CR-Scientific Minerals & Experimental Science newsletter
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Issue #10
In this issue:
I. How Hot is a Propane Torch?
II. Anthocyanins from Red Cabbage - with Experiments
III. Site News
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.
I. How Hot is a Propane Torch?
The use of the old-fashioned blowpipe in mineral
analysis requires considerable skill and patience to master. For many simple
mineralogy tests, a propane torch flame is highly satisfactory. The disposable,
blue cylinders of propane gas are available at hardware stores all over the
place. The most common torch nozzle gives a flame about 10 to 11 mm wide at
the base and having a distinct blue cone surrounded by a semi-luminous envelope.
Sometimes this nozzle is called a general-purpose, soldering, or "pencil"
torch1.
Just how hot is this flame? What is it capable of melting? It turns out
the latter question is much easier to answer; simply experimenting with minerals
having known melting points (using shards of uniform size) will provide a
useful list. The former question, unfortunately, is not so easy.
Some might find it tempting to use theory to calculate how hot a propane
torch flame will be. To get an estimate that doesn't appear ludicrous (i.e.,
plumber's torch nearly twice as hot as surface of the Sun), one has to consider
how much heat energy the burning propane-air mixture is losing to the surrounding
air. The brass torch nozzle also gets hot -- really hot-- meaning it, too,
is absorbing heat. Obviously, not all the heat energy of combustion of C3H8
in air is going toward "heating up the reaction products"... which in turn
would consist of nothing but carbon dioxide and water, if only reality were
as simple as the theory.
There are plenty of references available that give the burning temperature
of propane in air. The problem is, they don't often agree on just what
that temperature is. One source shows a peak adiabatic temperature for the propane-air
flame of roughly 2260-2280 °K (CIT, 2003). This is about
1987-2007 °C (3608-3644 °F). Another source gives the adiabatic temperature
as 1977 °C, or about 3590 °F (Babrauskas, 1997). "Adiabatic" means
no heat transfer to the surrounding air or any external objects... in short,
an idealized situation which can't exist. Anvilfire.com 's Gas Facts page lists
the propane torch flame in free air at 2950 °F, or about 1620 °C.
This is much more of a realistic figure, obtained possibly from optical pyrometry.
Other sources, too numerous to list, publish greatly divergent temperatures
ranging all the way from several hundred °C (most likely an open fire
of slowly-escaping propane, not a torch flame!) to upwards of 2000 °C.
The actual flame temperature depends on several factors, including: the nozzle
shape; the fuel-to-air ratio; the velocity of the fuel-air mixture; and what
part of the flame one is measuring. Assuming the hottest part, there are
still some variables to consider.
We can ask some relevant questions, but the answers are tough to quantify.
Some are downright intractable. We might wonder:
- How many moles of propane gas are combining with oxygen per second at any given region in the flame?
- How many kilojoules of energy are there per second at any given region of the gas flame?
- What is the size and shape of the region where the gas particles have maximal kinetic energy?
- How much of the heat energy of the flame is heating the brass torch tip or the surrounding air?
We started by asking, "how hot is a propane torch?", which seemed to be
an easy question. It turned out to be extremely intricate. To map out the
heat distribution quantitatively is beyond the capabilities of all but highly
specialized facilities. That's one reason we didn't find the answer
to our initial question printed all over the place, on the pages of every
textbook and on every website. The mathematical treatment of flames,
even the simplest kind, is an extremely complex proposition. How "hot"
the flame is cannot be a universal value that's somehow the same for all
conditions.
Let's go back, therefore, to what we know for a propane torch. We have the
published maximums for free-air: approximately 1600-1900 degrees C... at
the hottest portion of the flame... with what we assume to be a "typical"
torch nozzle... under the right conditions... and depending on what published
source one prefers. So really, what else can we know for certain? It
is safe to say that an object of any significant size will not melt in the
propane flame if its melting point is close to the maximum flame temperature.
Once again: the flame temperature drops as soon as we introduce something
into it. We can also be quite sure the flame is not 2000-2500 °C (the
adiabatic temperatures according to different sources); these do not consider
the heat-draining effects of the torch nozzle or the air.
So how much does the temperature drop, then, when we heat an object?
We must take into account the thermal properties of the object. Let's assume
we started at a free-air temperature of, say, 1600 °C, and let's narrow
it down to a very specific object: a 7 x 3 x 1.5 millimeter chip of
actinolite into the flame, held by stainless steel tweezers.
Let's focus on what we'll call a "practical temperature".
We decided to try a little experiment based on the knowledge that actinolite
melts at 1200 °C (see, for example, Hibbard, 2001). A small chip of this
mineral (7 x 3 x 1.5 mm, as mentioned above), held by the thinnest steel
tweezers possible, with as little of the steel in the flame as possible,
was heated at the tip of the blue cone for about five minutes. Actinolite,
having a melting difficulty of 4 on the 5-point fusibility scale of minerals
(Warren, 1921), is supposed to fuse along thin edges b.b. ("before the blowpipe").
The plumber's torch managed it, but barely. The rounding was visible only
on a corner of the chip at 10x magnification. This fused area was only about
three-fourths of a millimeter across.
Based on this information, we'll treat the "practical temperature" of the
propane torch as no more than 1200 °C, and only on small objects. To
be fair, we couldn't have arrived at such a sweeping figure without having
a blatant disregard for factors such as heat capacity, thermal conductivity,
and thermal diffusivity. Minerals do not conduct heat away as efficiently
as most metals. They also have different rates of emissive heat loss
compared to metals. It also takes different amounts of heat to raise
a given volume of, say, actinolite, to a desired temperature as compared
to a given volume of, for example, aluminum.
This article couldn't even pretend to descant
the vast subjects of flame thermodynamics and combustion science. 2 It arrives
only at some generalized truths: a huge torch flame will be "hotter" in every
practical sense than a tiny torch flame, given the same fuel-air mixture.
In a nozzle of fixed size, more fuel/air mixture flowing through it will
make a hotter flame. If there is more oxygen in the air stream, there
will be more collisions between fuel and oxygen molecules, resulting again
in more heat.
If one could make a big and concentrated enough
flame out of a sufficiently energetic fuel, surrounding it in an environment
that prevented the heat from escaping too rapidly, one might be able to melt
a walnut-sized chunk of actinolitecompletely (even if the goal weren't a
boiling, white-hot, miniature sun, this melting such a rock would require
a considerable amount of heat 3) With a typical propane torch, however,
a mineral chunk of walnut size is too much of a heatsink even to begin melting
it, unless it's something like stibnite.
A large chip will drain more heat away from the desired area than a small
one, so obviously it's wise to make the chip as small as is practical. No
walnut-sized chunks; not even fingernail-sized chunks.
There are two additional ways to lessen the inevitable heat loss from a
flame. The first, as mentioned before, is to use a bigger flame; but that
would mean a different nozzle type and / or a different fuel mixture, which
is outside the focus of the article. The second is to use something that
will contain the heat around whatever it is you're heating.
This brings to mind the age-old technique of making a kiln to contain some
of the heat and reflect it back toward the center. Supposing there
is at least a small portion of the flame that has a temperature of 1600 °C
or thereabouts; we can put this to work much more effectively with some firebricks
fashioned into a miniature oven. It might increase the size of the
chip that the flame can effectively melt, if only by a little bit.
Notes - "How Hot is a Propane Torch?"
1 The so-called "vortex" torch tips give a hotter flame and also usually provide built-in ignition mechanisms, but these tips are not as useful in mineral analysis. They produce a highly turbulent flame that lacks a clear demarcation between oxidizing and reducing areas (i.e., no "blue cone"). If it's simply melting you're after, the vortex tips are suitable. Back to article
2Mathematically modelling fires and flames can be a little off-putting for all but the mathematician. Have a look at http://fire.org/Download/Catchpole_spread.pdf , for example. For another heavily math-oriented page, have a look at http://www.win.tue.nl/casa/research/casaprojects/graziadei.html .
Back to article
3 Though "miniature sun" is a facetious term, turning common objects into miniature suns is something of which many science-minded people have dreamed. The most elusive part of the thing, of course, is to make it supply its own energy.
As for heating an oversized object with an undersized torch, this fuel-wasting mistake is common among beginners.
Back to article
References - "How Hot is a Propane Torch?"
Babrauskas, Vytenis. "Temperatures in Flames and Fires", Fire Science and Technology, Inc. [ Online article ]. 1997.
CIT (California Institute of Technology), Mechanical Engineering Department. "ME 96: Flame Temperature Measurement Experiment". [ Online PDF File ]. March 2003.
Hibbard, M. J. Mineralogy: A Geologist's Point of View. New York: McGraw-Hill, 2001.
Warren, Charles H. A Manual of Determinative Mineralogy. New York: McGraw-Hill, 1921.
II. Anthocyanins from Red Cabbage - With Experiments
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.
It is rightfully so that this experiment is so popular, for it 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. (Update: the
full name for red cabbage is Brassica oleracea var. capitata f. rubra; we had erroneously left
it as simply "Brassica oleracea",
which could have meant anything from wild mustard to kohlrabi. A red
cabbage could certainly be called Brassica
oleracea, but so could broccoli, kale, Brussels sprouts, etc.
They are actually all the same species, just different varieties and cultivars.)
Another property of "cabbage indicator" is that the study of it 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 this true? Wouldn't red cabbage be exhaustively
studied, even trite as a subject for a resarch 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).
Drawing from subsequent 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 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 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",
the teacher or amateur chemist can hope only to end up with a preparation
having as few contaminating substances-- in other words, non-anthocyanins--
as possible via the extraction means at hand. Below the university upperclass
level, we'll assume these 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 oygen 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.
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.
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.
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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.
It is worth further investigation to find out how much this happens at -20
C; it is, however, worth noting that the original source (1968) does
contain this procedure. Its authors may have overlooked that point.
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's odd 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. |
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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). |
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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. |
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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. |
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 very 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?)
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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 filter paper (Whatman #1
or #2, for example).2D-1. Cut strips of the paper about 5-6 cm wide and about 10-12 cm tall. 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".
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.
2F-2. Wash the 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?
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 will
probably 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.
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.
III. Site News
First off, a note about the newsletter: The program we were
using to make the HTML newsletter was an old one that had the bizarre property
of making the text jump around on the screen as one typed. The program would
try to scroll, but it didn't do it right-- instead of acting like a word
processor, it made the lines march up the page and wrap around again at the
bottom of the screen. It became virtually unusable with large amounts
of text (such as encountered here). During this strange scrolling, the program
sometimes also inserted typed letters in the wrong places-- as if the words
jumped out of the way before the letters were put in place. This resulted
in a great deal of unwanted re-typing. Overall, it made producing the newsletter
unbelievably time-consuming (and rough on the eyes).
About 50% of the current issue was produced using the old program before
it finally became unbearable. The good news is, we've switched to a
different editing program for HTML. It does have some bugs, but at
least the strange scrolling/marching effect is gone. That means easier
typing. That also means, likely, more frequent issues once again (at
least better than the once or twice a year it was turning into).
We're hoping to get back on track with the newsletter and come out with issues
more often. If you've any comments, suggestions, or corrections for
the newsletter, don't hesitate to email us.Product updates:
We recently got some new carbon blocks in stock. These new ones are
somewhat smaller, which should help lessen the tendency to drain heat away
from the sample (Readers might have been considering size of the carbon block
as another variable in the article about propane torches, above. It is sheer
coincidence, though, that this article and the smaller carbon blocks are
mentioned in the same newsletter issue).
The site also has some other recent additions, with more on the way -- have
a look!Reader feedback & suggestions:
We've had some requests for articles featuring proper uses of various glassware
products, with examples. This will be a possibility for upcoming newsletter
issues. We've also had requests for experiments in other fields we
haven't yet touched on, such as physiology. We'll also keep this in
mind for future issues, wherever possible.
A reader has recently shared some experimental results and questions relating
to mineralogy, which we hope to include in the next newsletter issue.
Keep those suggestions coming in!That concludes this issue of the CR Scientific Newsletter. Until next time, stay safe and have fun!
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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