Background: The most modern method of separating mixtures in organic chemistry is chromatography. Chromatography is defined as the separation of a mixture of two or more different compounds by distribution between two phases, one of which is stationary and the other moving. The method depends on the different solubilities, or adsorptivities, of the substances to be separated relative to the two phases between which they are to be partitioned. In thin layer chromatography (TLC) the ‘stationary’ phase is the adsorbent silica, which is bound to an aluminum-backed plate (also called a TLC plate). Silica is considered a polar substance since the surface of the crystals consists of polar hydroxyl (OH) groups. The ‘moving’ phase is an organic solvent system that, by capillary action, will move up the stationary silica coated plate. All solvent systems will be considered non-polar relative to the silica adsorbent. The sample mixture is usually applied as a small spot near the base of the TLC plate (called ‘spotting’). The plate is then put into a solvent reservoir where, by capillary action, the solvent will rise up the plate. As the solvent ascends the plate, the compounds in the sample are partitioned between the moving liquid solvent and the stationary solid phase. This process is called developing the TLC plate. When developing a TLC plate, the various components in the mixture are separated. This separation is based upon each compound’s distribution equilibrium between the solvent and adsorbent. See the figure below:
Compound Distribution Equilibrium
Each compound will have a unique distribution equilibrium depending mainly upon the polarity of the compound (based on intermolecular forces between the compounds being separated and the adsorbent). An example is that of a ‘polar’ compound vs. a non-polar compound. The distribution equilibrium of a polar compound will favor the adsorbent since the adsorbent is highly polar (“like dissolves/attracts like”). The ‘non-polar’ compound however, will have less affinity for the polar adsorbent and will have an equilibrium favoring solubility in the mobile solvent. The consequence of this is that polar compounds will ‘stick’ to the stationary TLC plate while non-polar compounds will separate and travel upward with the solvent. When developing a TLC plate we can state that each compound in the mixture will ascend the plate at a different rate; polar compounds ascend slowly, less polar compounds ascend quickly. In this experiment you will isolate a mixture of colored chlorophylls (see last page for structures) from spinach leaves and then separate this mixture into its individual components using TLC.
1. How would using a more polar solvent affect the equilibrium between solid and solution for a very polar compound? 2. Would you expect the above material to elute (move) faster or slower with a very polar solvent? 3. Very polar compounds are sometime purified by reverse phase chromatography where the stationary phase is very nonpolar while the solvent is very polar. Why might one want to use this technique with extremely polar compounds?
Materials: Spinach leaves, acetone, hexane, 70/30 hexane/acetone, anhydrous sodium sulfate, mortar and pestle, 2 centrifuge tubes, Pasteur pipets, hot plate, beakers, test tube, TLC plates, micro capillaries, filter paper, watch glass cover (or aluminum foil), pencil
Safety Precautions: Acetone and hexane are flammable, keep away from flames. Pasteur pipets break easily, handle with care. Get instructions before using the centrifuge.
Procedures: 1. Isolation of Pigments: Weigh about 0.5 g of fresh spinach leaves. Cut or tear the spinach leaves into small pieces and place them in a mortar along with 1 mL of acetone. Grind with a pestle until the spinach leaves have been broken into particles too small to be seen clearly. If too much acetone has evaporated, you may need to add an additional portion of acetone (0.5-1.0 mL) to perform the following step. Using a Pasteur pipet,
transfer the mixture to a centrifuge tube. Rinse the mortar and pestle with 1.0 mL of acetone and transfer the remaining mixture to the centrifuge tube. Centrifuge the mixture (be sure to balance the centrifuge). Using a Pasteur pipet, transfer the liquid to a centrifuge tube with a tight fitting cap. Add 2.0 mL of hexane to the tube, cap the tube, and shake with mixture thoroughly. Then, add 2.0 mL of water and shake thoroughly with occasional venting. It’s important to thoroughly dissolve the pigments in the hexane before adding the water. Centrifuge the mixture to break the emulsion, which usually appears as a cloudy, green layer in the middle of the mixture. Remove the bottom aqueous layer with a Pasteur pipet. Using a Pasteur pipet, prepare a column containing anhydrous sodium sulfate to dry the remaining hexane layer, which contains the dissolved pigments (see the figure on next page). Place a plug of cotton into a Pasteur pipet and tap it into position using a glass rod. Add about 0.5 g of sodium sulfate and tap the column with your finger to pack the material. Clamp the column in a vertical position and place a dry test tube under the bottom of the column. With a Pasteur pipet, transfer the hexane layer to the column. When all the solution has drained, add 0.5 mL hexane to the column to extract all the pigments from the drying agent. Evaporate the solvent by placing the test tube in a warm water bath and directing a stream of air into the vial. Once evaporated, dissolve the residue 7-10 drops of hexane.
Column for Drying the Extract TLC Development Chamber
2. TLC of spinach extract: Obtain two 1-inch TLC plates and micro capillaries from your instructor. These plates have a flexible backing, but they should not be bent excessively. They should be handled carefully or the adsorbent may flake off them. Also, they should be handled only by the edges; the surface should not be touched. Using a lead pencil (not a pen) lightly draw a line across the plates (short dimension)
about 1 cm from the bottom (on the coated side). At the center of this line make a light mark. This is the point at which the spinach extract will be spotted. To spot the TLC plate, fill the capillary tube by dipping one end into the spinach extract. Capillary action fills the pipet. Empty the pipet by touching it lightly to the thin-layer plate at the mark that is at the center of the 1 cm line from the bottom (The spot must be high enough so that it does not dissolve in the developing solvent). When the pipet touches the plate, solution is transferred to the plate as a small spot. The spot should be no larger then 2 mm in diameter and should be a fairly dark green. If you do not have a dark green spot, you may spot again using another sample of your spinach extract. Allow the solvent to evaporate completely between successive applications, and spot the plate in exactly the same position each time. It is important that the spots be made as small as possible and that the plates not be overloaded. When the first plate has been spotted it is ready to be placed in a development chamber. For a development chamber you will use your large beaker lined with a piece of filter paper and cover (see the figure above). When the development chamber has been prepared, obtain a small amount of the development solvent (70/30 mixture of hexane/acetone). Fill the chamber with the development solvent to a depth of about 1/4 inch (about 10 mL of solvent). Be sure that the liner is saturated with the solvent. The solvent level must not be above the spots on the plate or the samples will dissolve off the plate into the reservoir instead of developing. Place the spotted plate in the chamber and allow the plate to develop (the solvent will slowly move up the TLC plates). Since the backing on the TLC plates is very thin, if they touch the filter paper liner of the development chamber at any point, solvent will begin to diffuse onto the adsorbent surface at that point. When the solvent has risen to a level about 1 cm from the top of the plate, remove the plate from the chamber and, using a lead pencil, mark the position of the solvent front. Let the plate dry. Lightly outline all the observed spots with a pencil. Before proceeding, make a sketch of the plate in your notebook and label each spot by color. Using a ruler marked in millimeters, measure the distance that each spot has traveled relative to the solvent front. Under an established set of such conditions, a given compound always travels a fixed distance relative to the distance of the solvent front. This ratio of the distance the compound travels to the distance the solvent travels is called the Rf value. This can be expressed as a decimal fraction: Rf = distance traveled by substance/distance traveled by solvent front (see figure below). Calculate Rf values for each observed spot. Repeat the above TLC of you spinach extract but use a different proportion of hexane/acetone for your developing solvent (be sure to record the proportion used in your notebook). Once this plate has developed, sketch the plate in your notebook, label the spots by color and calculate Rf values for all the spots as described above.
Waste Disposal: Put all extra acetone and hexane solutions into the organic waste. Aqueous solutions can go down the drain. Rinse centrifuge tubes with water and place into the dirty glassware bin. Rinse the mortar and pestle with a small amount of acetone (put rinse into organic waste), rinse with water and put into the dirty glassware bin.
Spinach residue can go into the trash. Pasteur pipets should go into the glass waste container (unless otherwise instructed).
Report: 1. Try to identify each spot on your TLC’s by the type of compound it may be (see next page).
2. In your notebook, write a brief discussion of the experiment. In this section you should summarize your results (number of spots, colors, Rf values), note any interesting observations and make any possible conclusions about the experiment (successful vs. unsuccessful and reasons why).
3. Answer the post-lab questions.
4. Make a copy of your lab notebook pages (do not tear out the originals) and attach the question sheet with your answers. Staple all pages, put your name on the front and turn it in.
Post-Lab Questions: 1. What would happen to the Rf values of the pigments if you were to increase the relative concentration of acetone in the developing solvent (and why)?
2. Why are the chlorophylls less mobile than the carotenes on the TLC plate?
Below are the structures and description of chlorophylls and other components that are in the pigments.
The colored pigments from a plant fall into 2 categories, Chlorophylls and Carotenoids. Caroteniods are yellow pigments that are involved in the photosynthesis process.
Xanthophylls are oxygen-containing carotenes (OH and C=O) The carotenes are shown below. The Green pigments are the Chlorophylls that act as the principal photoreceptor molecules of plants. There are two different forms, chlorophyll a and chlorophyll b. The two forms are identical except that the methyl group that is shaded in the structural formula of Chlorophyll a is replaced by an aldehyde (C=O) group in chlorophyll b. Pheophytin a and pheophytin b are identical to chlorophyll a and b except that in each case the magnesium ion (Mg2+) has been replaced by two hydrogen ions.
On your TLC place you will see in the following in order of decreasing Rf value (top of plate to bottom); Carotenes (yellow), Pheophytin a (grayish), Pheophytine b (grey, may not be visible), Chlorophyll a (blue-green), Chlorophyll b (green), Xanthophylls (up to 3 spots, yellow)