Spinach Extraction - Column Chromatography PDF

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Download Spinach Extraction - Column Chromatography PDF

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EXPERIMENTS 5: Basic organic chemistry laboratory techniques: column chromatography Column Chromatography In this experiment you will continue to employ extraction and TLC techniques as you further explore purification techniques. Introduction: As is common in most plant leaves, spinach leaves contain a variety of colored pigments used in the capturing of light for photosynthesis. Generally these pigments fall into two main classes: chlorophylls and carotenoids. Chlorophylls a and b are green pigments that consist of highly conjugated (many alternating double bonds) chemical structures. The carotenoids are part of a larger collection of chemicals known as terpenes, which naturally occur as compounds containing 10, 15, 20, 25, 30, or 40 carbon atoms. Carotenoids such as lycopene or β-carotene are typically responsible for the red or orange color typically found in fruits and vegetables such as tomatoes, watermelons, and carrots. Additionally, β-carotene is a major dietary source of vitamin A, which is a major product of its decomposition upon digestion. In addition to these major pigments, spinach leaves also possess other pigments such as xanthophylls and pheophytins in smaller quantities. In this lab, you will work to separate at least the two main classes of pigments from each other based on the differences in their polarity. You will then analyze the success of your separation using TLC and spectrophotometry. Column Chromatography The structures of the primary pigments can be seen below. In order to separate these compounds, you will take advantage of the polarity differences they possess. This will be accomplished using a method known as column chromatography. Column chromatography works very similarly to TLC, but it is used on a much larger scale to obtain isolatable fractions of what previously just showed up as spots on a TLC plate. Like in TLC, column chromatography also often uses silica gel or alumina as a stationary phase and the mobile phase can be modified in polarity to control the movement of certain compounds down the column. One of the main advantages of this method is that, unlike in TLC where a plate must be run with one solvent, column chromatography allows the experimenter to change the solvent during the separating process in order to more cleanly separate compounds and also speed up the process.

Figure 1. Molecular structures of chlorophylls a and b as labeled on the left and β-carotene on the right. Safety:

Many chemicals used in the lab have health, environmental, and physical safety hazards. Wash hands thoroughly with soap and water before leaving the laboratory. Be sure to dispose of all materials in the appropriate waste containers and not in the sink. In general if any chemical comes in contact with your skin, immediately wash the infected area and notify your instructor. All experiments throughout the semester will be conducted in fume hoods and all PPE (flame-resistant lab coats, nitrile gloves, and safety goggles/glasses) must be worn at all times. Please review all current Material Safety Data Sheets for additional safety, handling, and disposal information as linked in the materials list below. Materials:        

Spinach Mortar and pestle Sodium sulfate Acetone Pasteur pipette and bulb Centrifuge tubes Centrifuge Hexane

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Test tubes Cotton Alumina Sand Methanol TLC jars TLC plates Forceps Capillary tubes

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Filter paper/Coffee filters (for TLC chambers and gravity filtration) UV-lamps Vernier Spectrovis Cuvettes

Procedure: 1. Weigh about 0.5-1.0 g of fresh spinach leaves (avoid stems and thick veins) and tear them into small pieces 2. Place the spinach in a mortar with ~0.5 g anhydrous sodium sulfate and ~2.0 mL of acetone 3. Grind this mixture with a pestle until the spinach leaves are too small to be clearly seen (you may use an additional 0.5-1.0 mL of acetone if too much evaporates during the grinding process) 4. Using a Pasteur pipette or spatula, transfer the mixture to a centrifuge tube 5. Rinse the mortar with 2 mL of cold acetone and transfer this to the same centrifuge tube 6. Centrifuge the mixture, making sure the centrifuge is balanced before turning it on 7. Add ~2.0 mL of hexane to the centrifuge tube, cap the tube, and shake the mixture thoroughly 8. Add ~2.0 mL of water and shake thoroughly with occasional venting 9. Centrifuge the mixture again – you should now see two layers. The top layer is a dark green hexane mixture containing most of your pigments. The bottom layer is the aqueous layer in which much of the acetone will have dissolved 10. Using a dry Pasteur pipette, carefully transfer the top organic layer into a clean test tube 11. Add another ~1.0 mL hexane to the centrifuge tube containing the aqueous layer, then cap and centrifuge this mixture 12. Again withdraw the top organic layer and transfer it to the organic test tube 13. Add a small amount (~0.5 g) of anhydrous sodium sulfate to the organic test tube until some of the particles are free-floating when shaken or swirled 14. After standing for about 5 minutes, transfer the hexane mixture to another clean test tube labeled “E” for extract 15. Rinse the leftover sodium sulfate with ~0.5 mL of hexane, swirl, and transfer the liquid to the “E” test tube 16. Transfer a small amount of this extract (~0.5 mL or less) to a small vial also labeled “E” for TLC 17. Clamp a clean, dry 5 ¾” Pasteur pipette vertically on your ring stand and push a small, loose cotton plug to the bottom 18. Weigh out ~1.25 g of alumina and use this to pack the pipette “column” 19. Gently tap the column to even out the alumina layer, then add a little less than 0.5 cm of sand to the top of the column and tap it again

NOTE: Once this part of the procedure is started, it should not be stopped until the column is completely finished! The alumina must be kept wet with solvent at all times. Since the column does not have a stopcock to control solvent flow, obtain separately labeled containers of 15 mL hexane, 15 mL 70% hexane-30% acetone, 15 mL acetone, and 10 mL 80% acetone-20% methanol solutions. Also label six test tubes 1-6 and a beaker as “waste solvent” 20. Once you are ready to begin your column separation, place the waste beaker under the column and carefully add a few mL of hexane to the top making sure not to force the sand layer into the alumina 21. Continue adding hexane as necessary until all of the alumina is wet and some of the hexane has begun draining into the waste beaker 22. Once the hexane has reached the top of the sand layer, add your spinach extract to the top of the column 23. As the extract pushes onto the alumina, you will likely see the separation into green and yellow bands 24. If separation is seen, add a few mL of hexane onto the column once the last of the spinach extract is right at the top of the sand and continue draining the solvent into the waste beaker until the yellow band reaches the bottom of the column 25. At this point, switch to test tube 1 collecting the yellow solution and continue to do so, adding more hexane to the top of the column as necessary, until the solution turns clear again. You may switch to test tube 2, 3, etc. as necessary when collecting this band. Once the solution is clear, swap the test tubes for the waste beaker 26. If the yellow band does not separate with pure hexane, switch to the next more polar solvent (70% hexane-30% acetone) when conducting steps 24-25. When changing solvents, do not add the new solvent until the level of the last solvent is at the top of the sand 27. Once the yellow band has completely eluted, add several mL of the next more polar solvent as directed in step 26 28. Now repeat this same chromatography practice to collect the green band, again upping the polarity of the eluent as necessary to make the green band move. Be sure to collect this band within your unused labeled test tubes 29. Setup a warm water bath using your hotplate and a small or medium sized beaker 30. Using the bath, evaporate the solvent out of (one of) your green test tube(s). If you had more than one green test tube, combine them all into one as it evaporates down. Do the same for your yellow tube(s). 31. Re-dissolve the green test tube and yellow test tube in ~3 mL hexane 32. Obtain a cuvette and prepare a blank by filling it approximately 2/3 with hexane and wiping the outside of the cuvette with a Kim-wipe 33. Handling only the top edges of the cuvette, gently tap the cuvette against the lab bench to dislodge any air bubbles and position the cuvette into the spectrometer so that the beam of light will shine through the clear sides of the cuvette 34. Turn on the LabQuest and attach the spectrophotometer via the USB cable then choose new from the File menu. The spectrophotometer should have been auto-detected by the device 35. Under “Sensors” select Calibrate and then USB: Spectrometer 36. Calibrate the spectrophotometer with the hexane blank so it gives an absorbance reading of 0. Allow the warmup and calibration to run for the full 90 seconds and then choose Finish Calibration and press OK when the “Calibration completed” message appears 37. Dump the hexane blank into your waste organic beaker and carefully add the yellow solution to the cuvette so that it is approximately 2/3 full 38. Place the cuvette into the spectrophotometer and press play to obtain a full spectrum of this pigment 39. Press stop, then go to File -> Email -> Text File and send yourself a text file of the absorbance vs. wavelength data 40. Pour the cuvette contents back into the test tube 41. Rinse the cuvette with a small amount of the green pigment and dump this into the waste beaker 42. Repeat steps 38-40 with the green pigment (now dissolved in hexane) 43. For TLC, concentrate your yellow and green samples down to ~1 mL using the warm water bath 44. Draw a pencil line ~1 cm from the bottom of your TLC plate and indicate the spotting location for your mixed extract as well as your two test tubes

45. Gently spot the three solutions on these locations using a different TLC capillary for each one and being sure to get a small concentrated sample of each – the spots should be fairly dark 46. Obtain a chromatography jar and add enough 70% hexane – 30% acetone to coat the bottom of the jar but not so much as to be over the spotting line of your plate 47. Run the TLC until the solvent is most of the way up the plate then carefully remove and quickly mark the solvent front in pencil 48. Circle the visible spots and use the UV lamp to aid in observing any others 49. Calculate the Rf values for each of the observed pigments and identify them using the following information: a. In order of decreasing Rf values, pigments in spinach are: i. Carotenes (yellow-orange, 1 spot) ii. Pheophytin a (gray, may be nearly as intense as chlorophyll b) iii. Pheophytin b (gray, may not be visible) iv. Chlorophyll a (blue-green, more intense than chlorophyll b) v. Chlorophyll b (green) vi. Xanthophylls (yellow, possibly 3 spots) Waste:  

All remaining extracts and organic solvents should go in the organic waste container Columns should be placed in a plastic bag fully intact for proper disposal

Lab Report: Please follow the lab report guidelines posted on Canvas. A few things to be sure to include in your report are the following:       

Discussion of reason for separation of compounds based on polarity A detailed image of your TLC plate showing pigment identification and Rf values Properly labeled graphs showing the full spectrum absorbance of the green and yellow fractions from your separation Discussion of the absorbance maxima observed in these graphs based on properties of color and light General comments on success of separation and why/why not it was successful Improvements that could be made to enhance the separation of the pigments found in spinach Discussion of how column chromatography can be used in experiments where bands are not colored...