish, though pretty well-known, Italian cigar. We placed it in the mortar and set to smash it into a fine powder, which we weighed (2.0053 g).

The brown powder was put in a small beaker and 50 mL of a 1% sodium hydroxide solution were added to start separating nicotine from some of the components of the plant (namely, tannins). This mixture was stirred for 15 minutes at room temperature.

Then, we filtered the suspension using a Büchner funnel: given that we were to pour a mixture full of components of the cell wall, we carefully covered the filter paper with a couple of tea spoons of Celite.

We poured the liquid in a separatory funnel and begin the extractions: we added 20 mL of dichloromethane thrice.

Sadly, however, the two phases produced a nasty emulsion, which obviously had to be broken. Now, this was no surprise, if one thinks about the solubility of dichloromethane in water, but, on the other hand, this should be quickly achieved just waiting and, then, opening the bottom valve and emptying the sep funnel very slowly.

In fact, as the emulsion reaches the tight end of the funnel, this should automatically break the system, separating the two phases completely. Unfortunately, this helped but wasn't enough.

So, every time the organic phase (hopefully full of nicotine) was collected in a beaker, the emulsion was placed in two test tubes we centrifuged for a short while.

Finally, the dichloromethane solution was dried with sodium sulphate and transferred to a 100 mL volumetric flask, whose final volume was reached with some extra dichloromethane.

1 mL of this last solution was put in a round-bottom flask to concentrate nicotine, evaporating the solvent with a rotavap. Once the solvent had gone, we added 10 mL of methanol and measured the absorbance of our sample at 262 nm.

Our reference was a 10 µg/mL nicotine standard solution, whose absorbance, at the wavelength we are using, was 0.4670.
Solving a simple proportion, we calculated the concentration (3.717 µg/mL) from the value we read (0.1736).

This means, the concentration of nicotine was 0.188%, which is consistent with what was expected (0.15% - 0.3%).

Meanwhile, we had prepared the mobile phase for our TLC: this solution consisted of a 60/10/1 mixture of dichloromethane/methanol/ammonia.

On a silica layer, a standard of nicotine was lined up with a solution yielded concentrating 20 mL of the 100 mL solution to approximately 1 mL.



This showed nicotine wasn't the only alkaloid we had extracted: more lipophilic metabolites (I guess, in descending order, nornicotine, cotinine, norcotinine and nicotine N-oxide) were also detected.

While waiting for the layer to be exposed to the UV-light, what had been left over of the 100 mL flask was placed in a round-bottom flask and the solvent evaporated. Then, we added 3 mL of methanol and 5 mL of a saturated picric acid solution in methanol: this test is the standard one for the tertiary amines which react with picric acid and yield yellow salts.



This was done to complete the analysis, following the procedure described in the European Pharmacopoeia.

Yesterday, I gave you an example of how biology can make the liefe of organic chemists easier. Today, I'm going to show you how organic chemistry can make biology less boring.

You see, I receive hundreds of emails at my new address (please, check it out) from people who are annoyed by the molecules you deal with when studying biochemistry. Take amino acids, for example: they all look pretty much exactly the same and, even if you play with pH, after a short time, you'll begin to yawn and ask yourself why, in the name of God, you have to go through this monotony.

Even proline, which, at first glance, may stand out as the most different amino acid, is a little bit boring.

But don't worry, ladies and gentlemen, because I've worked out a way to live things up a bit! Welcome everybody to Pimp My Amino Acid!

In the lab...ehm, at West Coast Custom, we have reckoned proline would be much better with a brand new protecting group bound to that nitrogen, instead of that rusty hydrogen. We chose Boc because it looks cool: the product will be big, noisy and heavy...like a proper, American muscle car. Oh, and it'll be stable at 25°C, too.

Still, the protection mustn't change the stereochemistry of natural L-proline, so that it will be able to race against the normal, natural ones, but, being much better-looking, ladies will love it!

I put 0.6048 g of L-Proline in a 100 mL round-bottom flask and dissolved the amino acid in 10 mL of a 5M NaOH solution, which yielded its sodium salt. This first reaction is, thus, a sort of activation of the amine.

Before I could add Boc, 4 mL of tert-Butanol had to be added to the aqueous solution, so that the protecting agent was dissolved and ready to react with the salt of the amino acid. Due to its low melting point, Boc was quickly weighed (1.2950 g) and added little by little, with the magnetic stirrer on.



We checked the pH of the mixture: the adduct we were trying to yield is perfectly stable when the solution is basic, but one of the features which has made Boc so popular is that the deprotection of the amine requires rather mild acidic conditions.

Then, theoretically, you should keep on stirring for 24 hours, but because we knew we would have finished the reaction 4 days after (it was Friday and on Tuesday it would have been the Labour Day), we switched off the stirrer. Someone would have turned them on at least 6 hours before our arrival on Wednesday, we were told.

So, after a worringly long time, it was time to go on pimping our proline! Actually, the magic had already happened, but, before we could unveil our masterpiece to the nerd guy who needed our help, a series of separations had to be performed.

First, we extracted with 25 mL of n-hexane twice: this gave us the oppurtunity to get rid of most of the Boc which hadn't reacted.



We collected the organic phases and extracted all the product which might have eventually ended up there, because protonated, with a saturated sodium bicarbonate solution (20 mL, twice): using a salt, you turn the acid into the its salt, as at the very beginning.

Then, with all the aqueous solution, it was time to decrease the pH to 2-4, using an agent (citric acid) too weak to deprotect the amine, but capable of protonating the carboxyl group.
25 mL of ethyl acetate were required for each of the four extractions we did, once the product was an acid again.

But that wasn't the end: there was still the proline which hadn't reacted to get rid of. To achieve this goal, another separation had to be done, this time, adding 25 mL of water. Twice.

Finally, our ethyl acetate solution was ready: it was dryed with sodium sulphate, filtered and the solvent evaporated with a rotavap.

Theoretically, you should get a white powder from this operation, but we had already been warned that the odds were we would have got an dense oil, instead.

Unsurprisingly, we had to crystallize the oil which resulted, placing the flask on ice, adding approximately 5 mL of n-hexane and scratching the inside of the round-bottom flask wtih a glass rod.

After 15 minutes, the crystals had appeared and we filtered them with a Büchner funnel.

The yield for this reaction, bearing in mind the huge number of extraction, was supposed to be nearby 60%. Since the weight of our product was 0.5891 g, our yiled was a mere 52.84%.
Still, the melting point perfectly matched that found in literature (135°C).



However, the final and most important test was the specific rotation. The value we had as a reference was from a 2g/mL solution in acetic acid (- 60.6°). Our sample was a 0.0504g/mL solution (concetrated acetic acid as solvent), instead, but, still, the result was a lovely -57.5° as specific rotation (optical rotation: -2.9).

Now you can thank us from pimping your Proline!

Nowadays, organic chemists normally cross the boundaries of traditional synthesis and play with stuff biologists thought to be their own, personal territory. This phenomenon has had a terrific impact: for instance, the conidia Curvularia Lunata plays a key role in the industrial preparation of cortisol acetate. It has made easier to yield the drug, skipping many steps, so, as a result, cortisol acetate has become cheaper, as its synthesis has been shortened.

During our half-decent lab course we, too, had a go at mixing synthesis and enzymes. Our aim was to prove how easily you can perform an enantioselective reduction of a beta-keto-ester (methyl acetoacetate) using normal brewer's yeast.
The most important advantage of the "biological" way lies in its simplicity: to yield this very product (with an asymmetric synthesis) would be incredibly more complicated, whereas the use of a yeast dramatically reduces the problem.

We weren't interested in the final yield very much: it was the specific rotation what we wanted to determine, above all.

In a 250 mL conical flask, with a ground-glass neck, we prepared the following solution: 64 g of sucrose, 0.4 of Na₂HPO₄ and 100 mL of water.
Because we will introduce a living being in this piece of glassware, we must create an environment where our cells could live and enzymes normally work. So, we set the temperature to approximately 35°C.

The flask was placed on the stirrer and we waited until all sucrose was dissolved. Then, we added 14 g of commercial brewer's yeast: I mean, it came from the supermarket just in front of the department, not from, say, a Chinese supplier.

We stirred the mixture again, but this time, following the instructions we had been given, we stirred rather vigorously for at least 15 minutes.  Actually, it took a little bit more than that to get a decent, homogeneous suspension.

Finally, we added the last, missing ingredient, the reagent, as 2 mL of methyl acetoacetate were poured in the flask.

Before leaving the lab, we assembled the reaction apparatus: the conical flask and a reflux valve (thanks to their ground glass junctions)  were joined together. At the other end of the valve, we put a rather short silicon tube, which provided the link to a Pasteur pipette.
This was partly immersed in a test tube full of glycerin. Obviously, we fixed this part of the apparatus so that it was in oblique position.

This system wasn't for show: it was a brilliant solution to exhaust the carbon dioxide produced by the yeast. Interestingly, when we arrived in the lab, on the next day, we could appreciate everything had worked from the small bubbles in the test tube.

So, once the mixture had been stirred overnight, at room temperature, we filtered the solution with the help of Celite, which is useful whenever cells are involved in such procedure, unless you want to have your filter rapidly blocked and useless (or so obstructed that the filtration becomes an endless agony).

We saturated the resulting solution with NaCl and, then placed it in a separatory funnel, where we extracted it thrice, with 30 mL of dichloromethane each time.
The organic phases were collected and traces of water removed with anhydrous sodium sulphate.

In this solution, however, there were still either methyl acetoacetate and our product. To check if that was really how things were, we ran a TLC (not shown) where the mobile phase consisted of a solution dichloromethane/methanol (98:2). The products were detected only once the layer had been treated with KMnO₄ and NaOH (yellow stains).

At this stage, the two molecules had to be separated chromatographically in a column packed with silica gel. Now, unfortunately, none of us actually prepared anything: the columns were prepared either by our professor or by PhD students.

Still, at least I can describe what they did: half of the volume of the column was filled with silica gel, which was then placed in a beaker. There we added 100 mL of the aforementioned mobile phase; meanwhile, a little bit of cotton was put right over the bottom valve of the column, together with 5 mL of mobile phase. Then, the suspension of silica gel was poured in, some mobile phase added, the bottom valve was opened, for a short amount of time, to let the stationary phase settle.

With a pipette, they loaded our sample (letting it running down the inner surface and poured some more MP). Before that, a bed of sand was laid in order to get an accurate loading and, as a result, an excellent separation, since sand provides a higher density, so your analytes gather right at the beginning of the stationary phase.

We discarded the first 10 mL and I began to (manually) collect everything coming out of the column in test tubes, while making sure the stationary phase never got dry.

While collecting the samples, I performed several TLCs (same conditions of the previous, preparatory one) to determine the content of any test tube that had already been filled, using a standard solution of methyl acetoacetate as reference.

Predictably, the first analyte to be eluted was the reagent (more hydrophobic than the product), while, at the sixth TLC, one stain confirmed all the product was in one test tube only. Nevertheless, we put the content of those collected immediately before and after, too, in a round-bottom flask.

 

Once we reached a constant weight (0.1118 g), the oil was placed in a 10 mL volumetric flask and chloroform was added to reach the final volume.

Finally, we measured the optical rotation (+ 0.40°): so, we really had yielded the S-(+)-methly-3-hydroxybutanoate we were looking for and its specific rotation (+ 35.78°) was pretty close to the one found in literature (+ 38°).