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Anthocyanins from Red Cabbage -
With Experiments

by C. Thorsten


Introduction:

A perennial favorite of chemistry and biology teachers is to have students extract the reddish-purple coloring matter from red cabbage, typically with the purpose of showing off the colored substance's use as a pH indicator.  The  experiment has all the characteristics of a good classroom demonstration:  a comparatively high degree of safety (depending on how it's done);  ready availability of materials;  visual interest;  and relative simplicity.  
Red cabbage is Brassica oleracea var. capitata f. rubra.  A red cabbage could certainly be called Brassica oleracea, but then so could broccoli, kale, Brussels sprouts, etc.  These vegetables are all the same species, just different varieties and cultivars.  It is the red or burgundy-colored cabbage that has the compounds we seek.  The study of these compounds can range from as elementary to as involved as one desires.  Last this writer checked, there were still papers appearing in the scientific journals about extraction and characterization of the pigments in Brassica oleracea capitata rubra, and there probably will be for some time.  Why is that?  Wouldn't red cabbage be exhaustively studied, even trite as a subject for a research project by now?  Not at all.  Let's examine why.  We'll have a look at the fascinating properties of the anthocyanins, the active coloring agents of the common red cabbage.
Early investigators, around the beginning of the19th Century, knew the coloring matter in red cabbage and certain other plants would turn different colors depending on the acidity or alkalinity of the solution. They knew how to extract the material with water, alcohol, and other solvents. By the early 20th Century they even had a rough idea of the pigments' molecular structure (Onslow, 1925).
Thanks to decades of experimental foundation, we know today that red cabbage contains several anthocyanins having different substituents and functional groups.  It has proven no small task to sort them out.  Cabbage anthocyanins are mostly in the form of glycosides;  in other words, they have sugar molecules chemically attached to them (Timberlake and Bridle, 1973;  USDA, 2003).  Some have acyl functional groups instead of or in combination with these sugar molecules;  the acyl groups can contribute greatly to color stability (Piccaglia, Marotti, and Baldoni, 2003). 

Inset:



Of Acyl Groups and Esters

The simplest acyl group is the "acetyl" group.  This would be CH3COOH without the OH, meaning a CH3CO group.  We sometimes write "CH3CO" simply as "Ac".  Depending on the source, "Ac" can refer to acyl groups other than the acetyl group.  However, to keep everything straight we might decide to write "Fer" if we're referring to ferulic acid, "Sin" if it's sinapic acid, and so forth.

If the acyl group were attached at an -OH group, it would form an ester linkage.  An acetyl group would thus give an acetate.   The simplest acetyl ester is methyl acetate.  We could write this as  CH3CO-O-CH3 or simply Ac-O-Me.  ("Me" for "methyl").

Now suppose we wanted to represent a sugar which had been acetylated at one of its OH groups.  If it were the OH at carbon 6, we would call this 6-O-acetyl glucose.  Sinapic acid esterified with the -OH of carbon 6 would instead give us 6-O-sinapoyl glucose. We might write shorthand "formulas" for these using strings of name-abbreviations.  To a biochemist, the abbreviation "Gluc" is actually much more informative than C6H12O6.

As a general rule, an acylated sugar or polyphenol will have the O-linkage (in other words, Ac-O-R, not Ac-R;  however, acylation does commonly occur at N-linkages, as in N-acetylglucosamine).

C-acylation is rare in sugars, polyphenols, and other molecules having hydroxyl groups, since -OH groups are attacked in preference to carbons.  C-acylation is apparently much rarer in biological systems than it is in the chem lab.
  


The anthocyanins can be difficult to separate from each other, as well as from other substances such as sugars. In the case of "red cabbage indicator", simple bench-top means allow for separation of the anthocyanins from interfering  substances, but not so much from one another.  Below the university upperclass level, we'll assume the methods don't include HPLC machinery or even affinity chromatography columns having specialized resins.
The compounds we're after from red cabbage are essentially cyanidin glucosides and molecules derived from them.  The USDA (2005) lists cyanidin-3-glucoside,  cyanidin-3,5-diglucoside, and several others (mainly cyanidin sophoroside-glucosides) 1.  Incidentally, the name "cyanidin" has nothing at all to do with cyanide;  there are no CN groups anywhere on the cyanidin molecule.
Figure 1 shows the structure of the cyanidin aglycone in its oxonium configuration; "aglycone" meaning the core molecule without any attached sugars, and "oxonium" because the oxygen in the central aromatic ring has taken on a positive charge. Sometimes this charged anthocyanin molecule is alternatively called the flavylium cation, deriving from its flavonoid structure.  The molecule in general is also a polyphenol, so named because it has OH groups attached to 6-membered aromatic rings. 
Anthocyanins assume the oxonium or flavylium cation configuration at low pH (being predominant at less than about 3 or 4), which is a good general representation considering that traditional extractions utilize hydrochloric or acetic acid, most often dissolved in alcohol (see for example Timberlake and Bridle, 1973; Piccaglia, et al. , 2003). The O+ would have a Cl- (not shown) associated with it in the case of cyanidin chloride.  This maintains electrical neutrality.

The cyanidin molecule

Figure 1: The cyanidin aglycone at acidic pH.  Starting at the O+ in the middle ring and going clockwise, the second carbon atom is numbered 3. It this carbon whose oxygen forms the 3-glycoside linkage; hence the name. Sometimes the oxygen attached to carbon 5 also forms a glycosidic bond.

Cyanidin-3,5-diglucoside
Figure 2: Cyanidin-3,5-diglucoside, sometimes called cyanin. The 3-monoglucoside is sometimes called asterin or chyrsanthemin  (Merck Index, 10th ed., entry 2682; see also Onslow, 1925). The sugars themselves do not have to be glucose.  Sophorose  (2-O-beta-D- Glucopyranosyl-alpha-D-Glucose: Merck Index, 10th ed., entry 8566) is another one encountered. The sugars may or may not be acylated with organic acid groups including malonic, coumaric, sinapic, and / or ferulic (USDA, 2003). See diagrams below for representations of these organic acids.

We drew these from scratch w/ image editing software.  Copyright CR Scientific LLC p-Coumaric Acid  
Ferulic acid Sinapic acid

Figure 3.  The above diagrams show the close structural similarities between at least three of the acid moieities found in acylated anthocyanins from red cabbage (in other words, the acyl groups are not random). The similarities are interesting but not surprising, since different biological compounds often have synthesis pathways that overlap on certain enzymes.  Furthermore, biological systems are not random throws of the dice;  some definite range of structure-function is expected.
We should be able to break the glycoside 2 linkages by treatment with hydrochloric acid.  After all, a glycosidic bond is really just a type of acetal, which can be converted back to a hemiacetal by treatment with aqueous acid; hemiacetal formation means the glycosidic bond must break.  According to Onslow (1925), the glycosidic linkages of anthocyanins will hydrolyze with ease in 20% HCl. Supposing we're starting with the molecule in Figure 2 (above), hydrolysis should leave us with cyanidin chloride and some free-floating sugar molecules, if all goes according to plan. While HCl can also attack certain types of acyl groups associated with anthocyanins (depending on pH, time, and temperature), the results will suggest this isn't of great consequence for the indicator properties of red cabbage-- which seem to rely only on the flavonoid nucleus, not on any attached groups.

General Procedures:
Avoid opening the concentrated HCl outside the fume hood. While concentrated HCl will emit visible fumes in moist air, HCl vapor will be invisible otherwise.  It is still there and can still corrode your lungs.
Though we are starting with a food substance (cabbage), treat all liquids as if they're toxic - even the red cabbage extract. Remember we're going to be using methanol, acids, etc. It's too easy to lose track of which extracts are poisonous and which are not, so assume they all are.
 
1. Extraction

1A. Extraction with Water
This method is extremely simple and requires a minimum of supplies. It is also the safest method with regard to classroom demonstrations.

1A-1. Chop some fresh red cabbage leaves into tiny pieces. Place them in a beaker and cover them with distilled water, amounting to about 1 1/2 times their volume.

1A-2. Slowly heat the beaker to 50 C and maintain for 10-15 minutes. It is not necessary to boil the liquid. Boiling is acceptable as long as you're planning only to use the material qualitatively as a pH indicator. 3

1A-3. When the colored liquid is cool, strain it through filter paper and save it. Discard the chopped cabbage and the filter paper.  If desired, centrifuge the liquid.  For later experiments involving crystallization, try to obtain a liquid with as little particulate matter as possible.

1A-4. (Optional:) Using a separatory funnel, shake the extract together with diethyl ether (make sure you hold the stopper on!). Allow the layers to separate and save the aqueous layer. According to Onslow (1925) the ether layer will take up some of the non-colored compounds from the extract.

1B. Extraction with Acetone
This extraction method does not show up very often in the literature due to the prevalent use of acid-alcohol, but it does work (see for example Durkee and Jones, 1968); one benefit is that the anthocyanins will dissolve in acetone, leaving behind salts, enzymes, and other unwanted components-- some of which would degrade the anthocyanins over time. Durkee and Jones (1968) used acetone chilled to -20 C to do a preliminary extraction, then created a powder from the once-extracted cabbage.  They then extracted this powder at least once more with cold acetone, combining the liquids and drying with concentrated sulfuric acid. Update:  A reader pointed out that some of the acetone will probably dissolve in the concentrated sulfuric acid, possibly reacting to yield mesitylene.  We are not sure offhand how much this would happen at -20 C;  it is, however, worth noting that the original source (1968) does call for drying the cold acetone with sulfuric acid.  Its authors may have overlooked that point, but presumably they were still able to continue with their task.
We can use a grossly-simplified version of the Durkee and Jones procedure, eliminating the drying step anyway;  losing some of the anthocyanins is alright for our purposes:

1B-1. Chop some fresh red cabbage leaves into tiny pieces. Place them in a beaker and cover them with acetone, amounting to about 1 1/2 times their volume.

1B-2. Cover the beaker and put in a cool, dark place. Let stand for at least 24 hours. Stir periodically if desired.

1B-3. Filter as in 1A-3. If desired, centrifuge the liquid. Acetone is miscible with water, so at this stage the cabbage extract could be used as an indicator.

1B-4. (Optional:) Using a separatory funnel, attempt extraction with light petroleum ether or chloroform. According to Hedin, et al. (1967) and Durkee and Jones (1968), this will remove some of the colorless polyphenols and a few other compounds from the extract (it is odd, though, that Durkee and Jones should suggest an acetone-chloroform extraction, since acetone is miscible with chloroform...)

1C. Extraction with Acid-Alcohol
Most experimental procedures surveyed (dating c.1914 to the present) call for a 1% v/v solution of HCl in methanol (less frequently, ethanol is used). Sometimes acetic acid is used in place of the HCl.

1C-1. Add enough concentrated HCl to methanol to make an approx. 1% v/v solution (It is sufficient for this application to mix 99 parts methanol with 1 part conc. HCl by volume;  where molarity and weight percent of a given lot of HCl are required, it's necessary to look these up in a chemistry handbook). 

1C-2. Chop some fresh red cabbage leaves into tiny pieces. Place them in a beaker and cover them with the HCl-methanol solution, amounting to about 1 1/2 times their volume.

1C-3. Cover the beaker and put in a cool, dark place.  Let stand for at least 24 hours.  Stir periodically if desired.

1C-4. Filter as in 1A-3. Save the colored liquid.  If desired, centrifuge it.

1C-5 (Optional) Extract with chloroform or light petroleum ether.  Save the alcohol layer.

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Cabbage extract in water with no added acid or alkali (tube on left); cabbage extract made to  approx 6 molar in HCl (tube on right).

The HCl solution was boiled for 5 minutes with no apparent loss of color.


Click for larger image.


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Went a little too far with the NaOH titration (see II: C-5), bringing the indicator past its naturally purple color.  Note the greenish, almost teal hue of the solution now.

Click for larger image.
 .

Titrated back to the "natural" purple color, though the pigment is somewhat diluted now.  When this evaporates it's going to have abundant NaCl crystals from the HCl-NaOH reaction.

Click for larger image.





2. Experiments with the Extract

2A. Crystal Growth
Onslow (1925) mentions the early efforts to crystallize anthocyanin pigments. They all seem deceptively easy. Red cabbage anthocyanins are supposed to crystallize from alcohol as red-brown needles (Onslow, 1925;  Merck Index, 10th ed., entry 2682).
There are several possible obstacles to growing these crystals: attached sugar molecules and acyl groups; other unwanted substances from the cabbage; salt and other compounds introduced during extraction; wrong solvent or combination of solvents; and possibly too-rapid evaporation.
Many of our own attempts failed despite trying a variety of conditions. Water extract did not yield crystals, even after hydrolysis of glycosidic bonds with HCl. Titrating back to neutrality with alkali and trying to grow cyanidin crystals did not work, either (see photos below for the effect that NaCl crystals had on the red solution). Drying and re-dissolving the amorphous reddish-purple material in alcohol was also unsuccessful. Even starting with an alcohol or acid-alcohol extract did not produce crystals, whether the evaporation was slow or fast.
Starting with an acid-alcohol extract and boiling some of this in an equal volume of concentrated HCl was a step in the right direction.  The resulting residue after evaporation had red-brown spots at least, though these weren't regular enough in shape to be called crystals in any sense this writer had ever encountered.   Trying again but controlling the rate of evaporation actually did yield something resembling red-brown needles (see photo below).


 
Photos taken with Mini-VID USB and Observer III microscope at 40x

Interesting development: The salt crystals (after neutralizing w/ NaOH) attracted and concentrated the dissolved, red pigment.  The affinity of the red pigment for NaCl, an ionic solid, suggests that it's in the oxonium chloride form.  The purple clumps, in contrast, seem to have no special affinity for the salt and are probably in the uncharged state (most likely, the quinoid form).



Partial success:  the photo at left shows the first instance of  "red-brown needles" that we were able to obtain-- only via slow evaporation of the HCl-hydrolyzed alcohol extract.  The "needles" actually appear to be elongated aggregates of smaller crystals. They do not dissolve appreciably in water, which is a good sign. 




Success (?):  the photo at left shows some crystals grown from extract after it has undergone:  (1) acid hydrolysis, (2) adsorption onto filter paper, (3) re-uptake with acetone, and (4) subsequent slow evaporation.  Note the curvature this time.

Anthocyanin crystals are tough to grow!





2B. Heat and Chemical Stability
There is no doubt that heat will eventually destroy or degrade some of the phytochemicals in red cabbage. Certain additives will of course accelerate this degradation. 
Here we'll focus on the colored pigments and the qualitative results of heating in the presence of acid, alkali, and salt.  Note that these procedures are not quantitative with regard to temperature; "boiling" may correspond to a different temperature in each, depending on what's dissolved in the liquid. (Step B-3, for example, could be expanded to try different salt concentrations, resulting in different boiling point elevations. How much heat does it take to destroy the anthocyanin color?)

2B-1. Put 3 mL of the red cabbage extract into each of five different 10 mL micro flasks-- or five, large (18 mm or bigger) test tubes. 4  Set the first one aside -- this is the control sample.

2B-2. To one of these flasks add 3 mL of concentrated HCl. Add 2-3 small boiling chips. Boil this flask IN A FUME HOOD for about 5 minutes. Do NOT heat it to dryness. If necessary, add distilled water to keep the total volume to at least 3 mL. After boiling, set aside and let cool.

2B-3. To another one of the micro flasks add 3 mL of saturated salt (NaCl) solution. Add 2-3 small boiling chips. Boil the liquid down until it thickens, but do NOT heat to dryness or anywhere near. If necessary, add distilled water so the total volume is never less than 2-3 mL. Continue this for about 5 minutes.  Check the temperature periodically and write it down. After boiling, set aside and let cool.

2B-4. To the fourth flask add about 3 mL of distilled water and a couple of small boiling chips. Boil for about 5 minutes. Do not heat to dryness-- if necessary, add distilled water to keep total volume at no less than about 3 mL.

2B-5. To the fifth flask add about 3 mL of ca.12 molar NaOH solution (strong but not quite saturated) and a couple of small boiling chips. Boil for about 5 minutes. Do not heat to dryness-- if necessary, add distilled water to keep total volume at no less than about 3 mL.

2B-6. When the flasks are cool, make sure the relative volumes are the same in each one. Add distilled water where necessary to make the volumes approximately equal. Compare the colors and their intensities against a white sheet of paper.  Which ones, if any, lost their color?
Save these solutions for the following procedures.

2B-7.  If desired, place one drop each of red cabbage extract in the wells of a spot plate.  Number them 1 through 7.   To the corresponding numbered wells, add a drop of ONE of the following solutions (warning: lead and hexavalent chromium are toxic!): 
1.) lead (II) ions
2.) copper (II) ions
3.) iron (II) ions
4.) iron (III) ions
5.) Cr(VI) (as chromate or dichromate)
6.) cerium (IV) ions
7.) hydrogen peroxide

Which, if any, give colored precipitates or complexes?  Which destroy the anthocyanin color altogether?  Why?  Are there any other transition metal ions which might have interesting effects on the anthocyanin molecule? 


2C. pH Experiments

2C-1. Using a clean spot plate or several test tubes, put enough red cabbage extract into each well or tube that you'll be able to see any color changes that take place. If you wish to do any future calculations, keep careful track of how many drops or milliliters you use. Keep it consistent.

2C-2. The first well or tube will be a control; do not add anything to it.

2C-3.   Since we are working with small amounts of extract, the plan is to add only one drop of acid or base to each. 
Use a dropper that you've calibrated so you know exactly how many drops make 1.0 mL.  Divide this by number of drops to get volume of each drop in mL.  It will probably be somewhere between 1/15 to 1/20 of a milliliter.  (It also depends on the viscosity and surface tension of the fluid being dispensed.)
In some clean test tubes or flasks, prepare a series of dilutions starting with 6 M HCl.  The dilution regimen does not have to be exactly as follows;  it is just an example:
Tube 1:  6 M HCl (undiluted)
Tube 2:  Starting with 6M HCl, make a 1:2 dilution (1 part HCl, 1 part water) for 3 M final conc. of HCl.
Tube 3:  Starting with 6M HCl, make a 1:6 dilution  (1 part HCl, 5 parts water) for 1 M final conc. of HCl.
Tube 4: Starting with 6M HCl, make a 1:60 dilution (1 part HCl, 59 parts water) for 0.1 M final conc. of HCl
Tube 5: Starting with 6M HCl, make a 1:600 dilution (1 part HCl, 599 parts water) for 0.01 M final conc. of HCl.  At 15 drops per mL, this would be 1 drop of 6M acid added to 39.9 mL of water.
One drop of acid solution from Tube 1 is added to the first well of the spot plate;  one drop from Tube 2 is added to the second well of the spot plate;  and so forth.  Then a fixed amount of cabbage extract is added to each well.
The reason we do it this way instead of just adding increasing amounts of very dilute HCl is that we are using such a small volume of sample.  The goal is to use the same number of drops of liquid per well, so that the final volume is some fixed number of drops.
Try to calculate the hydrogen ion concentration in the final sample if you assume the cabbage extract has no buffering capacity.  Check the pH with either pH paper or a calibrated meter, if available.

2C-4. Repeat the process on the other wells or test tubes, this time using base (NaOH or KOH) in increasing dilution.

2C-5. To about 1 mL of the solution from 2B-2 , slowly add just enough aqueous alkali to titrate the indicator back to its natural purplish color. (Does the indicator still work after it had been treated with boiling acid? Why or why not?)


Quinoid resonance structures of Cyanidin Left:  Quinoid resonance structures for the cyanidin aglycone, resulting when the flavylium structure loses a proton from one of its OH groups.  This happens at neutral pH or slightly above.  To retain maximum stability after loss of the H+, the molecule assumes the quinoid form.  The structure on the bottom is more commonly encountered in the literature.

At very high (strongly alkaline) pH, the molecule loses yet another hydrogen ion and assumes the quinoid anion form, in which the hydroxyl on the outlying ring is deprotonated and takes on a negative charge.


 

2D. Paper Chromatography
We can use the typical method of spotting some of the material onto a strip of filter paper (or even ordinary white paper) and then letting a solvent or mixture of solvents carry the pigments up the paper.
A good choice of paper is fine to medium-porosity, qualitative filter paper.

2D-1. Cut strips of the paper about 5-6 cm wide and about 10-12 cm tall, or if using a smaller disc, as big as the piece will allow. Obtain a suitable jar or beaker which is tall enough to accomodate the paper strip while covered.  The paper strip must not touch the sides of the vessel.
You can also do paper chromatography in a test tube, provided you cut the paper strips thinly enough.  The paper must not touch the sides or the solvent will take a "shortcut".   It is better to use a container with some room around the paper.
The solvent level must be somewhat below wherever you are going to spot the paper with sample.

2D-2. Dip the end of a capillary tube into the cabbage extract and spot this liquid down near the bottom edge of the paper strip (not too close-- see above). Across the bottom, in a neatly-spaced row, you may want to spot the different extract samples: water extract, acetone extract, acid-alcohol extract, hydrolyzed acid extract. It will be useful to compare their relative mobilities, especially the acid-hydrolysed one with the non-hydrolyzed one.

2D-3. Pour enough of the desired organic solvent into the tall jar to cover the bottom. It may require some different trials to get the best separation. A good starting mixture is either 1-butanol : acetic acid : water (4:1:5) or acetic acid : HCl : water (5:1:5) if you wish to try duplicating the results of Hedin, et al. (1967). Whatever solvent mixture you devise, it should dissolve the anthocyanins just enough to carry them up the paper, but not so thoroughly that they stay with the solvent front.  On the other hand, the solvent must have at least some affinity for the anthocyanins, or else they'll never progress beyond the initial spot.  In other words, the anthocyanins should have roughly as much affinity for the paper as they do for the solvent, otherwise they'll never separate out of the solvent as spots on the paper.  
Be sure to keep track of the solvent front and any visible spots.  Record the exact proportions of solvents used-- this is important. 
The presence of acid in the solvent system will mean the anthocyanins will be mostly in the oxonium / flavylium cation form.  However, keep in mind that this is in equilibrium with other forms (chalcone, quinoid) that may show up as separate spots. 

2D-4. Not all the anthocyanin spots will be readily visible. However, these compounds absorb short-wave UV light (254 nm) and will therefore show up as dark spots when a UV lamp is shined on the paper. This is especially effective if computer or writing paper was used, because these contain fluorescent whiteners (unfortunately, these types of paper may not give good spot separation without considerable experimenting to discover the right solvent mixture). Look for the dim or dark spots and circle them with a pencil.  The solvent front should have carried enough UV-absorbing components with it that you can locate it again if you lost track of where it was;  it may be very faint, even in the UV.

2D-5. Anthocyanin spots may be made more visible after development (not before, or you might immobilize them) by spotting with lead acetate solution (warning: toxic!). For example, Fuleki and Francis (1964) describe this method.  Their results indicate that lead acetate gives some color differentiation of different anthocyanins and their glycosides (1964).


2E. Absorbance Measurements
The characteristic absorbances of anthocyanins can be measured, at least in the long-wave UV and visible ranges, with a spectrophotometer.
It would be interesting to compare the UV and VIS absorbances of a given anthocyanin extract against (1) extracts made via different procedures from the same red cabbage; (2) extracts from other reddish-purple plants suspected to contain cyanidin; and (3) "dummy" solutions of synthetic food dyes mixed to approximate the hue and visible color intensity of a red cabbage extract. Additionally, the solutions left over from 2B-2 through 2B-5, above, could be checked.
Absorbance in the 500-540 nm range as a measure of anthocyanin content is common in the literature surveyed. See, for example, Piccaglia, et al. (2003); Hedin, et al. (1967); Timberlake and Bridle (1973). The latter source also suggests measuring the IR absorbance, with characteristic peak at 1640 cm-1 (1973).


2F. Elution or Adsorption chromatography
Elution chromatography is a promising method for separation of the cabbage anthocyanins from interfering compounds. Several investigators have used it and continue to do so.  Timberlake and Bridle (1973) give a survey of the methods tried up until that time; they mention elution chromatography and also thin-layer chromatography.
It may be interesting in some future newsletter article to try the method of Coutinho, et al. (2004) using a poly-{methyl methacrylate} resin.  In the meantime, you can experiment with a sort of rudimentary form of elution / adsorption chromatography with the following steps:

2F-1.  Filter some aqueous cabbage extract through filter paper after hydrolysis with HCl.  Much of the pigment should adhere to the cellulose fibers of the paper.  For better results, soak multiple shreds of filter paper in the extract, allowing them to absorb as much color as possible.

2F-2.  Wash the reddened filter paper with cold water to wash away sugars and other water-soluble compounds.  Note that water does not release the pigment in any significant amount (provided you've completed acid hydrolysis of the glycosides).  The pigment at this stage consists primarily of the cyanidin aglycone, which is not all that water-soluble.  It has much more affinity for the stationary phase (cellulose) than the mobile phase (water). 

2F-3.  Next, wash the filter paper with acetone into a clean beaker.  The acetone acts as the eluent;  it will release the pigment almost entirely, giving a reddish solution.  You may want to let some of the acetone evaporate slowly at room temperature in the dark.  Then you can try growing crystals with this solution.

Now, supposing we were to cut the filter paper into small pieces and pack a column with it, we'd have a very simple elution chromatography column.  Because wet paper shreds can clog the column too easily, it would be better if we could have the cellulose in the form of little beads (as in "real" column chromatography.


2G.  Other possibilities
Even the few methods we've touched on can provide a great deal of future study.  What happens to the anthocyanins in the presence of various transition metal ions, for example?  When treated with potassium chromate, the reddish-purple extract (in acid-alcohol) turns very dark brown, almost black.  Some flower-shaped crystals begin to grow (probably chromate), visible at 100x on a microscope slide.  Does the chromate completely degrade the anthocyanins and other polyphenols, or does it produce any interesting oxidation products?  

microphotograph of red cabbage extract treated with chromate Microphoto of red cabbage extract treated w/ chromate


3.  Conclusions and Discussion

Most of the techniques tried in this article were successful after only one or two attempts.  The major exception proved to be the crystal growth experiment.  Growing crystals of a particular phytochemical has inherent complexities and pitfalls that aren't present in growing, for example, sodium chloride and other inorganics, or even in growing crystals of certain organic compounds.  Biological compounds in general have several features that make crystal growth an uncertain proposition:  (1) larger molecules having more complex hydrophobic and hydrophilic entanglements with interfering substances;  (2) difficulty in separating the desired compound from the rest, resulting in doubtful purity of extracts;  (3) within the same group of desired compounds (e.g., the anthocyanins) there can be considerable variation in the attached sugars and other organic molecules-- such a jumble doesn't lend itself to the ordered structure of a crystal.  
It is quite easy to find a seemingly mundane object around the house or garden for which there turns out to be years of advanced research in store.  A head of red cabbage is one such object.  A student, teacher, or hobbyist can use simple experimental techniques to uncover questions that still have no known answer.  Conversely, even the techniques which seem extremely easy at first glance (such as crystallizing the anthocyanins) require a great deal of effort and patience, and even then they don't always go as planned.
The components of red cabbage still hold many secrets and experimental challenges; this article was only a look at some simple extraction and characterization techniques which have proven successful in the past.   We may  explore a couple of these techniques in greater depth at some later date;  paper chromatography, for example, could have an entire newsletter article devoted to it.




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Notes - "Anthocyanins from Red Cabbage"

1The USDA source also seems to have the most comprehensive list of chemical constituents of Brassica oleracea var. capitata f. rubra that's available on the Internet. Other interesting phytochemicals it lists in red cabbage include kaempferol, beta-sitosterol, methylamine, methoxybrassitin, and campesterol. Back to article

2Note the seemingly inconsistent usage of "glucoside" and "glycoside". The prefixes "gluco-" and "glyco-" are synonyms, at least in theory. In practice, "glycoside" often refers to a generic sugar linkage that may or may not be specifically with glucose, while "glucoside" typically refers to a linkage with glucose in particular (as opposed to arabinose, galactose, sophorose, etc). Not all sources stick to this convention, though.  Back to article

3If you do choose to boil the water to speed the extraction process, do not use a test tube. Cabbage leaf pieces will choke up the tube and lead to sudden pressure buildup that inevitably results in ejection of boiling liquid from the test tube.  Back to article

4Generally speaking, do not use small diameter test tubes to boil liquids. The smaller the test tube, the greater the likelihood of "bumping". 10, 12, and even 13 mm test tubes are pretty much out of the question (though boiling chips should still be used just in case cautious heating results in unwanted boiling).  The depth of the liquid relative to the tube's circular cross-section has a great effect on the likelihood of "bumping";  if the tube is very narrow and has a great deal of liquid, forget it.  A wide tube with a relatively small amount of liquid in the bottom is the best situation for heating, though evaporation may be rapid.  Back to article


References - "Anthocyanins from Red Cabbage"

Coutinho, M. R., M. B. Quadri, R. F. P. M. Moreira, and M. G. N. Quadri. "Partial Purification of Anthocyanins from Brassica oleracea (Red Cabbage)." Separation Science and Technology 39 , 3769-3782 (2004).

Fuleki, Tibor and Francis, F. J. "Lead Acetate as Chromogenic Reagent for Anthocyanins". Phytochemistry 6, 1161-1163 (1967).

Hedin, P.A., J.P. Minyard, A.C. Thompson, R.F. Struck, and J.Frye. "Constituents of the Cotton Bud - VII: Identification of the Anthocyanin as Chrysanthemin." Phytochemistry 6, 1165-1167 (1967).

Merck Index. 10th ed. Rahway, NJ: Merck & Co., Inc., 1983.

Onslow, Muriel. The Anthocyanin Pigments of Plants, 2nd Ed. London: Cambridge University Press, 1925.

Piccaglia, R., M. Marotti, and G. Baldoni. "Effects of Fertilization and Environmental Conditions on Anthocyanin Content of Red Cabbage" [Online PDF File ]. Department of Agroenvironmental Science and Technology, Bologna University, Italy. 2003.

Timberlake, C. F. and Bridle, P. "The Anthocyanins". In The Flavonoids , eds. J. B. Harborne, T. Mabry, and H. Mabry. New York: Academic Press, 1973. 215-266.

USDA, ARS, National Genetic Resources Program. Phytochemical and Ethnobotanical Databases. [Online Database ] National Germplasm Resources Laboratory, Beltsville, Maryland. 25 March 2005.






While the information in this newsletter is thought to be accurate to the best of the authors' current knowledge, it is not guaranteed to be free of errors or to be suitable for any particular use. The procedures and experiments outlined within can be dangerous or even fatal if carried out improperly. If you choose to attempt any of them, you proceed entirely at your own risk.


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