Fordham University TLC & Column Chromatography Lab Report Organic chemistry 1 lab homework assignment based on the concept of TLC chromatography. I need so | Course Hero
Fordham University TLC & Column Chromatography Lab Report Organic chemistry 1 lab homework assignment based on the concept of TLC chromatography. I need someone who is in expert in organic chemistry! I also attached lectures notes and procedure of the experiment to guide the tutor through the problems, so they can better understand how to solve the questions. Chromatography
Chromatography is a technique used for separating the components of a mixture from each other.
Chromatography can also be used to analyze a mixture of compounds and determine how many
components are present in a mixture.
Note: chromatography is based on differential adsorptivites of the compounds in a mixture between a
stationary phase and a mobile phase.
Analogy:
Rain!
Note: In this analogy, the earth is the stationary phase and rain water is the mobile phase.
Note: The opposing forces of the rain water and inertia of the rocks causes different distances to be
traveled by the rocks and thus become separated from each other!
Note: At the molecular level, differences in polarity and hydrogen bonding ability among compounds of
a mixture to the stationary phase causes different compounds to travel different distances upon the
influence of the mobile phase.
Note: The most common chromatographic methods are thin-layer chromatography (TLC) & column
chromatography (CC)
Note: In TLC, the stationary phase is a thin layer (0.1 mm) of silica gel (SiO2xH2O) or alumina
(Al2O3xH2O) supported on a rectangular plate (5 X 10 cm). The TLC plate may be glass, plastic
or a light metal like aluminum.
Note: In TLC, the mobile phase is a liquid which is placed in a capped jar.
Note: In TLC, the sample to be analyzed is placed on the TLC plate using a capillary tube (this is called
spotting the plate) before being placed in the jar (see figure 3-28 on page 154 of lab textbook).
Note: In TLC, the plate is placed inside the jar that contains the mobile phase. The mobile phase moves
upward on the TLC plate by capillary action.
Note: Different compounds present in a mixture move upward on the TLC plate at different rates and
become separated.
Note: The TLC plate is viewed under UV light and compounds that absorb UV light appear as dark spots
on the TLC plate. Or, the TLC plate is placed in a jar containing iodine and some compounds
become stained (yellow/brown) by iodine vapor.
Note: In CC, the stationary phase (silica gel or alumina) is placed inside a vertical glass or plastic
column.
Note: In CC, the sample whose components are to be separated are dissolved in a small quantity of a
solvent and then placed on top of the stationary phase using a Pasteur pipette.
Note: In CC, the mobile phase is added to the top of the stationary phase after the sample has been
applied to the stationary phase. The mobile phase moves downward by gravity.
Note: Since silica gel and alumina are highly polar substances, polar molecules are adsorbed strongly to
the stationary phase while those that are less polar are adsorbed less strongly.
Note: Consequently, less polar compounds move faster while more polar compounds move slower.
Note: Mobile phases that are more polar desorb the components of a mixture more readily while less
polar mobile phases desorb the components less readily.
Order of increasing polarity of mobile phases
compounds
Order of decreasing desorption of organic
Petroleum-ether (mixture of hydrocarbons)
Chloroform (CHCl3)
Methylene chloride (CH2Cl2)
Ether (CH3CH2OCH2CH3)
Ethyl acetate (CH3CO2CH2CH3)
Acetone (CH3COCH3)
Ethanol (CH3CH2OH
Methanol (CH3OH)
Acetic acid (CH3CO2H)
alkanes/cycloalkanes
alkenes/alkynes
ethers (R-O-R)
haloalkanes (R-X)
ketones (RCOR)
aldehydes (RCHO)
esters (RCO2R)
alcohols (R-OH)
carboxylic acids (RCO2H)
Extraction of pigments from spinach
and their
separation by column chromatography
some chlorophyll (Chl) pigments
Spinacia oleracea
some carotenoid pigments
?-carotene
lutein (a xanthophyll)
The pigments found in the leaves of green plants fall mostly into two compound classes: chlorophylls (green/blue-green) and
carotenoids (yellow to orange/orange-red). Spinach leaves contain chlorophyll a & b and ?-carotene as major pigments, as
well as smaller amounts of other pigments such as xanthophylls (oxidized versions of carotenes; note that the structure of
lutein above contains hydroxyl [-OH] groups, which are not present in ?-carotene). The structures of the major pigments are
shown above. ?-Carotene is a nonpolar hydrocarbon, while the chlorophylls (all similar in structure) contain polar C-O & C-N
bonds, and also a magnesium ion chelated to the nitrogen atoms, making them much more polar than??-carotene;
chlorophyll a has a methyl group in a position where chlorophyll b has an aldehyde (-CH=O), making the latter slightly more
polar than the former. After you extract the pigment mixture from spinach leaves you will take advantage of this difference in
polarity to separate the mixture of pigments using column chromatography.
In this experiment you will:
(a) Extract a mixture of pigments from spinach;
(b) Separate the pigments by column chromatography, based on their differences in polarity;
(c) Tentatively identify pigment types by their color and the order they elute from the column.
YOU WILL BE WORKING IN PAIRS FOR THIS EXPERIMENT: ONE PERSON WILL PERFORM THE EXTRACTION WHILE THE OTHER
PREPARES THE COLUMN. PREPARE AND PLAN CAREFULLY IN ORDER TO AVOID COMPLICATIONS AND TO COMPLETE THE
EXPERIMENT ON TIME.
Before coming to lab remember to consult the MSDS for all chemical substances. As always, avoid direct contact
with all chemicals/solvents by working in fume hood as much as possible and wearing gloves, eye protection and a
fully buttoned lab coat. NO FLAMES will be allowed in the lab as the eluents you will be using are highly
flammable.
Part 1. EXTRACTION OF PIGMENTS FROM SPINACH
In a mortar, grind approx. 30 g of fresh or previously frozen spinach in 25 mL of methanol using a pestle. Decant the
methanol solution and put it aside (discard this in the NON-HALOGENATED LIQUID ORGANIC WASTE container at the end of
the lab period). Next, grind the spinach with a mixture of methanol (12 mL) and low-boiling petroleum ether (20 mL; bp 3060 oC). Carefully filter the mixture into a separatory funnel through a small layer of cotton loosely placed in a glass funnel,
leaving the spinach behind in the mortar. Regrind the spinach in the mortar with more methanol (12 mL) and petroleum
ether (40 mL), and then filter the liquid through the cotton in the glass funnel into the separatory funnel. The above
operations will extract most of the water in spinach into the methanol phase and much of the organic pigments into the
petroleum ether phase.
Drain off the lower methanol layer into an Erlenmeyer flask and set it aside (discard this in the NON-HALOGENATED LIQUID
ORGANIC WASTE container at the end of the lab period). Add water (25 mL) to the petroleum ether remaining in the
separatory funnel and shake it for a minute. Allow the layers to separate in the funnel, and then drain off the lower aqueous
phase into a beaker (discard this down a sink drain at the end of the lab period). Repeat this extraction with a second
portion of water (25 mL) and drain off the lower water layer into the beaker containing the first water layer. The purpose of
washing the petroleum ether layer with water is to remove any dissolved methanol.
Pour the petroleum ether layer through the top of the separatory funnel into a 125-mL Erlenmeyer flask and dry the solution
over anhyd. sodium sulphate for 5 min. Decant the liquid from the Erlenmeyer flask into a 100-mL round-bottomed flask and
evaporate the solvent using a rotary evaporator until the volume of the solution is reduced to about 2-3 mL. THIS IS YOUR
CONCENTRATED EXTRACT OF PIGMENTS FROM SPINACH; SAVE THIS SOLUTION FOR THE NEXT PART OF THE EXPERIMENT.
Part 2. SEPARATION OF SPINACH PIGMENTS BY COLUMN CHROMATOGRAPHY
Prepare a chromatography column by first placing a small plug of cotton into the bottom of the
column. Pour in a small amount of sea sand to form a level layer approx. 5-10 mm in height. Weigh
approx. 10 g of alumina into a small beaker, and then fill the column approx. 2/3 with high-boiling
petroleum ether (ligroin; bp 60-90 oC). Slowly pour the alumina through a funnel into the column,
and tap the column with your scoopula to ensure that the alumina packs evenly. Use extra ligroin to
wash down any alumina that is adhering to the inner column surface. After youve added all the
alumina, add a 5-10 mm layer of sea sand to the top of the column packing. From this point onward,
be sure there is always enough solvent in the column to cover the packing; i.e., never let it go dry (!)
and add more solvent as necessary. Open the stopcock at the bottom of the column to allow the
solvent to go just below the level of the sand at the top of the column but not below the alumina,
and then close the stopcock. Add all but approx. 0.5 mL of the spinach extract (from part 1) via pipet
to the top of the column without disturbing the packing. Open the stopcock and wash down any
pigments on the inner surface of the column with ligroin (1 mL) and repeat this twice more, each
time letting the solvent go below the level of the sand before adding the next 1-mL portion. Close
the stopcock and fill nearly the entire column with a mixture of 10 parts acetone to 90 parts ligroin.
Open the stopcock and start collecting column fractions into clean, dry Erlenmeyer flasks.
As you watch the two major groupings of colored bands of pigment (green for chlorophylls and yellow for carotenoids) elute
from the column, attempt to collect them in separate 50-mL flasks while collecting the dead volumes (where no colored
compounds are eluting) into other flasks. Which compound class will elute first from the column: chlorophylls or
carotenoids? After the first band(s) of the same color have eluted from the column into the same flask you can increase the
polarity (eluotropic strength) of the eluent by adding methanol (20 mL) to the column to elute the more polar pigments,
which better adhere to the stationary phase; add more MeOH as needed. After recording your observations/results related
to the fractions youve collected you can discard them in the NON-HALOGENATED LIQUID ORGANIC WASTE. Use a rubber
bulb to force all remaining solvent off the column and discard the used alumina in a container labeled WASTE ALUMINA.
Analysis of Pharmaceutical Drugs by Thin-layer Chromatography (TLC)
Goal: identification of an unknown OTC pharmaceutical drug by TLC analysis in comparison with authentic samples of CAFFEINE,
ASPIRIN, ACETAMINOPHEN, & SALICYLAMIDE. The table below indicates the components present in each of the six OTC drugs, and
each drug has a unique composition. Your unknown will be one of the six commercial tablets shown in the left column.
OTC PHARMACEUTICAL
caffeine
aspirin
acetaminophen
salicylamide
X
X
X
X
X
X
X
X
X
X
X
Note: before using microcapillary tubes to spot dilute solutions of analgesics on your assigned TLC plate, please practice spotting
and visualizing spots on scrap plates that will be made available to you.
Step in procedure
Detailed comments
1) Prepare at least five microcapillary tubes for spotting dilute solutions
of the analgesics on TLC plates.
Microcapillary tube preparation will be demonstrated in the lab. THE
SOLVENTS USED IN THIS EXPERIMENT ARE FLAMMABLE AND THE USE
OF BUNSEN BURNER FLAMES WILL ONLY BE PERMITTED DURING A
DESIGNATED TIME AT THE START OF THE EXPERIMENT. BE SURE
THERE ARE NO FLAMMABLE SOLVENTS NEARBY WHEN USING A
FLAME, AND VICE VERSA.
2) Sign out an unknown pharmaceutical tablet and, after youve
practiced spotting on scrap plates, obtain a 5 x 10 cm silica gel TLC plate
with fluorescent indicator.
Avoid touching the surface of TLC plates; oils from your skin could
contaminate it.
3) Approx. 2 cm from one of the short edges of the plate draw a light
pencil mark across the plates surface.
This line identifies the solvent front, used to indicate how far the eluent
(the solvent(s) used for the mobile phase in TLC) advanced up the plate.
4) Using a pencil, mark a light line across the plate at 1.5 cm above the
other short edge of the plate, and then lightly draw five short hash
marks equidistant from each other and from the edges.
This line identifies the origin, where you will adsorb compounds to the
plates surface.
5) Using a microcapillary tube, spot a methanol solution of your
unknown at the hash mark in the center, and spot each of the four
standards (dilute solutions in methanol of authentic analgesics) at the
other four marks (refer to detailed comments in the right hand column).
Note: analgesic components in the drug tablets are present in unequal
amounts, and a component present in relatively small amount might
show up as an easily overlooked faint mark if you spot too lightly;
therefore, spot the unknown a little more heavily than you would the
authentic analgesics.
The standards are 2-5% solutions in methanol of the following
analgesics: acetaminophen, aspirin, caffeine and salicylamide. Your
unknown is part of a crushed analgesic tablet (one listed in the table
above) provided to you in a small test tube, to which you should add 1
mL of methanol followed by brief swirling of the resulting suspension.
The analgesic components present should all be extracted into the
methanol solvent; however, there will be much material that will NOT
be soluble in methanol, so dont expect the entire tablet to dissolve!
KEEP THE SPOTS SMALL by touching the microcapillary tube gently upon
the plates surface and then immediately withdrawing it. IF YOUR
SPOTS ARE TOO LARGE (as visualized under UV light), TURN THE PLATE
o
BY 180 and spot again, using the solvent front as the new origin. At
the top of the plate lightly write in pencil a small code identifying which
standard is spotted there.
Step in procedure
Detailed comments
6) Prepare a TLC chamber by filling a commercial chamber (a jar with a
screw-cap lid) to a depth of approx. 0.5 cm with a solution (15 mL) of
toluene, ether, glacial acetic acid and methanol (120:60:18:1); this
solution is the TLC eluent (i.e., the mobile phase for TLC). Place a halfpiece of large filter paper around the inner perimeter of the chamber.
KEEP THE CHAMBER COVERED AS MUCH AS POSSIBLE TO AVOID
EVAPORATION OF THE ELUENT, WHICH WOULD CHANGE ITS
COMPOSITION AND POSSIBLY AFFECT YOUR RESULTS.
The paper in the chamber acts as a wick from which solvent can
evaporate and fill the chamber with its vapor. This will decrease the
rate of solvent evaporation off the face of the plate, thereby allowing
the eluent to move more rapidly up plates surface.
7) Use forceps to carefully place the plate in the developing chamber.
Immediately cover the chamber with its lid and leave it undisturbed
until the eluting solvent reaches the line marked for the solvent front
(this should take approx. 10 min or less).
If you measure correctly, THE SPOTS ON THE PLATE SHOULD BE ABOVE
THE SOLVENT LEVEL IN THE CHAMBER. The plate should not touch any
surface along its sides; only the top and bottom of the plate should be
touching the chamber. Position the plate in the chamber such that the
top corners are resting against the filter paper wick. The solvent (which
is the TLC mobile phase) will slowly be drawn up the surface of the plate
by capillary action.
8) When the eluting solvent reaches the pencil line marking the solvent
front, remove the plate from the chamber with forceps.
9) Hold the plate with forceps while drying it with the hot air from a
heat gun, and then examine it under UV light. Use a pencil to lightly
outline the spots you see and note your observations (i.e., number of
spots in each lane, appearance of spots, etc.) in your lab notebook.
DO NOT look directly into the UV light as it will damage your eyes, and
avoid exposing your skin to the light.
10) Place the plate in an iodine (I2) chamber and note which spots are
visualized by iodine and their appearance. Make your observations
immediately since the color of the stain fades fairly rapidly.
CAUTION: IODINE IS A CORROSIVE SOLID WITH A HIGH VAPOR
PRESSURE. AVOID INHALING ITS VAPORS OR EXPOSING YOUR EYES OR
SKIN TO EITHER THE VAPORS OR THE SOLID ITSELF. THE CHAMBER
SHOULD ALWAYS BE KEPT CLOSED BETWEEN THE TIMES THAT PLATES
ARE BEING TRANSFERRED IN AND OUT.
11) Use the ruler provided in your lab drawer to make the
measurements that you will need to calculate Rf values for each spot.
You must record all measurements in your lab notebook while you are in
the lab since you must leave your plate in your lab drawer. Do not
destroy the plate until youve received your graded report form, after
which you can discard the plate in the household waste container.
Rf = distance traveled by a spot_________
distance traveled by the eluting solvent (origin to solvent front)
The abbreviation Rf = ratio to front (or retention factor) and is simply a
measure of how far the compound has eluted, or traveled, up the
plates surface; the distance a spot travels is the distance measured
from the origin to the center of gravity of the spot.
The MEASUREMENTS (i.e., the raw data) must be recorded while you
are in the lab, but the actual Rf value calculations may be performed in
your notebook at a later time outside of the lab. If time permits,
perform the calculations in the lab; otherwise, do them in your
notebook at a later time outside of the lab.
12) Identify each spot you observe in the unknowns TLC lane by
matching their Rf values and visualization characteristics with those of
the authentic standards you spotted on the same plate. By identifying
the analgesic components present in your unknown and referring to
the table on page 1, you can establish which of the six assigned
pharmaceutical drugs it is.
An unknown report form will be posted on our Blackboard website;
please submit the completed form on the due date (to be announced
on Blackboard).
WASTE DISPOSAL:
Please discard the used eluent in the NON-HALOGENATED LIQUID
ORGANIC WASTE container and leave the chamber, along with its
filter paper wick, in your hood with the chamber lid removed.
Discard used or broken microcapillary tubes in a GLASS WASTE
container.
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