state a hypothesis relating polarities and solubilities of pigments

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6.2: Photosynthetic Pigments

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Extract and Separate the Pigments

• Lay a strip of filter paper on the bench.
• About 2 cm from the bottom of the strip, place a fresh spinach leaf and rub a coin across the leaf to transfer pigment to the strip.
• On a separate strip, the instructor will apply the Spirulina extract approximately 2cm from the bottom of the strip.
• Suspend the strips by a dowel or paper clip in a tube with about 3ml chromatography solution (2 isooctane: 1 acetone: 1 diethyl ether ).
• Develop the strips until the solvent reaches about 2 cm from the top.

Chromatography Analysis

• How many different pigments separate from the spinach extract? From the spirulina?
• Are all pigments represented between the two extracts?
• The mobile phase is non-polar. What are the properties of each pigment?
• Measure the R f of each pigment.

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1.8: Experiment 6 - Polarity and Solubility

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Learning Objectives

By the end of this lab, students should be able to:

• Explore the relationship between polarity and solubility.
• Determine the solubility of polar and nonpolar solutes, and an ionic solute in different solvents.

Prior knowledge:

• 8.7: Bond Polarity and Electronegativity
• 8.8: Bond and Molecular Polarity

Introduction

Have you noticed the common solubility problems in our everyday life? If you get sap from a pine tree on your clothes, or wax from a candle on the table, or bike chain grease on your pants, these substances will not be easily removed with just water. Do you think this demonstrates a solubility problem?

In this lab, you will explore how polarity affects whether substances dissolve in each other. What is the meaning of “like dissolves like”? Why are nonpolar and polar substances immiscible? You will explore what it means chemically for a substance to be polar or nonpolar and how you can use this to eventually get the stains out of your clothes.

Pre-Lab Primer

This pre-lab assignment is an individual assignment to be completed on your own with the help of the "Prior Knowledge" links at the top of this page. All work must be in your own words. Do not copy and paste information from the internet. The assignment will be due 10 minutes before your lab begins.  Late work will not be accepted.

The document below is a preview only. Please do not try to screenshot or print it off. You will be able to find your assignment to work on in your Google Classroom.

Interactive Element

In-Lab Assignments

Individual assignment.

The first assignment for this week will be an individual assignment. Your lab instructor will play a video for everyone at the beginning of lab. Based on the contents of the video, you will answer the questions in the document. After everyone has finished and turned this assignment in, your instructor will put you into your Breakout Room with your group on Zoom to complete the second assignment.

Group Assignment

The second assignment will be a worksheet that you complete with your group members. You will use the data collected from the Individual Assignment to complete the questions for this assignment.

Each person can type in this document at the same time. Remember, part of your grade comes from your participation during lab, so there will be a Peer Evaluation this week. Make sure you are contributing to discussion and to the completion of the worksheet. The worksheet will be due by the end of your lab session, and late work is not accepted. Be sure to turn your assignment in on Google Classroom.

Post-Lab Problem Set

After you have had a chance to work on the data analysis with your group during lab, you will be given the Post-Lab Problem Set. This is an individual assignment that must be completed on your own, and it is based on your Pre-Lab Primer and your In-Lab Assignments. This assignment will be due the day after your lab meets by 5 p.m. For example, if your lab is on Monday, the Post-Lab Problem Set will be due on Tuesday at 5 p.m. No late work is accepted. 

Paper Chromatography of Plant Pigments

Learning Objectives

After completing the lab, the student will be able to:

• Extract pigments from plant material.
• Separate pigments by paper chromatography.
• Measure R f (retention factor) values for pigments.

Activity 2: Pre-Assessment

• The leaves of some plants change color in fall. Green foliage appears to turn to hues of yellow and brown. Does the yellow color appear because carotenoids replace the green chlorophylls? Explain your reasoning.
• Examine the molecular structures of photosynthetic pigments in Figure 10.1. Photosynthetic pigments are hydrophobic molecules located in thylakoid membranes. Will these pigments dissolve in water?

Activity 2: Paper Chromatography of Plant Pigments

Paper chromatography is an analytical method that separates compounds based on their solubility in a solvent.

The solvent is used to separate a mixture of molecules that have been applied to filter paper. The paper, made of cellulose, represents the stationary or immobile phase. The separation mixture moves up the paper by capillary action. It is called the mobile phase. The results of chromatography are recorded in a chromatogram. Here, the chromatogram is the piece of filter paper with the separated pigment that you will examine at the end of your experiment (see Figure 10.4).

We separate the compounds based on how quickly they move across the paper. Compounds that are soluble in the solvent mixture will be more concentrated in the mobile phase and move faster up the paper. Polar compounds will bind to the cellulose in the paper and trail behind the solvent front. As a result, the different compounds will separate according to their solubility in the mixture of organic solvents we use for chromatography.

This video demonstrates the principles and examples of chromatography. You will experiment with only paper chromatography in this lab; however, you will see that you are already familiar with some uses of thin layer chromatography.

Safety Precautions

• Work under a hood or in a well-ventilated space and avoid breathing solvents.
• Do not have any open flames when working with flammable solvents.
• Wear aprons and eye protection.
• Do not pour any organic solvent down the drain.
• Dispose of solvents per local regulations.
• Use forceps to handle chromatography paper that has been immersed in solvent and wash your hands after completing this activity.

For this activity, you will need the following:

• Plant material: intact leaves of spinach and Coleus (one leaf of each plant per pair of students)
• Filter or chromatography paper
• Ruler (one per group)
• Colored pencils
• Beakers (400 mL) (Mason jars are an acceptable substitute)
• Aluminum foil
• Petroleum ether: acetone: water in a 3:1:1 proportion
• If no hood or well-ventilated place is available, the mixture can be substituted with 95 percent isopropyl alcohol. Note that, if isopropyl alcohol is used, the pigment bands will smear. You may not be able to separate and identify the chlorophylls or carotene from xanthophyll.

For this activity, you will work in pairs .

Structured Inquiry

Step 1: Hypothesize/Predict: Discuss with your lab partner what color pigments will likely be present in the spinach leaves. Write your predictions in your lab notebook and draw a diagram of how you think the pigments will separate out on the chromatography paper.

Step 2: Student-led Planning: Read step 3 below. Discuss with your lab partner the setup of the experiment. Then agree upon the dimensions of the filter/chromatography paper that you will use. To allow good separation, the paper should not touch the walls of the container. The paper must fit inside the container while being long enough for maximum separation. Write all your calculations in your lab notebook.

Step 3: Follow the steps below to set up your filter paper and perform the chromatography experiment.

• Prepare the chromatogram by cutting a piece of filter paper. Transfer pigments from spinach leaves as in Activity 1. A heavy application line will yield stronger colors when the pigments separate, making it easier to read results. Allow the pigments to dry between applications. Wet extracts diffuse on the paper and yield blurry lines.
• Form a cylinder with the filter paper without overlapping the edges (to avoid edge effects). The sample should face the outside of the cylinder. Secure the top and bottom of the cylinder with staples.
• Pour enough separation mixture to provide a mobile phase while staying below the origin line on the chromatogram. The exact volume is not critical if the origin, the start line where you applied the solvent, is above the solvent. See Figure 10.4.

• Label the beaker with a piece of tape with your initials and your partner’s initials.
• Lower the paper into the container with the band from the extraction in the lower section. The paper must touch the solvent, but not reach the band of pigment you applied. Why must the band be above the solvent line? Write your answer in your notebook.
• Cover the container tightly with a piece of aluminum foil.
• Track the rising of the solvent front. Can you see a separation of colors on the paper?
• When the solvent front is within 1 cm of the upper edge of the paper, remove the cylinder from the beaker using forceps. Trace the solvent front with a pencil before it evaporates and disappears! Draw the colored bands seen on your chromatography paper in your lab notebook immediately. The colors will fade upon drying. If no colored pencils are available, record the colors of the lines.
• Let the paper dry in a well-ventilated area before making measurements because the wet paper is fragile and may break when handled. This is also a precaution to avoid breathing fumes from the chromatogram.
• Discard solvent mixture per your instructor’s directions. Do not pour down the drain.

Step 4: Critical Analysis: Open the dried cylinder by removing the staples. Measure the distance from the first pencil line to the solvent front, as shown in Figure 10.5. This is the distance traveled by the solvent front. Measure the distance from the pencil line to the middle point of each color band and the original pencil line. Record your results in your notebook in a table modeled after Table 10.1. The retention factor (R f ) is the ratio of the distance traveled by a colored band to the distance traveled by the solvent front. Calculate R f values for each pigment using the following equation:

R f=Distance traveled by colored band/Distance traveled by solvent front

Step 5: After determining the color of the band, tentatively identify each band. Did your results support your hypothesis about the color of each band? Discuss which aspects of the experiments may have yielded inconclusive results. How could you improve the experiment?

Guided Inquiry

Step 1: Hypothesize/Predict: What type of pigments are present in Coleus leaves and where are the different colors located? Can you make a hypothesis based on the coloration of the variegated leaves? Write your hypothesis down in your lab notebook. Would there be a difference if you performed chromatography on pigment composition from different colored regions of the leaves?

Step 2: Student-led Planning: Cut the chromatography/filter paper to the dimensions needed. Apply pigments from different parts of the Coleus leaves following the procedure described under Activity 1, keeping in mind that a darker line will yield stronger colors when the pigments are separated, which will make it easier to read the results. Allow the pigments to dry between applications. Wet extracts diffuse on the paper and yield blurry lines.

Step 3: When the solvent front reaches 1 cm from the top of the filter paper, stop the procedure. Draw the pigment bands you see on the filter paper in your lab notebook. Clearly indicate the color you observed for each band.

Step 4: Let the cylinder dry and measure the distance the front traveled from the origin and the distances traveled by each of the pigments. If the bands broadened during separation, take measurements to the middle of each band.

Step 5: Critical Analysis: Calculate R f for each of the bands and record them in a table in your notebook. Compare the R f you obtained with those of other groups. Are the R f values similar? What may have altered R f values?

Assessments

• Carotenoids and chlorophylls are hydrophobic molecules that dissolve in organic solvents. Where would you find these molecules in the cell? What would happen if you ran the chromatography in this lab with water as the solvent?
• All chlorophyll molecules contain a complexed magnesium ion. Your houseplant is developing yellow leaves. What may cause this, and how can you restore your plant’s health?
• Seeds that grow under dim light are said to be etiolated, which describes their pale and spindly appearance. They soon waste away after exhausting their food reserves. Can you explain this observation?

Lab Manual for Biology Part I Copyright © 2022 by LOUIS: The Louisiana Library Network is licensed under a Creative Commons Attribution-NonCommercial-ShareAlike 4.0 International License , except where otherwise noted.

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Science in School

Colour, chlorophyll and chromatography teach article.

Author(s): Josep Tarragó-Celada, Josep M Fernández Novell

Use thin-layer chromatography to discover the variety of pigments that play a role in photosynthesis and give leaves their colour.

Looking out over a lush green valley or forest, it is fascinating to see the array of different shades. Leaves range from light to dark and even speckled. The colours are determined by the presence of different pigments, many of which are responsible for one of the most interesting and important metabolic reactions in living organisms: photosynthesis.

Photosynthetic pigments are located in the chloroplasts of the leaf. They capture energy from the visible light spectrum, which they use to synthesise carbohydrates from inorganic matter. There are many types of photosynthetic pigments, but the two main groups are chlorophylls and carotenoids (which are further split into two classes: carotenes and xanthophylls). Each type absorbs a different wavelength, so that together they capture more light.

Chlorophylls are the pigments primarily responsible for photosynthesis. They absorb red and blue light, and reflect green light, which is what gives leaves their green colour. Carotenoids, on the other hand, reflect yellow, orange and red – the colour of leaves during autumn. During this time of year, chlorophyll breaks down so the carotenoid pigments become visible.

Carotenoids assist with photosynthesis by absorbing wavelengths of light that chlorophylls cannot absorb. They transfer energy to chlorophyll molecules and also help to protect the leaf from excess light – they absorb surplus light energy and dissipate it as heat to prevent it from damaging the leaf.

Other non-photosynthetic pigments, such as anthocyanins or other flavonoids, determine the colour of flowers, so their absorption spectra vary. The function of these pigments is to attract insects or birds for pollination.

Separating leaf pigments using thin-layer chromatography

This article presents a simple laboratory experiment to understand leaf pigments. Students use thin-layer chromatography to separate the various pigments that are present in two different leaf extracts. They identify each pigment and determine whether the two extracts have any pigments in common. The experiment is suitable for students aged 11–16 and takes 1–2 hours to complete.

Note that we used leaves from Epipremnum aureum (commonly known as devil’s ivy) and Ficus benjamina (commonly known as weeping fig) , but any species could be used for the leaf extracts. You might also like to carry out the experiment using a brightly coloured flower, such as those in the Petunia genus, and also a yellow or orange leaf.

For the thin-layer chromatography, we use a combined mobile phase of hexane, acetone and trichloromethane (3:1:1) as it provides the best separation result. However, it requires part of the activity to be carried out inside a fume hood by the teacher. This mobile phase separates the pigments most clearly, but you could adapt the activity to use mobile phases of hexane or ethanol alone, which the students can carry out themselves. Both hexane and ethanol successfully separate the pigments, but the distinction between each pigment is not as clear as when the combined solvent is used.

• Leaf samples (e.g. E. aureum and F. benjamina ), cut into pieces measuring approximately 2 cm x 2 cm
• Thin-layer chromatography plates (10 cm x 5 cm) pre-coated with silica gel
• 3 parts hexane, C 6 H 14
• 1 part acetone, (CH 3 ) 2 CO
• 1 part trichloromethane, CHCl 3
• A beaker and watch glass (or chromatography chamber)
• Spotting tile
• Mortar and pestle
• 1 ml Pasteur pipettes (one for each leaf sample)

Safety note

A lab coat, gloves and eye protection should be worn. The solvents used in this experiment are flammable, so they must not be used near flames. The combined solvent (hexane, acetone and trichloromethane) must only be used inside a fume hood due to the volatility, smell and health risks associated with it.

The following steps should be carried out by the students:

• Place your first leaf sample in the mortar. Pipette 1 ml of acetone into the mortar and use the pestle to grind the sample until the leaf is broken down.
• Transfer the mixture to a well of the spotting tile using the pipette.
• Wash the mortar and pestle, and repeat steps 1–2 using the second leaf sample. Use a new pipette to add 1 ml of acetone and use this pipette to transfer the mixture to a new well of the spotting tile.
• Take the chromatography plate and draw a horizontal line 1.5 cm from the bottom using a pencil. Take care not to touch the plate with your fingers.
• Using your first pipette (take care not to mix up which pipettes were used for each leaf sample), draw up some of your first leaf sample. Apply a single, small drop to the pencil line on the left hand side of the chromatography plate. Make sure to leave enough space to fit the second sample on the right hand side.
• Wait a few seconds until it dries, and apply a second drop on the same spot. Continue until you have added around 10 drops.
• Using your second pipette, repeat steps 5 and 6 for the second leaf sample by adding it to the right hand side of the plate.
• Allow the plate to dry completely.

The following steps must be carried out by the teacher:

• Inside the fume hood, combine the solvents in the following proportions: hexane, acetone and trichloromethane, 3:1:1.
• Add the combined solvent to the beaker. You should add only a shallow layer of solvent, so that the pencil line on the chromatography plate will not be submerged.
• Place the chromatography plate vertically into the beaker, with the pencil line at the bottom, and cover the beaker with a watch glass. Students can watch as the solvent moves up the plate and the pigments separate.
• Wait until the solvent has travelled roughly 6 cm from the starting point (this will take approximately 15–30 minutes) before removing the plate from the beaker, leaving it inside the fume hood.
• Use a pencil to quickly mark the furthest point reached by the solvent. Allow the plate to dry completely before removing it from the fume hood.
• Photograph the chromatogram as soon as it is dry. The colours will fade within a few hours. Print out a copy of the photograph for your notes.
• Using the chromatogram photo, try to work out how many pigments are present in each leaf extract.
• Now look at the chemical structures of different pigments (see figure 1). Can you determine which pigment is which (see the explanation section for more guidance)? Write down your answers.
• Measure the distances travelled by the solvent and the pigments, and calculate the retardation factor (Rf) using the following equation: Rf = (distance travelled by pigment) / (distance travelled by solvent) 

Record your results in a table. Compare these to the values in table 1: were your answers correct?

Explanation

The different pigments in a leaf extract are separated based on their affinities for the stationary phase (the silica on the thin-layer chromatography plate – a polar substance) and the mobile phase (the solvent – a nonpolar substance). Compounds with a high affinity for the solvent (i.e. nonpolar compounds) will move much further than compounds with a high affinity for silica (i.e. polar compounds).

In our example (see figure 2), both leaf extracts contained four pigments. Pigment 4 moved a shorter distance than pigment 1, indicating that pigment 4 is more polar and pigment 1 is less polar. By looking at the chemical structures of different pigments and the polar and nonpolar groups, students can try to identify the pigments in each of the leaf extracts.

They will need to know that, of the functional groups present in the pigments in figure 1, alcohol groups are the most polar, ester and ether groups the least polar, and aldehyde and ketone groups are in between. From this, we can deduce that carotenes are the least polar pigments (no polar groups), and xanthophylls are the most polar (two alcohol groups, one at each end of the molecule). Therefore, pigments 1 and 2 are likely to be carotenes, and pigment 4 is likely to be a xanthophyll. Pigment 3 is likely to be chlorophyll, since it is more polar than carotenes but less polar than xanthophylls. You can observe the characteristic green colour from chlorophyll on the chromatogram.

Now look at the Rf values, which range between 0 and 1, with 0 being a pigment that does not move at all, and 1 indicating a pigment that moves the same distance as the solvent. The Rf value varies depending on the solvent used, but the general order of the pigments (from the highest to the lowest Rf value) usually remains the same, because the nonpolar compounds move further than the polar compounds. Rf values for various pigments (using hexane, acetone and trichloromethane (3:1:1) for the solvent) are shown in table 1.

After the experiment, you can ask your students some of the following questions to gauge their understanding of plant pigments and thin-layer chromatography.

• Look at absorption spectra for various plant pigments. Which pigments absorb the most light from the red end of the spectrum? What colour are they?
• If chlorophyll is the most important photosynthetic pigment, which colours of the visible spectrum are most useful to a plant for photosynthesis?
• Seaweeds are often yellow-brown in colour. Do you think light from the red end or the blue end of the spectrum penetrates water best?
• What species of plants have non-green leaves? How could you find out what pigments they contained?
• Where are photosynthetic pigments located within a leaf?
• Why is it useful for plants to contain several different photosynthetic pigments?
• Why is it important to use a nonpolar solvent (such as hexane, acetone and trichloromethane) and not a polar solvent (such as water) to investigate plant pigments using thin-layer chromatography?
• Why should you avoid touching the thin-layer chromatography plate?
• Why should the plate be completely dry before putting it into the beaker?
• Why do some pigments have a larger Rf value than others?
• Reiss C (1994) Experiments in Plant Physiology . Englewood Cliffs, NJ, USA: Prentice Hall. ISBN: 0137012853
• For an infographic explaining the chemicals behind the colour of leaves, visit the Compound Interest website .
• Read more about the chemical structure of different plant pigments by visiting the Harvard Forest website from Harvard University.

Josep Tarragó-Celada is a PhD student in biochemistry at the faculty of biology in the Universitat de Barcelona, Spain. His work focuses on the metabolic reprogramming of cancer metastasis.

Josep M Fernández Novell is a professor in the department of biochemistry and molecular biomedicine at the Universitat de Barcelona.

Together, they presented this activity at the 2018 Hands-on Science conference in Barcelona, and they frequently organise and participate in educational activities to help bridge the gap between university and secondary school students.

Combining the outdoor element of nature with the identification of different chemical structures produces a perfect applied science lesson. The analysis of the different pigments in leaves has a clear visual outcome that can then be related to the chemical structures of the different photosynthetic pigments.

This practical activity affords students the opportunity to move beyond basic paper chromatography to the more complex technique of thin-layer chromatography. This cross-curricular task will engage students who enjoy biology-based topics such as photosynthesis as well as students who enjoy the problem-solving aspect of analytical techniques in chemistry.

The activity is most suitable for students aged 14–16 as part of a science club or extension activity. In addition to the main method, the authors provide suggestions for using different solvents to enable students to carry out the experiment entirely independently. With further detail, the activity could also be useful for students aged 16–19.

Many new terms are introduced, so the article provides an excellent chance to challenge students to understand concepts such as mobile and stationary phases, polarity of molecules and how biology is fundamentally based on chemical building blocks.

Caroline Evans, head of chemistry, Wellington College, UK

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Separation of Plant Pigments by Paper Chromatography

The separation of plant pigments by paper chromatography is an analysis of pigment molecules of the given plant. Chromatography refers to colour writing . This method separates molecules based on size, density and absorption capacity.

Chromatography depends upon absorption and capillarity . The absorbent paper holds the substance by absorption. Capillarity pulls the substance up the absorbent medium at different rates.

Separated pigments show up as coloured streaks . In paper chromatography, the coloured bands separate on the absorbent paper. Chlorophylls, anthocyanins, carotenoids, and betalains are the four plant pigments.

This post discusses the steps of separating plant pigments through paper chromatography. Also, you will get to know the observation table and the calculation of the Rf value.

Content: Separation of Plant Pigments by Paper Chromatography

Paper chromatography, plant pigments, steps of plant pigment separation, observation, calculation.

It is the simplest chromatography method given by Christian Friedrich Schonbein in 1865. Paper chromatography uses filter paper with uniform porosity and high resolution.

The mixtures in compounds have different solubilities . For this reason, they get separated distinctly between the stationary and running phase.

• The mobile phase is a combination of non-polar organic solvents. The solvent runs up the stationary phase via capillary movement.
• The stationary phase is polar inorganic solvent, i.e. water. Here, the absorbent paper supports the stationary phase, i.e. water.

Plant pigments are coloured organic substances derived from plants. Pigments absorb visible radiation between 380 nm (violet) and 760 nm (red).

They give colour to stems, leaves, flowers, and fruits. Also, they regulate processes like photosynthesis, growth, and development.

Plants produce various forms of pigments. Based on origin, function and water solubility, plant pigments are grouped into:

• Chlorophylls (green)
• Carotenoids (yellow, orange-red)
• Anthocyanins (red to blue, depending on pH)
• Betalains (red or yellow)

Chlorophyll : It is a green photosynthetic pigment. Chlorophyll a and b are present within the chloroplasts of plants. Because of the phytol side chain, they are water-repelling . Their structure resembles haemoglobin. But, they contain magnesium as a central metal instead of iron.

Carotenoids : These are yellow to yellow-orange coloured pigments. Also, they are very long water-repelling pigments. Carotenoids are present within the plastids or chromoplasts of plants.

Anthocyanins : These appear as red coloured pigments in vacuoles of plant cells. Anthocyanins are water-soluble pigments. They give pink-red colour to the petals, fruits and leaves.

Betalains : These are tyrosine derived water-soluble pigments in plants. Betacyanins (red-violet) and betaxanthins (yellow-orange) are the two pigments coming in this category. They are present in vacuoles of plant cells.

You can separate all the above pigments using paper chromatography.

Video: Separation of Plant Pigments

Preparation of Concentrated Leaf Extract

• Wash spinach leaves in distilled water.
• Then take out the spinach leaves and allow the moisture to dry out.
• After that, take a scissor and cut the leaves into the mortar.
• Take a little volume of acetone into the mortar. Note : Acetone is used instead of water to mash the leaves because it is less polar than the water. This allows a high resolution of the molecules in the sample between the absorbent paper.
• Then, grind spinach leaves using a pestle until liquid paste forms. Note : The liquid in the crushed leaf paste is the pigment extract.
• After that, take out the mixture into the watch glass or Petri dish.

Load the Leaf Extract onto Absorbent Paper

• Take Whatman filter paper and draw a line above 2 cm from the bottom margin. You can use a pencil and scale to draw a fainted line. Note : A pencil is used because pencil marks are insoluble in the solvent.
• Then, cut the filter paper to make a conical edge from the line drawn towards the margin end. You can use a scissor to cut the Whatman filter paper. Note : The conical end at the bottom of the filter paper results in better separation.
• Put a drop of leaf extract on the centre of a line drawn on the absorbent paper.
• Then, at the same time dry the absorbent paper.
• Repeat the above two steps many times so that the spot becomes concentrated enough.

Setup the Chromatography Chamber

• Take a clean measuring cylinder and add rising solvent (ether acetone) up to 4 ml.
• Bend the strip of paper from the top. Then, using a pushpin attach the paper to the bottom of the cork.
• Adjust the length of the paper. The absorbent paper should not touch the surface of the measuring cylinder.
• After that, allow the solvent to move up the absorbent paper.
• When the solvent front has stopped moving, remove the paper.
• Allow it to dry for a while until the colours completely elute from the paper.
• At last, mark the front edge travelled by each pigment.

Over the dried paper strip, you will see four different bands. Different colour streaks form because of different affinities with the mobile phase (solvent).

• The carotene pigment appears at the top as a yellow-orange band.
• A yellowish band appears below the carotene, which indicates xanthophyll pigment.
• Then a dark green band represents the chlorophyll-a pigment.
• The chlorophyll-b pigment appears at the bottom as a light green band.

Observation Table

1. Light green spot indicates chlorophyll-b pigment.

• Rf value= Distance chlorophyll-b travelled / Distance solvent travelled = 2/10 = 0.2

2. Dark green spot represents chlorophyll-a pigment.

• Rf value= Distance chlorophyll-a travelled / Distance solvent travelled = 3.7/10 = 0.37

3. The yellow band represents xanthophyll pigment.

• Rf value= Distance xanthophyll travelled / Distance solvent travelled = 5.6/10 = 0.56

4. The yellow-orange band indicates carotene pigment.

• Rf value= Distance carotene travelled / Distance solvent travelled = 9/10 = 0.9

Factors affecting the Rf values of a particular analyte are:

• Stationary phase
• The concentration of the stationary phase
• Mobile phase
• The concentration of the mobile phase
• Temperature

The Rf value of compounds in the mixture differs by any changes in the concentration of stationary and mobile phases.

Temperature affects the solvent capillary movement and the analyte’s solubility in the solvent. Rf value is independent of the sample concentration. Its value is always positive .

Related Topics:

• Difference Between Budding and Grafting
• Phototropism in Plants
• Potometer Experiment

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5 Research Project: Pigments in Plant Leaves

To determine the composition of pigments in different types of plant leaves.

Expected Learning Outcomes

• Extract, separate, and characterize compounds from a mixture.
• Describe the purpose of analytical separations.
• Conduct thin-layer chromatography (TLC), column chromatography and high performance liquid chromatography (HPLC).

Reading Assignment

The focus of this experiment is on liquid phase extractions/separations.  While separations are mentioned briefly in Tro,  Chemistry – Structures and Properties , 2nd Ed, Ch. 1.2, the following readings from Nichols, L.,  Organic Chemistry Lab Techniques will help you understand this experiment:

• Chromatography: Ch. 2.2 , 2.3(e) , 2.4(b)
• Extraction: Ch. 4.2 , 4.3

In addition, we will introduce HPLC (high performance liquid chromatography.  Please read Barkovitch, M.,  High Performance Liquid Chromatography and watch the following video produced by the Royal Society of Chemistry:

About This Project

So far, most of the experiments we have done have  definite, well known answers, and we’ve been doing experiments that either (a) confirm what you’ve learnt or (b) illustrate concepts from the lecture course.  We will continue to do those experiments, not least because the combination will support your learning and help you develop laboratory skills.

However, in real life, we often need to study problems with no well-defined solution or problem.  We are dealing with unknown problems.

As you may know, there are a number of pigments in plant leaves (that make it the color they are) which are critical to photosynthesis. [1]   However, there is a mixture of them, and as leaves have different colors, how would the pigments vary – what pigments are there, and how much?  We will do this using two forms of column chromatography as explained below.

Chemical Principles

Most of the substances we find in nature (wood, air, petroleum, etc.) are not pure but are instead more or less complex mixtures.  Some of these mixtures require little or no modification before being put to practical use, but many of them must be at least partially separated into their components before they can be used to the best advantage.

• Crude oil is a mix of different hydrocarbons (compounds formed from carbon and hydrogen) that is separated into different components as shown below.
• Hard water is “softened” by separating out dissolved calcium carbonate (limescale) from water.

There are a number of chemical and physical processes used to separate mixtures but two of the most important for scientists today are the closely related techniques of extraction and chromatography.

Separation Method 1: Extraction

Extraction is the process of separating different substances by their differing solubilities in the same solvent (or solvent mixture).

When coffee is made using either a percolator or a drip-type coffee-maker, the hot water acts as a solvent to dissolve caffeine, tannic acid, and a large number of other chemical compounds, leaving the coffee grounds undissolved.  This process is an extraction because of the differing solubilities of different substances: the caffeine and tannic acid are much more soluble in hot water than other components of the coffee grounds are (cellulose, for example).

Coffee-making illustrates the main requirements for a successful extraction:

• The mixture to be separated must be thoroughly mixed with the solvent (for example, by the “percolating” action).
• Other components must not dissolve in the solvent.

For a successful extraction we must find a solvent (or solvent mixture) that does a good job of dissolving the components we want but leaves undissolved components we don’t want.  Since the solubility of most solids and liquids goes up with increasing temperature we can sometimes get the best results by using a hot solvent for the extraction.  This is true for instance for most of the substances we want in our coffee, so we always start with very hot water (even if we are planning to end up with iced coffee).  Other ways include using different solvents.

Separation Method 2: Chromatography

Chromatography is a general technique that has several points in common with extraction; for example, differing solubilities of different substances play a part in both techniques.  Chromatography differs from extraction in that this always involves the movement of one phase (the “mobile phase”) past a non-moving phase (the “stationary phase”) to bring about the separation.  Conditions are set up so that different components of the mixture are carried along with the mobile phase at different speeds as they pass through the stationary phase.

Different inks can be separated into different compounds/pigments using various forms of chromatography.

The big idea here is that there are two substances in which a substance can be dissolved:

• The  stationary phase is a substance that is fixed along the entire structure where chromatography is conducted.
• The mobile phase which is typically a liquid or gas, and can move through the  stationary phase.

Depending on the nature of the two phases and the substance concerned, there are varying ratios of solubility between the mobile phase and the stationary phase for different compounds.  When a substance is more soluble in the mobile phase, it would tend to move along with the solvent front (the point where pure solvent has flowed to).  Conversely, when a substance is more soluble in the stationary phase, then it would tend to stay on the stationary phase and not move.

For the inks on the TLC plate above, the extent to which the sample is soluble in the ethanol:water mixture relative to the silica (stationary phase, surface on the TLC plate) determines how far it migrates.  Relatively speaking, the yellow ink is more soluble in ethanol:water (relative to silica) compared to the black ink.

While TLC is useful for quickly analyzing a sample and determining what compounds are present in that sample, it is very difficult to remove compounds from a TLC plate, thus it is not generally used to purify compounds. If you want to be able to collect the components of a mixture after separating them, column chromatography is more useful.  In this technique, you place the stationary phase (for this experiment, silica gel) in a column (which is constructed using a glass, disposable Pasteur pipet).  The sample is then loaded, and mobile phase is loaded to drive the sample until it elutes out of the bottom of the column.  Each component is therefore isolated from each other and can be studied using absorption spectroscopy.  This is the main method used to purify, among other things, proteins for biochemical experiments.

High Performance Liquid Chromatography (HPLC)

In this course, we have previously described how chromatography can be used to separate different components in a mixture.  However, better separations can be made with columns created using specialized micrometer-sized particles (often silica).  As the size of the particles decreases, the separation of the column improves.

However, with such fine particles the column would not function under room pressure.  As such, high performance liquid chromatography (HPLC) instruments operate by forcing the solvent through the column under high pressure.  In each case, the difference in affinity of solutes between the mobile and stationary phases is exploited to separate the substances.

While different column resins exploit different types of separation, one of the most common approaches to separation is by polarity of the solute.  One of the most prevalent resins is that for reverse-phase HPLC (RP-HPLC), where a non-polar stationary phase is used in conjunction with a polar mobile phase.  The most common resins for reverse phase HPLC are the octyl (C8) and the octadecyl (C18) columns.

The sample will then be detected by some means (often, UV-vis absorption) and a  chromatogram is formed.  Such a chromatogram is a plot of the signal as a function of time elapsed since the sample is injected.

Absorption Spectrum

Depending on the chemical structure of the molecule, different molecules can absorb light of different wavelengths.  The color of the compound is the complementary color of the color of light absorbed – as seen in the color wheel, where the color observed is on the opposite side of the wheel:

In turn, each color of light corresponds to a different frequency/wavelength of light.  The absorbance of light at different wavelengths can be plotted in an  absorption spectrum , which can be used to help characterize a compound:

• This is a multi-week experiment.  Your notebook will be graded when you finish the HPLC part of the experiment.
• It will be conducted in pairs.
• Before conducting this experiment, please review the directions on vacuum filtration and using the SpectroVis Plus in Using Laboratory Equipment .
• As this is a research project and particularly since it’s the first time we’ve tried this, the procedure is prone to revisions and change.

Week 1: Extraction and Thin-Layer Chromatography

This week, you will

• extract the pigments using organic solvents and hot water.
• analyze the extracted pigments using TLC.

Required Special Materials and Chemicals

• Mortar and pestle
• sand (a pinch)
• Sodium sulfate (15 g)
• saturated sodium chloride solution (2 mL – use from provided bottle)
• acetone (15 mL)
• hexane (15 mL)
• TLC plates (2)
• 110 mm diameter filter paper (2)
• forceps (1)
• Observe and record the appearance and species of the leaves.  Take a photograph of the leaves.
• Weigh out 1.2 g of fresh leaves. Record the species and mass to the nearest 0.001 g, as well as a description of the leaves.
• Tear or cut the leaves into small pieces and place them in a mortar with a pinch of sand (to assist with grinding up the leaves). Add 6 mL of acetone and 6 mL of hexane and grind the mixture for 3–5 minutes in the fume hood. If little liquid is left in the mortar (<3 mL), add another 3-6 mL of each solvent and grind for another minute. You’re looking for at least 3 mL of a dark green liquid in your mortar.
• Filter the green solution using gravity filtration , into a clean 50 mL graduated cylinder then transfer it to a clean test tube. Note: do NOT filter directly into the test tube as it is likely that the test tube will break if you do.
• Add 2 mL of saturated aqueous sodium chloride solution to the test tube.
• Use a pipette and pipette the mixture up and down until it is well mixed.
• Place the test tube in the test tube rack and allow the mixture to separate.

Create a filtering pipet using a short (5″) disposable Pasteur pipet and cotton plug by tearing off a VERY SMALL (as small as possible) piece of a cotton ball, putting it into the top of the 5″ (short) Pasteur pipet and gently pushing it to the bottom of the pipet with a 2nd, longer (9″) pipet (the point of the cotton plug is just to prevent the sodium sulfate from falling out of the pipet, so don’t use too much cotton and don’t pack the cotton down too tightly). Fill the filtering pipet half-way with solid sodium sulfate using a weigh paper funnel. The final traces of water are removed by treating the organic solution with sodium sulfate as a drying agent.

• Filter the green organic layer through the filtering pipet into a screw cap vial by using a transfer pipet to move the green top layer from the test tube into the top of the filtering pipet and letting the liquid flow through the sodium sulfate and into the screw cap vial. Avoid adding any of the aqueous layer to the filtering pipet.
• Spot and run two TLC plates with your filtered green organic solution (see below for details).
• Loosely cap and label (initials, date, leaf pigment in hexane/acetone) the vial and store the vial in a small beaker (so it can’t tip over) in your drawer for next week. Be sure to NOT have the cap screwed on tightly, otherwise the solvent won’t fully evaporate. The solvent will evaporate away during this time leaving the pigment as residue. We will redissolve this residue next week.

Thin Layer Chromatography

Create the developing chambers:

• Place a 110 mm diameter filter paper into a 400 mL beaker such that it is pressed up against the side of the beaker and touching the bottom of the beaker (you will need to fold the filter paper over a bit to create a flat bottom in order to get it to fit into the beaker). You will need 2 beakers, 1 for hexane and 1 for acetone (I recommend labeling each beaker with tape so you don’t forget which beaker contains which solvent).
• Create a lid for each beaker out of aluminum foil by covering the top of the beaker with a square of aluminum foil and shaping it to the top of the beaker to seal it as tightly as possible.
• If you draw up and expel the eluent in your transfer pipet several times before transferring it to the beaker, the solvent won’t drip out of the pipet as much.
• Always hold pipets up and down (perpendicular to the floor), never sideways (parallel to the floor) as holding it sideways will cause the liquid to spill out.
• At this point, there should be a layer of solvent in that bottom of the beaker that’s ~0.5 cm thick. If it is significantly less than this, add a little more solvent.

Spot your TLC plates with your samples:

• Never touch the side of the TLC plate that has the silica gel with gloves. It can damage the silica gel and make the TLC plate unusable. Only touch the back (the plastic side) or the sides of the TLC plate.
• Never bend a TLC plate as this will cause the silica gel to flake off and make the TLC plate unusable.
• Using a NON-MECHANICAL PENCIL (the ink in pens will dissolve in organic solvents, and mechanical pencils will scratch off the silica gel), label the top of each of your TLC plates with your initials and the date.
• Label the bottom of your TLC plate with the eluent you will use to develop that plate (meaning one TLC plate should be labeled as hexane and the other one as acetone).
• Make 2 small horizontal lines 1 cm from the bottom of the TLC plate on each edge of the TLC plate (see fig. 1).
• Make 3 small (~1 mm) vertical tic marks 1 cm from the bottom of the TLC plate and ~0.5 cm away from the sides of the TLC plate and each other and label these tic marks “1”, “2”, and “3”.
• Don’t put your finger over the top opening of the capillary tube, doing so will prevent the liquid from going in and out of the tube.
• You can usually make 2-3 spots from a single dip into your organic solution, but if you’re concerned you won’t have enough solution left in your capillary tube to make your next spot (keep an eye on where the solvent level is in your capillary tube before and after making each spot), just dip the capillary tube back into your organic layer.
• After the organic solvent has evaporated from the spots on your TLC plate (when the spot is no longer as dark; this should only take a few seconds), create a 2nd spot of the same size on top of the spots in the “2” and “3” lane. This is known as double-spotting and is done if you’re concerned that adding a single spot won’t add enough of the compounds of desire to see after running.
• Add a 3rd spot to the “3” lane after the organic solvent has evaporated again. This means that you should now have 3 lanes, each with a single dot over the tic mark . The “1” lane is single-spotted, the “2” lane is double-spotted, and the “3” lane is triple-spotted. This way it is more likely that one of the lanes will have the right amount of the mixture to be able to see each compound as a distinct spot.
• Repeat this process of creating a single, double, and triple spot on the 2nd TLC plate.

Run your TLC plate:

• Place your prepared TLC plate into the appropriate developing chamber such that the bottom of the TLC plate is in the solvent and the top of the TLC plate is resting against the side of the chamber at a ~45-degree angle and immediately replace the aluminum foil lid on top of the beaker (seal as tightly as possible).
• Make sure that the TLC plate is oriented such that you can easily watch the solvent travel up the plate (places with solvent will be darker).
• Make sure the solvent level is well below where you spotted your compounds, otherwise they can dissolve into the solvent. If this happens, you will need to make a new eluent chamber and a new TLC plate.
• I’ve found that 10 minutes usually comes first.
• During these 10 minutes, you should be doing something productive, such as cleaning up, so make sure you pay close attention to each TLC plate and exactly when each plate started running so that you know when to stop each plate.
• Using a pair of forceps, grab the top of the TLC plate and remove it from the developing chamber.
• IMMEDIATELY draw a line showing where the solvent made it on the TLC plate.
• The solvent will start evaporating immediately, so the longer you wait to draw this line, the less accurate your results will be.
• The solvent might “smile”, meaning that the solvent closer to the edges ran slightly faster than the solvent in the middle. If this happens, the line you draw should reflect this.
• Once all of the solvent has evaporated from your TLC plate (i.e., it’s no longer dark anywhere), make a sketch of the TLC plate in your lab notebook (including a ruler) and take a picture of the TLC plate (including a ruler).

Week 2: Column Chromatography

• separate the pigments using microscale column chromatography.
• measure the absorption spectrum of each of the pigments.
• silica gel (about 1-2 cm 3 )

Column Chromatography

We will now separate out the different pigments in the pigment sample you extracted above, to isolate different pigments and characterize their absorption spectra.

• Once you start the elution process, it cannot be stopped. You must go to completion.  Be sure to read all directions for this part before proceeding.  Everything must be staged so that the process can occur seamlessly.
• One of the most important rules is that columns must not be allowed to go dry.  A continual flow of solvent is therefore necessary to ensure that the resin remains wet and intact.
• As glass cuvettes are fragile, be sure to handle these carefully.  These will need to be shared between all members of the class.
• The first fraction eluted is beta-carotene, and the second is chlorophyll (you may be able to separate those further, but likely not).
• If necessary, add ~0.4 mL of hexane (measure using a transfer pipet) to your vial and gently mix to reconstitute (redissolve) your pigments.
• Get a small piece of cotton or glass wool and wedge this into the narrow end of a short Pasteur pipet the same way you did last week for the filtering pipet. This should be relatively loose so liquid can get through but solid cannot.
• Scoop silica gel carefully into the column using a weigh paper funnel and make sure the column is collapsed so the powder falls to the bottom. Fill the pipet until it is about 2/3 full.
• Gently tap the side of the pipet for one minute to pack the silica.
• Add 0.5 cm of sand to the top of the pipet.
• Place the pipet column in a pipet clamp and secure it to the ring stand.
• Prepare 2 clean, small test tubes to collect the column fractions and a 100 mL beaker for waste. Label the TOP of the test tubes with your initials, the date and what fraction you will collect in that test tube (fraction 1 or fraction 2).
• Obtain 15 mL of acetone and 15 mL of hexane in separate beakers (keep these beakers covered as much as possible to minimize evaporation).
• Before elution, the column must be equilibrated (i.e. made to have similar solvent conditions as the mobile phase). To do this, place the waste beaker under the pipet column and elute 4-6 mL of hexane through the column. Note: If air bubbles or cracks appear in the column, discard and repeat Step 2.
• Note: It is important here that while there is a little mobile phase left, there isn’t too much hexane left on top of the column. If there is too much then this would dilute the pigment solution and “blur” the separation between the pigments.
• Allow the solution to adsorb onto the silica by letting it run into the column. Once the solution has adsorbed into the column (moved just below the sand) but before the silica is exposed to air, continue adding hexane continuously. Observe and record the migration of the bands that appear.  Continue collecting all clear liquids in a waste beaker.
• A yellow band will appear and begin to separate from the green band. Continue adding hexane until you finish collecting the yellow fraction in a clean test tube.
• Once you have collected the yellow fraction, change the solvent by adding acetone instead of hexane. Continue adding the solvent and collect the clear portion in the waste beaker.
• The green band should be moving down the column. Collect this fraction in a clean test tube.

Note: Not all of the pigments will be removed from the column.

• Measure the absorption spectrum of each of the fractions using a small test tube (instead of a cuvette).

Week 3/4: High Performance Liquid Chromatography (HPLC)

You will analyze the final leaf extract using HPLC.  The column we use is a C-8 column with 5 mm diameter (4.6 mm internal diameter x 100 mm length).  The solvents used are:

• Solvent A: 80:20 methanol:water
• Solvent B: 95:10 acetonitrile:water

These solvents are a mixture of methanol (CH 3 OH) and water or acetonitrile (CH 3 CN) and water.  A uniform solvent is used for elution.   We will measure the absorbance of the sample at 440 nm and use this to generate the chromatograph.

The method is as follows with a flow rate of 1 mL/min

• 0 min-5 min: linear gradient from 100% solvent A to 100% solvent B
• 5 min-20 min: 100% solvent B

Your instructor will provide more information about the method closer to the time of the experiment.

• Your instructor will do this step: Load the method directed by your instructor and allow the system to equilibrate.
• The system needs to be equilibrated so there’s nothing in the column before you start another round.  The chromatogram should be blank for a couple of minutes before you start.
• Look around and make sure there are no leaks in the system.
• Clean the syringe by drawing up 50:50 hexane:acetone mixture into the syringe and ejecting it into waste.
• Make sure the manual injector is in the upward direction.
• Get the sample vial from last week and redissolve in a small amount of 50:50 hexane:acetone mixture.
• Gently draw up a little more than 20 μL of the liquid with the special syringe, making sure there are no air bubbles. If there are air bubbles, eject all the liquid and try again more slowly, making sure that the tip of the needle is always submerged in the liquid. Inject the sample into the injector.
• When the injection is completed, push the injector knob downwards so that the sample is injected. Make sure no air bubbles are injected it into the system.  The HPLC system will record the starting time.
• Wait until the method is complete.  Make a PDF of the report and email the PDF to yourself.

Sharing of Data

You will post the PDF along with information about the leaves (species and overall appearance) onto Canvas by midnight the day you complete your lab report.

Data Analysis

• Locate spectra of different plant pigments and compare against the spectra for the pigments you extracted in the first week.  Could you identify these pigments?
• For the HPLC experiment, refer to Wright, S. W. et al, Improved HPLC method for the analysis of chlorophylls and carotenoids from marine phytoplankton .   Mar. Ecol. Prog. Ser. 1991,  77 , 183-196 and compare the peaks observed to yours.  Note that since the method is slightly different I would not expect your spectra to look exactly the same as the spectra in this paper.

Acknowledgment:

The extraction and column chromatography procedure is based on a protocol used previously for extracting pigments from spinach leaves for organic chemistry.

• For further details, please read Hoefnagels, Biology: The Essentials , 4th Ed, Ch. 5.2 (Photosynthetic Pigments) or OpenStax Biology 2e , Ch. 8.2 . ↵

IU East Experimental Chemistry Laboratory Manual Copyright © 2022 by Yu Kay Law is licensed under a Creative Commons Attribution-NonCommercial 4.0 International License , except where otherwise noted.

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