Photosynthesis Lab Combinepdf (4) PDF

LABORATORY 6 Photosynthesis Objectives After completing this lab, you should be able to: Explain the process of photosynthesis as expressed by the Hill reaction Measure photosynthetic activity using a spectrophotometer Conduct an experiment to measur...

LABORATORY 6 Photosynthesis Objectives After completing this lab, you should be able to: Explain the process of photosynthesis as expressed by the Hill reaction Measure photosynthetic activity using a spectrophotometer Conduct an experiment to measure the Hill reaction in isolated chloroplasts Analyze and interpret data obtained in the experiment Readings Fenton et al., (2023) Section 6.2 (Photosynthesis: An Overview), pages 138 – 140 Introduction Photosynthesis is the process by which green plants, algae, and certain bacteria convert light energy and CO2 into organic molecules. In green plants, algae, and cyanobacteria, the overall process of photosynthesis can be summarized by the equation: 6 CO2 + 12 H2O C6H12O6 + 6 O2 + 6 H2O While this equation is chemically correct, the actual process of photosynthesis is very complex and involves many separate reactions, which can be grouped into two categories: the light-dependent reactions and the light-independent reactions (Calvin Cycle). In the light-dependent reactions, the energy of sunlight is trapped and converted to ATP and NADPH by the combined activities of an electron transport chain (ETC) and chemiosmosis. The products of the light-dependent reactions are used to drive the Calvin Cycle, in which CO2 is reduced to carbohydrate. Thus, while the light-independent reactions do not directly require light, they are dependent on the products of the light-dependent reactions. A diagram of the key events of the light-dependent reactions is shown in Figure 6.1. Light absorbed by photosynthetic pigments in the antennae complexes funnel energy to the reaction centre chlorophyll a of Photosystems I (PSI) and II (PSII). Electrons in the reaction centre chlorophylls are boosted to a higher energy level and then passed to the primary electron acceptor associated with the photosystem. 1 In PSII, excited electrons first flow through protein complexes towards PSI. The energy released by the ETC is used to transfer protons from the stroma to the thylakoid space. This generates an electrochemical gradient (Proton Motive Force) across the thylakoid membrane that powers ATP synthesis via chemiosmosis. After donating electrons to the ETC, the reaction centre chlorophyll a in PSII become very strong oxidizing agents and can strip electrons from water to replace the ones they have lost. The splitting of water into protons and oxygen in the presence of light is called photolysis (“light splitting”). Light energy is funnelled to the reaction centre of PSI in the same way as it is in PSII. Absorbed energy causes the excitation of electrons in the reaction centre chlorophyll a which are then transferred to a separate ETC. In linear electron transport, the electrons are used to reduce NADP+, which is reduced to NADPH, the source of reduction potential for the Calvin cycle. Electrons lost by the reaction-centre chlorophyll of PSI are replaced by electrons donated by plastocyanin. Figure 6.1 Schematic diagram of the light-dependent reactions THE HILL REACTION From Figure 6.1, you can see that the oxygen released by plants is derived from water, not carbon dioxide. However, for many years, it was thought that CO2 was the source of oxygen, and many people still hold this misconception today. How did we learn that the oxygen comes from water? In 1937, Robert Hill discovered that isolated chloroplasts produce oxygen when they are exposed to light and provided with an electron acceptor, even if no CO2 is present. The Hill reaction is formally defined as the reduction of any electron acceptor (“A” below) by electrons and protons from water, with the production of oxygen, when chloroplasts are exposed to light: ! H2O + A AH2 + " O2 2 You will study how different concentrations of an inhibitor influences the light reactions of photosynthesis by monitoring the Hill reaction in chloroplasts isolated from plant leaves. To measure the progress of the Hill reaction, you will use an artificial electron acceptor, the pigment DCPIP, which is a stronger oxidizing agent than NADP+, thus replacing NADP+ as the final electron acceptor in the light-dependent reactions. DCPIP is blue when oxidized and becomes colourless when reduced (Figure 6.2). As the Hill reaction proceeds, the reduction of DCPIP can be measured as a change in light absorbance at 620 nm. A standard curve can be used to determine the concentration of DCPIPoxidized produced over time. Figure 6.2 Redox states of DCPIP What is the experimental question you are investigating? What is an appropriate null hypothesis? What is an appropriate alternate hypothesis? What is the independent variable in this experiment? What is the dependent variable? What is an appropriate control for this experiment? Is there more than one? What factors will you want to keep constant? 3 ISOLATING CHLOROPLASTS FROM PLANT LEAVES The course technicians have isolated chloroplasts from fresh spinach (Spinacia oleracea) by grinding the leaves followed by centrifugation to separate the chloroplasts from other cellular components. The isolated chloroplasts were suspended in cold sodium-potassium phosphate buffer (pH 7.3). Working in Pairs Collect a tube of chloroplast solution from the back bench. Wrap tube in aluminum foil and keep it on ice. Why is it important to wrap the tube in foil? Experiment I: Measuring the Hill Reaction 1. Label 3 test tubes “1” through “3”. Wrap Tube #2 in aluminum foil and place all 3 tubes in the test tube rack on your bench. Label 3 cuvettes “1” through “3” (on the frosted area) at the top of each cuvette. Wrap Cuvette #2 in aluminum foil and place all 3 cuvettes in a cuvette rack. 2. Set up the light source and beaker of water as illustrated in Figure 6.3. The beaker will act as a heat shield, protecting the chloroplasts from the heat produced by the light. Do not turn the lamp on until Step 9! Figure 6.3 Arrangement of light source, heat sink, and cuvettes 3. Ensure that the spectrophotometer is on and set to 620 nm. 4. Add 3 mL buffer and 1 mL RO water to Tube #1; do not add DCPIP. What is the purpose of Tube #1? 5. Add 3 mL buffer and 1 mL DCPIP to both Tube #2 and Tube #3. What is the purpose of Tube #2? What is the purpose of Tube #3? 4  Perform Steps 6-8 as quickly as possible to improve the accuracy of your measurements 6. Prepare the spectrophotometer cuvettes. a. Invert the chloroplast solution several times to ensure the chloroplasts are re-suspended. Transfer 1 mL into each of Tube #1, Tube #2, and Tube #3. Return the chloroplast solution to your ice bucket. b. Use Parafilm to seal each of Tube #1, Tube #2, and Tube #3. Invert to mix. c. Pour an aliquot from Tube #1 into Cuvette #1, from Tube #2 into Cuvette #2, and from Tube #3 into Cuvette #3. d. Place Cuvette #1 near the spectrophotometer so you can use it as a blank throughout the assay. e. Measure the initial absorbance of Cuvette #2. Record the absorbance in Table 6.1 on page 7. Keep Cuvette #2 wrapped in foil and place it in a cuvette rack. f. Measure the initial absorbance of Cuvette #3. Record the absorbance in Table 6.1. Place Cuvette #3 in a cuvette rack 20 cm away from the heat sink (Figure 6.3). Turn on the lamp and start timing. 7. At 2 minutes, stop the timer and turn off the lamp. Quickly invert Cuvette #3 and measure the absorbance – don’t forget to seal the cuvette with parafilm and to blank the spectrophotometer with Cuvette #1. Return Cuvette #3 to the cuvette rack, turn on the light, and restart the timer. 8. Measure the absorbance of Cuvette #3 every 2 minutes for 14 minutes. Remember to stop the timer, turn off the light, invert the cuvette, and blank the spectrophotometer at each reading. 9. At 14 minutes, unwrap Cuvette #2, seal with parafilm, invert, and measure the final absorbance. 10. Turn off the lamp, rinse the tubes and cuvettes, and leave them in the test tube rack upside down to dry. 11. Transfer your data from Table 6.1 to the class data sheet. 5 Experiment II: Effect of an Inhibitor on the Hill Reaction DCMU is a chemical used in many herbicides. In Experiment II, you will investigate the effect of various concentrations of this inhibitor of photosynthesis on the Hill reaction. 1. A stock solution of DCMU (labelled “Stock DCMU”) has been provided. Prepare a 1:10 and a 1:100 dilution of the inhibitor stock solution using RO water. a. Label a test tube “1:10 DCMU” and another “1:100 DCMU”. b. Add 1.8 mL RO water to both tubes c. Aliquot 0.2 mL DCMU stock to the 1:10 tube and vortex to mix d. Aliquot 0.2 mL from the 1:10 tube to the 1:100 tube and vortex to mix 2. Label 4 test tubes “1” through “4” and place all 4 tubes in the test tube rack at your bench. a. Add 2 mL buffer and 2 mL RO water to Tube #1 (do not add DCPIP) and vortex to mix b. Add 2 mL buffer, 1 mL DCPIP, and 1 mL Stock DCMU to Tube #2 and vortex to mix c. Add 2 mL buffer, 1 mL DCPIP, and 1 mL 1:10 DCMU to Tube #3 and vortex to mix d. Add 2 mL buffer, 1 mL DCPIP, and 1 mL 1:100 DCMU to Tube #4 and vortex to mix Perform Steps 3–5 as quickly as possible to improve the accuracy of your measurements 3. Label 4 cuvettes “1” through “4” (on the frosted area) at the top of each cuvette and place all 4 cuvettes in a cuvette rack. a. Invert the chloroplast solution several times to ensure the chloroplasts are re-suspended. Transfer 1 mL into each of Tube #1, Tube #2, Tube #3, and Tube #4. Return the chloroplast solution to the ice bucket. b. Use Parafilm to seal each of Tube #1, Tube #2, Tube #3, and Tube #4. Invert to mix. c. Pour an aliquot from Tube #1 into Cuvette #1, from Tube #2 into Cuvette #2, from Tube #3 into Cuvette #3, and from Tube #4 into Cuvette #4. d. Place Cuvette #1 near the spectrophotometer so you can use it as a blank throughout the assay. e. Measure the initial absorbance of Cuvette #2. Record the absorbance in Table 6.2 on page 7. Place Cuvette #2 in a cuvette rack 20 cm away from the heat sink (Figure 6.3). f. Repeat Step 3e for both Cuvette #3 and Cuvette #4. g. Turn on the lamp and start timing. 6 4. At 2 minutes, stop the timer and turn off the lamp. a. Quickly invert Cuvette #2 and measure the absorbance – don’t forget to seal the cuvette with parafilm and to blank the spectrophotometer with Cuvette #1. Return Cuvette #2 to the cuvette rack. b. Repeat Step 5a for both Cuvette #3 and Cuvette #4. c. Turn the light on and restart the timer. 5. Measure the absorbance of all three cuvettes every 2 minutes for 14 minutes. Remember to stop the timer, turn off the light, invert the cuvettes, and blank the spectrophotometer at each reading. 6. After the last measurement, turn off the lamp, rinse the tubes and cuvettes, and leave them in the test tube rack upside down to dry. 7. Transfer your data from Table 6.2 to the class data sheet. Table 6.1 Absorbance readings for Experiment I Time (min) Cuvette 0 2 4 6 8 10 12 14 #2 ¾ ¾ ¾ ¾ ¾ ¾ #3 Table 6.2 Absorbance readings for Experiment II Time (min) Cuvette 0 2 4 6 8 10 12 14 #2 #3 #4 7  LABORATORY 5 Fermentation and Biofuels Objectives After completing this lab, you should be able to: Explain the process of fermentation to produce ethanol. Carry out an experiment to compare the biological efficiency of the fermentation process Analyze and interpret data obtained in your experiment. Readings Bourne JK. 2007. Biofuels: boon or boondoggle? Green dreams. National Geographic October 2007 [Internet]. [Cited 2011 May 4]. Available from: http://ngm.nationalgeographic.com/2007/10/biofuels/biofuels-text Potters G, Van Goethem D, Schutte, F. 2010. Promising Biofuel Resources: Lignocellulose and Algae. Nature Education. 3(9):14. Fenton et al. (2023) Section 5.7a (In Eukaryotic Cells, Low Oxygen Levels Result in Fermentation) Background A. Economic and Environmental Impacts of Ethanol Biofuel Production from Plants Biofuels, such as ethanol made from corn or biodiesel made from soybeans, have attracted a lot of interest as alternative sources of energy that are touted as being more sustainable and “environmentally friendly” energy sources. Many countries around the world have been pouring money into biofuel production. In 2007, for example, the Canadian government committed $1.5 billion to encourage greater production of biofuels. More recently, investments have been made into the construction of biodiesel refineries in Canada, with the first being completed in Prince George, B.C. in the summer of 2023. As of December 2010, all gasoline sold in Canada must be blended with at least 5% ethanol. In the U.S., the government announced in 2007 the goal of reducing reliance on foreign oil supplies by increasing the annual production of corn ethanol through the Renewable Fuel Standard Program. Over a third of all corn grown annually in the U.S. has been devoted to the production of fuel ethanol since 2008. Ethanol production in the U.S. has increased from 13.6 billion gallons per year in 2011 to 17.5 billion gallons in 2021. Several concerns about using corn ethanol as fuel have been raised by scientists, environmentalists, and legislators (Bourne 2007; Potters et al. 2010). One concern is the relatively low yield of corn in terms of amount of ethanol that is produced from a hectare of the crop. For example, if the US, the leading producer of corn ethanol, dedicated all the land on which corn is grown to producing corn for ethanol, it would still only produce enough ethanol to replace 12% of the country’s total gasoline 1 demand (Bourne 2007). If the energy costs necessary to produce this much ethanol are considered, there would be a net energy gain of only 2.4% of the total American energy market. The environmental costs of producing ethanol from corn also include increased eutrophication and pollution of groundwater and aquatic ecosystems by the herbicides and fertilizers needed for growing corn. Nitrogen fertilizers are also converted to N2O by soil microbes and thereby contribute to the production and release of greenhouse gases. Thus, even if corn ethanol is a “carbon-neutral” fuel in terms of CO2 emissions (but see Potters et al. 2010 for discussion of this point), its production still contributes to greenhouse gases. While corn is the primary biofuel crop in North America, there are other plants used for ethanol production. How do they compare in terms of economic and environmental impacts? Sugarcane At first glance, sugarcane-based ethanol production has considerable advantages over corn in terms of greenhouse gas emissions. Not only does land devoted to sugarcane produce much more ethanol than land devoted to corn, but in Brazil, the world leader in sugarcane-based ethanol production, ethanol refineries derive most of their energy from burning sugarcane residue. They thus avoid fossil fuel burning and further cut the greenhouse gas emissions associated with the production process. Some estimates claim that sugarcane ethanol produced on established plantations offers an 80% reduction in greenhouse gas emissions compared to traditional gasoline. Brazil has successfully translated a massive investment into the process of independence from oil imports. Unfortunately, there is a serious problem with Brazilian sugarcane-based ethanol production: much of it is done on newly cleared lands from the country’s rainforests. In the global carbon cycle, plants and soil contain three times more carbon than exists in the atmosphere. When Brazilian rainforests are cleared to make room for sugarcane production, about 25% of the carbon dioxide previously stored in the forests’ trees and plants is released into the atmosphere due to cutting and burning of trees. Even more carbon dioxide is released in the first 20–50 years of farming former rainforest lands, as tree roots decay and other organic matter in the soil decomposes. When greenhouse gas emissions related to the clearing of tropical rainforests are considered, almost 50% more greenhouse gases are emitted than when a comparable amount of traditional gasoline is produced and burned (Bourne, 2007). In addition, the clearing of Brazil’s rainforest to produce more sugarcane for the ethanol industry will have a serious impact on biodiversity in the country, as countless habitats are destroyed. Furthermore, both sugarcane- and corn-based production of ethanol has the potential to drive food prices up and decrease global food security. Corn is a leading source of animal feed in Canada (and the United States), representing a key input for the dairy, poultry, and beef industries. As demand for corn to supply the rising number of ethanol refineries soars, so do corn prices, resulting in higher prices for consumers on a wide range of food products. A dramatic illustration of this food versus energy struggle occurred in 2007 in Mexico, when thousands of people took to the streets of the capital city to protest the rising prices for corn tortillas, a food staple of the country’s poor. 2 Wheat The biofuel industry in Canada is much smaller than in the U.S. or Brazil, but there are several plants across the country and more in development. While the focus in eastern Canada is on corn, the main crop of interest in western Canada is wheat. The grain of wheat is rich in starch, and it has been suggested that genetic engineering could further boost starch content, making wheat an even better fuel. As for corn and sugarcane, ethanol production from the grain of wheat means using a food crop for fuel. However, a different approach to producing biofuel would be to convert the cellulose in vegetative plant parts (e.g., stems) to ethanol, rather than just using the parts high in sugars or starch, which would eliminate the competition for food crops. Some biofuel producers are investigating the use of wheat stems as a source of cellulose biofuel. How do these different plants and different plant parts compare in terms of serving as the starting point for the biological process by which ethanol is produced? Is one best, in terms of the efficiency of the basic process that produces ethanol? B. The Biological Process of Ethanol Production from Fermentation of Plant Material Bioethanol is produced when carbohydrates are fermented by yeast. “Yeast” is a general term for a type of fungus that grows as a single cell and reproduces by budding. While there are many different species of yeast, the one most familiar to us is Saccharomyces cerevisiae, which has been used by humans throughout history to produce ethanol by fermentation. Although ethanol has the obvious ability to intoxicate, in the past alcoholic beverages were widely consumed because they provided nutritious (carbohydrate-rich and sometimes protein-rich), and safe (free of pathogenic bacteria) beverages during times, such as the Middle Ages in Europe, when food was not often both safe and available. Some alcoholic beverages, such as certain types of beer, use different species of Saccharomyces in their production. Humans also rely on fermentation by S. cerevisiae to make bread, although the strains used in baking differ from those used in alcohol production in terms of the relative amounts of ethanol and CO2 produced. The carbon dioxide produced in fermentation causes the bread to rise; the ethanol produced evaporates during baking. In more recent times, the process of fermentation by yeast has been co-opted to produce ethanol fuel. Henry Ford’s first car ran on pure ethanol (Bourne, 2007); modern ethanol fuel is 200 proof (100% concentration) and contains an additive that allows producers to avoid paying the ethanol-as-beverage tax. For food or fuel, the starting point in the ethanol production process is to provide the yeast with a source of carbohydrates (e.g., corn, sugarcane, grapes, barley, etc.) that it uses as a source of both energy and carbon. The chemical equation for the fermentation process is: C6H12O6 + H2O 2 C2H5OH + 2 CO2 + H2O Glucose ethanol 3 The most important part of this process for the yeast is not depicted in the above equation; the yeast breaks down the carbohydrates to release energy that it can use for its metabolic processes during this reaction, as shown in Figure 5.1: Figure 5.1 Ethanol fermentation Carbohydrates other than glucose can be used but they must first be broken down to simple sugars (like glucose) that can be fermented. Consider the plant materials available in this lab (sugarcane, corn, and barley) as substrates for fermentation: Which one(s) do you think contain a greater proportion of simple carbohydrates (sugars)? Which one(s) contain a greater proportion of complex carbohydrates (such as starch)? Which one(s) contain a greater proportion of structural carbohydrates (such as cellulose)? The ethanol produced in fermentation becomes toxic to the yeast cells once the alcohol concentration reaches a certain level; this is why alcoholic beverages produced solely from fermentation (e.g., beer, wine) don’t reach alcohol concentrations exceeding ~18% without supplementation. Therefore, the fermentation of raw materials, such as sugarcane or corn must be followed by distillation to remove excess water in the mixture and reach the desired 200 proof (100%) concentration. Notice that carbon dioxide (CO2) is another by-product of the fermentation process. Does this CO2 production present any problems for the argument that ethanol is a green (environmentally friendly) fuel? Why or why not? 4 Procedure for measuring ethanol produced in fermentation Before you conduct your experiment, read through the procedure below and complete the questions throughout, which will be a valuable resource as you prepare for the Lab Exam. Your task is to carry out an experiment comparing the fermentation of the three plant materials to determine if different plant materials produce different amounts of ethanol (biofuel). What is the experimental question you are investigating? What is an appropriate null hypothesis? What is an appropriate alternate hypothesis? You will quantify the amount of ethanol produced over time by measuring the volume of carbon dioxide released during the fermentation of each plant material. Why can carbon dioxide production be used as a measure of fermentation/ethanol production? (Review Figure 5.1) What is the independent variable in this experiment? What is the dependent variable? Each experimental treatments will consist of yeast solution mixed with one of the plant materials. You will run three replicates of each experimental treatment assigned to you and three replicates of a control treatment. What is an appropriate control for this experiment? What factors will you want to keep constant? Procedure – work in a group of 4 Two people from the group will prepare the yeast solution (step 1) while the other two people collect the necessary equipment and plant material (step 2). 1. The yeast solution consists of 5%(w/v) yeast and 0.14%(w/v) NaCl in warm water. Weight/volume percentage concentration (w/v%) is one way to indicate the concentration of a solute in a solution. w/v% can be determined by dividing the mass of the solute, usually in grams, by the volume of solvent, usually in millilitres, and multiplying by 100%. The units for w/v% are g/100mL and tells us how many parts of solute are present in 100 parts of solvent. A 5%(w/v) yeast solution contains 5 g yeast in 100 mL water. How many grams of NaCl are in 100 mL of yeast solution? 5  To have enough yeast solution to complete three replicates of the four treatments (3 plant materials + the control), your group will make 500 mL of yeast solution. What mass of yeast will you require? What mass of NaCl will you require? a. Collect the yeast and NaCl from the side bench. Use a separate balance and weigh boat for each. b. Collect a flask of water from the 40°C water bath on the side bench. c. Use the graduated cylinder to pour 250 mL water into the Erlenmeyer flask at your bench. Add the NaCl and yeast to the flask, cover with Parafilm, and swirl to mix. d. Add the another 250 mL water to the flask, and again cover with Parafilm, and swirl to mix e. Place the flask in the water bath at your bench 2. Your group will need one beaker and one fermentation tube set for each replicate (12 beakers and 12 fermentation tube sets)! Label the beakers and large fermentation tubes with the type of plant material and replicate number (you do not have to label the small fermentation tubes). You will label one beaker and one large fermentation tube: “sugarcane replicate 1” You will label another beaker and large fermentation tube: “sugarcane replicate 2”, etc. a. Weigh 2 g of each plant material into the appropriate beaker 3. Each fermentation tube set consists of a small test tube, containing a mixture of yeast solution and plant material, inverted inside a large test tube, as shown in Figure 5.2. Figure 5.2 A fermentation tube set 6 4. Stagger the time you start each set of fermentation tube set so that you are not trying to read each tube at the same time. As soon as you mix the yeast solution and plant material, the reaction starts. For each of the fermentation tube sets: a. Thoroughly stir the yeast solution and transfer a 30 mL aliquot into one of the beakers. Use a spatula to mix the yeast solution into the plant material. b. Pour the yeast-plant material mixture into one of the small fermentation tubes until it is completely full. c. Holding the small tube upright, slide the large fermentation tube (labelled with the correct plant material and replicate number) over top of the small tube and push the small tube all the way up inside the large tube with your finger (see Figure 5.3). d. Carefully invert the tubes such that the large tube is now upright, with the small tube upside down inside the large tube (Figure 5.3). This is time = 0 for this replicate. e. Some of the yeast-plant material mixture may overflow into the large tube. In Table 5.1, record the initial height (in mm) of fluid in the large tube (time = 0 minutes) by placing a ruler next to the tube. You will subtract this value from all later height readings to correct for this overflow. f. Place the fermentation tube set in a test tube rack to keep it stable. Figure 5.3 Preparation of one set of fermentation tubes 7 5. As fermentation proceeds, the CO2 gas produced will form a bubble at the “top” of the small tube. This will force some of the liquid in the small tube into the large tube, and thus the volume of liquid in the large tube will increase (see Figure 5.4). a. Every 5 minutes for 20 minutes, record the height (in mm) of fluid in the large tube in Table 5.1. If the small tube starts to float upwards inside the large tube, push it down with your finger before you read the fluid height. If the plant material sticks to the top of the small tube, push the small tube down with a finger and gently tap the bottom of the large tube against the palm of your hand (not the bench). Figure 5.4 As fermentation proceeds, a CO2 gas bubble forces liquid into the large tube 6. Clean all your tubes using the test tube brushes at the back sink. Rinse the tubes and beakers with Reverse Osmosis (RO) water and place them upside down at your bench to dry. Data Conversions 1. You need to correct for any overflow that occurred when you inverted the fermentation tube set for each replicate (see step 4e). Subtract the initial height of fluid in the large tube from the fluid height recorded at teach time point. Record the Corrected Fluid Height in Table 5.1. 2. Convert the fluid height at each time to volume of carbon dioxide using the conversion factor below and record the values in Table 5.1. 7 mm fluid in outer test tube = 1 mL CO2 3. Calculate the mean volume of carbon dioxide produced at each time for each treatment (3 plant materials + the control) and record the values in Table 5.1. 4. Record the mean volume of carbon dioxide produced at each time point for each treatment on the lab data sheet. 8  Table 5.1 Mean Volume of CO2 Produced by Fermentation of Various Plant Materials Over Time by Saccharomyces cerevisiae 9  LABORATORY 4 Eutrophication Procedures Each group of 4 students will set up one microcosm (see Figure 4.1). Your group will be assigned to 1 of 6 different treatments (see Table 4.1); The fertilizer that you will be using contains 15% nitrogen by weight, 30% phosphorous, 15% potassium, and trace amounts of other elements required for plant growth. Figure 4.1 Microcosm parts and assembly. This material has been adapted with permission. Christopher Harendza, Ph.D., Eutrophication Lab Exercise, presented at the 2008 ABLE Conference at the University of Toronto, Mississauga Table 2.1 Microcosm treatments. All microcosms are given 100 mL of distilled water three times per week for three weeks (total volume of water/fertilizer provided = 900 mL). The water in microcosms 2–6 is replaced with 100 mL of fertilizer solution at specific intervals as indicated below. Calculate the total amount of fertilizer added in three weeks and enter this value in the right-hand column to complete the table. Fertilizer Total Volume of Total amount of Fertilizer Microcosm concentration Watering/Fertilization Fertilizer added added Over Three Weeks # Frequency (g L–1) Over Three Weeks (g) 1 Distilled water All waterings with distilled water only 0 Water replaced with 100 mL of fertilizer 2 0.78 200 mL (0.2 L) solution once per 14 days Water replaced with 100 mL of fertilizer 3 1.56 200 mL (0.2 L) solution once per 14 days Water replaced with 100 mL of fertilizer 4 3.12 200 mL (0.2 L) solution once per 14 days Water replaced with 100 mL of fertilizer 5 3.12 300 mL (0.3 L) solution once per week Water replaced with 100 mL of fertilizer 6 3.12 900 mL (0.9 L) solution three times per week 2 A. CONSTRUCTING THE LOWER CHAMBER OF THE MICROCOSM 1. Place 4 cm of fine gravel in the bottom of the lower chamber of your microcosm. 2. Measure 500 mL of distilled water and add it to the bottom chamber of your microcosm. This will be the “pond” in your microcosm. 3. Choose four aquatic plants. Gently blot them dry with paper towel and determine the total mass of the plants; record this value in Table 7.1 on the class data spreadsheet. Do this step quickly—do not allow the plants to dry out completely! 4. Anchor the plants in the gravel, spacing them evenly around the bottom. 5. Add an additional 250 mL of distilled water to the pond. B. ADDING MICROBES TO THE LOWER MICROCOSM CHAMBER 1. On the side bench, there are cultures of phytoplankton and zooplankton. These organisms are representative of the algae and protozoa found in many natural ponds. Gently swirl each culture flask to mix it and, using a new sterile pipette tip for each culture, add 500 µL of each culture to the “pond” in your microcosm. 2. Once you have inoculated the “pond” with the phytoplankton and zooplankton, gently mix the pond with a stir stick to evenly mix contents (be careful not to disturb the aquatic plants while you mix). C. CONSTRUCTING THE UPPER CHAMBER OF THE MICROCOSM 1. To set up the upper or “field” chamber of your microcosm, first place 3 or 4 pieces of large gravel in the bottle cap at the base of the funnel- shaped top chamber. 2. Fill the top container with the soil mixture until the mixture is 3 cm from the top of the container. Do NOT compact this soil too much: tap the container on the bench to settle the soil as you add it and when the soil surface is 3 cm below the top of the container, lightly press down on the soil so there are no large air holes in the soil. 3. Place 10 terrestrial plant seeds on top of the soil. Cover the seeds with a very thin (3–5 mm) layer of vermiculite. Mist the soil with distilled water to ensure that it is totally moist. 4. Finish the assembly of your microcosm by placing the middle connecting section into the top of the bottom “pond” chamber, as shown in Figure 4.1, then placing the top “field” chamber into the middle connection section. Tape around the seams between the middle and bottom sections, and between the top and middle sections, with labeling tape. Label the tape on your microcosm with your lab section, group name, and treatment. 5. Carefully place your microcosm into the designated space on the light cart. Always pick up and carry your microcosm by the base! Note: The course technicians will look after watering/ fertilizing of all microcosms for the duration of the experiment. 3 D. QUANTIFYING ZOOPLANKTON ADDED TO THE “POND” To determine the effect of eutrophication on zooplankton, you will need to know how the density of the population in each microcosm was affected, i.e., did the population increase or decrease in size? This means that you need to know the density of plankton (# cells per mL of “pond” water) at the start of the experiment. One pair of students in your group should carry out Steps 1 and 2; then each pair of students should carry out Steps 3–6 below: 1. Gently swirl the culture flask of zooplankton to mix it. Using a new sterile pipette tip, add 500 µL of the culture to a 1.5-mL microcentrifuge tube. 2. Add 500 µL of Lugol’s iodine to the microcentrifuge tube, close the lid, and invert tube several times to mix the contents. Lugol’s iodine will kill the cells, enabling you to count the number of cells in each volume of liquid accurately. 3. Obtain a Palmer-Maloney counting slide, which contains a reservoir holding 100 µL of sample (Figure 4.2). Carefully place the coverslip over the reservoir and add a 0.1-mL (100-µL) aliquot of the mixture in the microcentrifuge tube into one of the side ports on the slide. Figure 4.2 Palmer-Maloney counting slide. 4. Work with a partner to use a light microscope to count the number of cells as demonstrated by your TA. Use the 10X objective lens and sequentially select five fields of view, moving from left to right across the slide. One person should count the number of cells visible in each field of view, while the other person records these values and keeps track of the number of fields of view that you examine. Then calculate the final cell density of zooplankton per mL of “pond” water using the following equation: (𝑣𝑜𝑙𝑢𝑚𝑒 𝑜𝑓 𝑐𝑢𝑙𝑡𝑢𝑟𝑒 𝑎𝑑𝑑𝑒𝑑) 𝑛𝑢𝑚 𝑜𝑓 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑠𝑙𝑖𝑑𝑒 = 𝑐𝑒𝑙𝑙𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 × , 𝑎𝑟𝑒𝑎 𝑜𝑓 𝑓𝑖𝑒𝑙𝑑 𝑜𝑓 𝑣𝑖𝑒𝑤 𝑎𝑡 10𝑋 𝑝𝑜𝑤𝑒𝑟 × 𝑑𝑒𝑝𝑡ℎ 𝑜𝑓 𝑠𝑙𝑖𝑑𝑒 𝑟𝑒𝑠𝑒𝑟𝑣𝑜𝑖𝑟 = 4 < × 𝑛𝑢𝑚 𝑜𝑓 𝑓𝑖𝑒𝑙𝑑𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 100 𝑚𝑚! 𝑛𝑢𝑚 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑠𝑙𝑖𝑑𝑒 = # 𝑐𝑒𝑙𝑙𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 ∗ 0.02 𝑚𝑚" 𝑋 0.4 𝑚𝑚 𝑋 5 4 5. The # cells/slide is the # cells present in 0.1 mL of solution from the microcentrifuge tube. But we want to express cell density as # cells per mL of the culture. To convert our calculated value to # cells per mL, we first multiply # cells/slide X 2 because you diluted your sample by ½ when you added the iodine and then multiply by 5 to convert cells in 100 µL to cells per 500 µL (the size of the aliquot you took from the microcentrifuge tube). 𝑖. 𝑒. , # 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 500 µL 𝑜𝑓 𝑐𝑢𝑙𝑡𝑢𝑟𝑒 = 𝑛𝑢𝑚 𝑜𝑓 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑠𝑙𝑖𝑑𝑒 𝑋 10 6. However, we ultimately need to know the number of cells per mL of “pond.” To determine this value, divide the cell density of the zooplankton culture by the total volume of the “pond” (750 mL): 𝑛𝑢𝑚 𝑜𝑓 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 500 µL of culture 𝑖. 𝑒. , # 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑚𝐿 𝑜𝑓 “𝑝𝑜𝑛𝑑 𝑤𝑎𝑡𝑒𝑟” = 𝑣𝑜𝑙𝑢𝑚𝑒 𝑜𝑓 𝑝𝑜𝑛𝑑 (𝑚𝐿) 7. Compare your results to those obtained by the other pair of students. Determine the mean for # cells per mL of the pond of zooplankton and record this value in the space below and in the appropriate space on the class data spreadsheet. 8. Follow the directions posted in the fume hood to clean your Palmer- Maloney slide. 5 Formulation of Hypotheses Work with your group members to formulate hypotheses that predict the effects of increasing fertilizer concentration on various organisms in the microcosms. State a null hypothesis for each parameter and be sure to make each hypothesis as specific as possible. 1. What effect will increasing fertilizer concentration have on the health of the terrestrial plants? In thinking about effects on plant health, consider both growth and leaf colour (i.e., chlorophyll content). 2. What effect will increasing fertilizer concentration have on the biomass of the aquatic plant? 3. What effect will increasing fertilizer concentration have on the density of phytoplankton in the “pond”? 4. What effect will increasing fertilizer concentration have on the population of zooplankton (that feed on algae) in the “pond”? 5. How do you expect N levels to change in each microcosm over the duration of the experiment? 6. How do you expect P levels to change in each microcosm over the duration of the experiment? 6  LABORATORY 4 Eutrophication Objectives After completing this lab, you should be able to: Define the term “eutrophication” and explain how this process occurs and its importance in aquatic ecosystems. Develop and test hypotheses as to how nutrient enrichment will influence the various components of terrestrial and aquatic microcosms (terrestrial and aquatic plants, water chemistry, and aquatic microorganisms). Differentiate among major sources of scientific information and use CSE formatting to cite literature properly Readings Lab 4 Manual Assignment Guidelines Khan FA and Ansari AA. 2005. Eutrophication: An Ecological Vision. The Botanical Review. 71(4), 449–482. (Posted on D2L) Background We tend to think of aquatic ecosystems—lakes, rivers, etc.—as being separate from terrestrial ecosystems but the reality is that water bodies provide resources for many terrestrial organisms (e.g., fish as food for bears), and many nutrients cycle back and forth between aquatic and terrestrial ecosystems. One example of the interactions between aquatic and terrestrial ecosystems is eutrophication of water bodies. Healthy aquatic ecosystems are relatively low in dissolved nutrients and are described as oligotrophic (oligo = few; trophic = relating to nutrition). In contrast, eutrophic (eu = good, true) aquatic ecosystems have high nutrient levels and tend to be dominated by algae and/or other aquatic surface plants, which is generally not a desirable state. Eutrophication is the process by which a water body becomes enriched in nutrients from terrestrial ecosystems and can happen by natural processes (e.g., as the banks of a river are eroded), but human activities (i.e., cultural eutrophication) such as the addition of fertilizers to aquatic systems are often the cause of, or at least speed up, the process. The primary nutrients needed by plants and algae include nitrogen (N), phosphorous (P), and potassium (K), which are the major constituents of fertilizers. The three numbers indicated on containers of fertilizers (e.g., 20–15–20) refer to the percentage of N, P, and K, respectively, in the fertilizer. Plants and algae require large quantities of nitrogen and phosphorous because they are major constituents of proteins and DNA; potassium is essential because of its important role in maintaining turgour pressure in plant cells and in the activation of many enzymes. Unlike green algae, cyanobacteria can fix gaseous nitrogen into a form that can be used by cells, and so are able to survive in nitrogen-poor environments. In many ecosystems, including the agroecosystems humans have created to grow our major crop plants, the availability of these three mineral nutrients, primarily N, is the major limiting factor for plant (and algal) growth. We add fertilizers and/or animal wastes to our cropland to enhance the naturally occurring forms of these nutrients. Unfortunately, these nutrients are normally added in quantities higher than what the plants can use immediately, and so excess nutrients are carried into water bodies as topsoil is eroded or via runoff; nutrients are also introduced via effluent from sewage treatment plants. Eutrophication due to human activities has become a major problem in many aquatic ecosystems (Carpenter et al. 1998). In this lab, you will set up microcosms to investigate the effects of increasing fertilizer application on terrestrial plants and on an aquatic ecosystem over the course of three weeks. Each microcosm consists of an upper chamber (that simulates a terrestrial ecosystem or “field”) containing soil, which drains into a lower chamber containing water (simulating an aquatic ecosystem or “pond”). Both chambers have air holes to allow for gas exchange. We will grow terrestrial plants in the upper chamber, and aquatic plants as well as phytoplankton (aquatic photoautotrophs, including cyanobacteria and eukaryotic algae) in the lower chamber. Phytoplankton are the foundation for most aquatic food webs, and tend to be quite short- lived, responding to increased nutrient availability with rapid increases in population growth. We will also introduce a zooplankton that feeds on phytoplankton, into the lower chamber. After three weeks, we will harvest the microcosms and collect data on growth of terrestrial and aquatic plants, changes in populations of aquatic microorganisms, and changes in water quality in response to different fertilizer concentrations.  LABORATORY 3 Enzymes II Procedures I. Measuring Kinetics of b-Galactosidase Work in pairs 1. Collect the enzyme solution that has been prepared for you – keep the solution on ice. This is a 1:10 dilution of the stock enzyme solution you used in Lab 2. 2. You will be assigned to one of two groups. Your group determines the ONPG concentrations you will use. a. Group 1: 0, 5, 10, 20, and 40 mM ONPG b. Group 2: 0, 3, 7.5, 15, and 30 mM ONPG 3. Label 5 tubes with the ONPG concentrations assigned to your group. To each of these tubes, add 3.5 mL phosphate buffer and 0.5 mL of the appropriate ONPG solution that has been prepared for you. Which tube is the blank? Stagger the time you add enzyme to each tube, so the tubes don’t all finish at the same time. Start with the lowest concentration of ONPG and run each reaction for 4 minutes. 4. Add 0.5 mL enzyme solution to each tube, start the timer and vortex to mix. 5. After 4 minutes, stop the reaction by adding 0.5 mL sodium carbonate and vortex to mix. 6. Zero the spectrophotometer using your blank and measure the absorbance of each reaction tube, starting with the lowest concentration and ending with the most concentrated. Remember to zero the spectrophotometer and rinse the cuvette between samples. 7. Record the absorbance values in Table 3.1 below. If you measure an absorbance greater than 1.000, report the absorbance as 1.00 If you measure an absorbance less than 0.000, report the absorbance as 0.000 8. Use the standard curve you generated in Lab 2 to convert absorbance to o-nitrophenolate concentration and determine the reaction rate at each ONPG concentration. 9. Record the reaction rate at each ONPG concentration in the lab spreadsheet 1 II. Investigating Competitive and Non-Competitive Inhibitors Work in pairs You will work with an unknown chemical, “Inhibitor A”, and determine whether the chemical is a competitive or non-competitive inhibitor. 1. Use the same enzyme solution from Part 1 above – make sure it is kept on ice 2. Label 5 tubes with the ONPG concentrations assigned to your group. To each of these tubes, add 1.5 mL phosphate buffer, 2.0 mL Inhibitor A, and 0.5 mL of the appropriate ONPG solution. Which tube is the blank? Stagger the time you add enzyme to each tube, so the tubes don’t all finish at the same time. Start with the lowest concentration of ONPG and run each reaction for 4 minutes. 3. Add 0.5 mL enzyme solution to each tube, start the timer and vortex to mix. 4. After 4 minutes, stop the reaction by adding 0.5 mL sodium carbonate and vortex to mix. 5. Zero the spectrophotometer using your blank and measure the absorbance of each reaction tube, starting with the lowest concentrated and ending with the most concentrated. Remember to zero the spectrophotometer and rinse the cuvette between samples. 6. Record the absorbance values in Table 3.2 below. If you measure an absorbance greater than 1.000, report the absorbance as 1.00 If you measure an absorbance less than 0.000, report the absorbance as 0.000 7. Convert absorbance to o-nitrophenolate concentration and determine the reaction rate at each ONPG concentration. 8. Record the reaction rate at each ONPG concentration in the lab spreadsheet Clean Up 1. Rinse all cuvettes and tubes and place them upside down to dry 2. Empty the waste beaker into the waste carboy in the fume hood 3. Refill the water bottles with RO water 4. Tidy your bench and push in your stool 2 Table 3.1 Rate of ONPG hydrolysis catalyzed by b-galactosidase at various ONPG concentrations Concentration of ONPG Absorbance o-Nitrophenolate Reaction Rate (mM/min) (mM) (420 nm) Concentration (mM) 0 Table 3.2 Rate of ONPG hydrolysis catalyzed by b-galactosidase at various ONPG concentrations in the presence of “Inhibitor A” Concentration of Absorbance o-Nitrophenolate Reaction Rate (mM/min) ONPG (mM) (420 nm) Concentration (mM) 0 3  LABORATORY 3 Enzymes II Objectives Upon completion of this topic, you should be able to: Design an experiment to investigate the effect of inhibitors on enzyme activity Present, describe, and interpret your results Readings Lab 3 Manual Assignment Guidelines Fenton et al. (2022) Sections 3.5 (Role of Enzymes in Biological Reactions) and 3.6 (Factors that Affect Enzyme Activity) I. Measuring Kinetics of b-Galactosidase In this week’s lab, you will calculate the two main parameters of enzyme kinetics: The maximum rate of reaction for a given enzyme and substrate, or Vmax. The substrate concentration at which the reaction rate is half of its maximum value, or Km. To determine these parameters, reaction rates are measured over a range of substrate concentrations. Reaction rate can then be graphed as a function of substrate concentration, as shown in Figure 3.1. Vmax is the maximum rate at which an enzyme can catalyze the reaction under the conditions of your experiment. At Vmax, all enzyme molecules are complexed with substrate molecules and adding any additional substrate will not speed up the reaction. Km is the substrate concentration at which the reaction proceeds at its maximum catalytic efficiency. It is determined by reading the substrate concentration at half of its maximum rate (i.e., ½ Vmax). The type of graph shown in Figure 3.1 is known as a Michaelis-Menten plot, named for the researchers who developed this model of enzyme kinetics. (Maud Menten was a Canadian medical researcher and was one of the first women in Canada to earn a medical degree.) 1 Figure 3.1 Example of a Michaelis-Menten plot. E = enzyme; S = substrate; ES = enzyme-substrate complex We can obtain some information about the kinetics of an enzyme from a Michaelis-Menten plot, but one limitation of this type of plot is that Vmax is an asymptote (i.e., the curve will never reach the horizontal line that represents Vmax) and its value can only be determined if the reaction is run at high enough concentrations of substrates. We can correct this problem if we do a reciprocal graphing of our Michaelis-Menten plot to obtain a straight line, i.e., if we graph 1/V on the y-axis and 1/S on the x-axis, as shown in Figure 3.2. This graphical representation of enzyme kinetics is known as a Lineweaver-Burk plot, in which the slope of the line equals Km/Vmax, and the y-intercept equals 1/Vmax. The line can be projected back past the y-intercept to intersect with the x axis; the point at which the line intersects the x-axis equals –1/Km. Thus, using a Lineweaver-Burk plot allows us to determine Vmax and Km with much greater precision than a Michaelis-Menten plot. To determine Vmax and Km for b-galactosidase, you need to measure the effect of increasing ONPG concentration on the rate of its hydrolysis, and then graph reaction rate as a function of substrate concentration. All the necessary solutions have been prepared for you. Figure 3.2 Example of a Lineweaver-Burk plot 2 II. Investigating Competitive and Non-Competitive Inhibitors An enzyme’s ability to bind to other molecules and catalyze a reaction is determined by its structure (shape), which is determined by the sequence of amino acids that are linked together. Once the amino acids have been linked together, the chemistry of the individual amino acids and their order in the chain will drive the folding of the polypeptide chain into a three-dimensional structure. This folding brings specific amino acids close together to form the active site, where substrates bind and are converted into products. Changes in the three-dimensional structure of an enzyme (and thus the position of specific amino acids) will affect the enzyme’s ability to catalyze a reaction. If the entire shape of the enzyme is disrupted (i.e., if it is denatured, for example by exposure to high temperatures), all catalytic activity is lost. However, even more subtle changes in enzyme shape can increase or decrease its catalytic ability. These shape changes can occur because of changes in environmental temperature or pH or can be caused by genetic mutations. The presence of inhibitors also alters enzyme activity. Some inhibitors are naturally present in cells and play an important role in regulation of enzyme activity, while other inhibitors are drugs that we have designed to regulate enzyme activity. Inhibitors can be grouped into two general categories: Competitive inhibitors are similar in shape to an enzyme’s substrate and compete with the substrate for binding to the active site (Figure 3.3a). Since the two molecules compete for the active site, the effect of a competitive inhibitor on reaction rate depends on the relative concentrations of the substrate and the inhibitor. In competitive inhibition, Vmax is unchanged while Km increases. Why? Non-competitive inhibitors do not compete with the substrate for the active site, but instead bind to other sites on the enzyme (Figure 4.3b). This binding changes the shape of the enzyme such that even if the substrate can still bind to the active site, the active site functions less effectively, reducing the rate at which the substrate is converted to product even at high concentrations of substrate. In non-competitive inhibition, Vmax is reduced but there is no effect on Km. Why? Figure 3.3 Enzyme inhibitors 3  LABORATORY 2 Enzymes I Procedures Preparing a standard curve A stock solution of o-nitrophenol has been prepared for you. You will need to dilute the stock solution several times to create solutions with different concentrations of o-nitrophenol. Refer to Table 2.1 to determine the dilutions you will have to make. Table 2.1 Dilutions of a 1.5 mM stock solution of o-nitrophenol and corresponding absorbance readings (420 nm) to prepare a standard curve for o-nitrophenol. Sodium Standard Curve Final Concentration Phosphate Absorbance Carbonate (o-nitrophenol) Dilution of o-nitrophenolate Buffer (mL) (420 nm) (mL) (mL) (mM) 4.5 0.5 0.0 4.4 0.5 0.1 4.3 0.5 0.2 4.2 0.5 0.3 4.1 0.5 0.4 4.0 0.5 0.5 3.9 0.5 0.6 3.8 0.5 0.7 3.7 0.5 0.8 3.6 0.5 0.9 3.5 0.5 1.0 Use of Micropipettes Micropipettes are used to measure very small volumes of liquid accurately. In this course you will use two different pipettors able to measure liquids in the range of 10–100 µl (yellow button) or 100–1000 μl (blue button). The barrel of the micropipette houses a spring- loaded piston attached to a button that is depressed with your thumb to draw up and expel liquid (Figure 2.6). 1  Figure 2.6 Use of micropipettes 1. Set the volume required by releasing the lock and turning the setting ring. Never turn the ring below 100 μl (shown as “010” from top to bottom) or above 1000 μl (shown as “100”). 2. Use a clean sterile tip each time you measure a different solution to prevent cross- contamination of solutions. The tips have been autoclaved and are the only part of the instrument that is sterile. Allow only the tip, not the barrel, to contact the solution. The tip must be placed securely on the barrel of the micropipette so that there is an airtight seal. If it is too loose, the tip will leak air and not give an accurate measurement, or it may even fall off. Discard used tips into the disposal bags on your bench. 3. As shown in Figure 2.6, the control button on top of the micropipette has three positions: At Rest: The position of the button when it is not being used. Stop 1: This measures the volume selected in the window. Depressing the button from the “At Rest” position to Stop 1 measures the volume selected in the window. Stop 2: This blows out the last drops when expelling liquid from the tip. 4. To measure a required volume of liquid: a. Securely attach the micropipette tip. b. Press control button down to the first stop and hold. c. Hold micropipette vertically and immerse tip approximately 3 mm into the liquid. d. Allow control button to glide back SLOWLY. Never allow it to snap back. e. Slide tip out of the liquid along the inside of the vessel to remove any remaining droplets. Remember that: 0.1 mL = 100 μl and 1.0 mL = 1000 μl 5. To transfer this volume of liquid: 2  a. Place the tip in the receiving vessel, holding the tip at an angle against the inside of the vessel. b. Press the control button slowly down to the first stop. c. Press the control button down to the second stop (blow-out) to empty the tip completely. d. Hold down the control button. Slide tip out along the inside of the vessel. e. Let the control button glide back slowly so that it is fully released. f. To eject the tip after the liquid has been dispensed, hold the micropipette over the disposal bag and depress the eject button. This will shoot the tip off the end of the micropipette into the disposal bag. Be careful to aim correctly. Micropipettes are expensive and delicate. Please handle them carefully; do not drop them and do not exceed the maximum or minimum volumes. When not in use, keep the micropipettes in the micropipette stands—do not lay them down on the bench. Zeroing the Spectrophotometer One potential problem in any experiment using a spectrophotometer is that there may be other molecules in a solution besides the pigmented product that also absorb light. For example, what if the buffer or the enzyme in the enzyme-buffer solution absorbed light? You want to be sure that you are measuring only how much light is absorbed by the pigmented product, so you need to prepare a blank solution that contains all components of the solution other than the pigmented product molecule. This blank is used to set the spectrophotometer to zero. What should your blank tube for the standard curve contain? Using the Spectrophotometer 1. Set the spectrophotometer to read 420 nm. Ensure that the spectrophotometer is set to measure absorbance (Abs), not transmittance. 2. Follow the demonstration of your TA to create dilutions of the stock solution of the pigment molecule as indicated in Table 2.1. Make sure you are careful with the pipettes and the accuracy of your measurements. Mix the contents of the tubes using a vortex as demonstrated by your TA. 3. Transfer an aliquot of your blank solution to a cuvette. Do NOT overfill the cuvette! 4. Wipe the outside of the cuvette with a Kimwipe to remove fingerprints or moisture, which can scatter or absorb light. Insert the cuvette into the sample compartment as demonstrated by your TA. 5. Zero the spectrophotometer and remove the blank cuvette; DO NOT throw away the blank solution. You will need it to generate your standard curve and to measure enzyme activity. The spectrophotometer is now ready to measure the light absorbance of your pigment. 6. Measure the absorbance of each dilution. You should blank the spectrophotometer between each reading to improve accuracy. Record the absorbance of each dilution in Table 2.1. 7. Use the graph paper provided to prepare the standard curve for o-nitrophenolate. If you do not obtain a straight line, ask your TA for help. 3 Dilution of an Enzyme Stock Solution You can determine the activity of b-galactosidase by measuring the absorbance of the o- nitrophenolate produced by the hydrolysis of ONPG and using your standard curve to convert the absorbance value into concentration and then dividing by the reaction time. The course technicians have prepared a stock solution of b-galactosidase by grinding a LactaidTM tablet in 0.1 M phosphate buffer (pH 8). 1. Obtain a sample of the stock solution of b-galactosidase (labelled E for enzyme) and place in an ice bucket. At this point, the activity of the enzyme stock is unknown, so you first need to determine a proper concentration of b-galactosidase to use for the rest of your experiment. 2. Clearly label four microcentrifuge tubes with the dilutions shown in Figure 2.7. Prepare four serial (1:10) dilutions of the stock solution and place them in the ice bucket. Keep all solutions for the duration of the experiments in case something goes wrong, and you need to restart. Figure 2.7 How to prepare a serial dilution Measuring Enzyme Activity It’s important to choose a concentration of enzyme that is neither too dilute (it is hard to measure activity if there isn’t much enzyme) nor too concentrated (there is an upper limit to the absorbance that can be measured by the spectrophotometer). Use the procedure below to determine the activity of the b- galactosidase enzyme at each dilution. 4 1. Clearly label four new test tubes with the enzyme dilutions (e.g., 1:10, etc.) 2. Pipette 3.5 mL of buffer and 0.5 mL ONPG to each tube. It Is important to stagger the time you add the enzyme to each tube, so that the incubation time for the tubes don’t finish at the same time 3. Start the reaction by adding 0.5 mL of the lowest enzyme dilution (1:10 000) to the appropriate tube and start a timer. Vortex the tube to mix the contents. After 4 minutes, stop the reaction by adding 0.5 mL sodium carbonate. Vortex the tube to mix the contents. 4. Zero the spectrophotometer using the blank solution you used for the standard curve. Don’t forget to wipe the outside of the cuvette with a Kimwipe. 5. Transfer an aliquot from your reaction tube to a cuvette. Insert the cuvette into the sample holder of the spectrophotometer and record the absorbance reading in Table 2.2. Rinse the cuvette. 6. Repeat steps 3–5 for the remaining three enzyme dilutions. 7. Use your standard curve to convert absorbance to the concentration of o-nitrophenolate produced in the reaction. Record these values in Table 2.2. 8. Divide the o-nitrophenolate concentration by 4 minutes (the time interval during which the reaction occurred) to determine the reaction rate (mM o-nitrophenolate formed per minute). Record these values in Table 2.2. Table 2.2 Product formation and reaction rate for different enzyme concentrations. Enzyme Absorbance o-Nitrophenolate Reaction Rate Tube –1 Dilution (420 nm) Concentration (mM) (mM × min ) #1 1:10 #2 1:100 #3 1:1000 #4 1:10 000 Briefly describe the trend you observe (i.e., as enzyme concentration increases, what happens to the reaction rate?) 5  LABORATORY 2 Enzymes I Objectives Upon completion of this topic, you should be able to: Explain how spectrophotometry can be used to study the kinetic parameters of an enzyme and to measure enzyme activity Prepare and use a standard curve Prepare dilutions of a stock solution Readings Lab 2 Manual Fenton et al. (2023) Sections 3.5 (Role of Enzymes in Biological Reactions) and 3.6 (Factors that Affect Enzyme Activity) Introduction Enzymes are biological catalysts that increase the rates of biochemical reactions in cells by facilitating the conversion of a specific substrate to a product by lowering the activation energy required for that reaction to occur. They are essential for all cellular processes; for example, they aid in the conversion of light or chemical bond energy into ATP, the transformation of nutrients acquired from the environment into usable forms, the replication of DNA, and the synthesis of proteins. Most enzymes are proteins, although RNA can also act as a catalyst in some processes (e.g., in protein synthesis). The catalytic effect of an enzyme is determined by its shape, particularly the shape of its active site, which will bind only with certain substrates (reactants), forming an enzyme- substrate complex (Figure 2.1). The enzyme facilitates a chemical change, allowing the substrate to be converted into a product molecule. The product is released from the active site, and the unchanged enzyme is free to combine again with other substrate molecules. The rate of an enzyme-catalyzed reaction depends on the concentrations of both enzyme and substrate, as well as enzyme sensitivity to high temperatures or extreme pH, and the presence of regulatory compounds that alter activity. Enzyme kinetics involves studying how fast an enzyme converts substrate to products, as well as investigating the effects of varying the conditions of a reaction. One key parameter of enzyme kinetics is enzyme activity: the rate or velocity of the reaction catalyzed by the enzyme. Figure 2.1 Diagram of enzyme function 1  How Can We Measure Enzyme Activity? In general, enzyme activity can be determined by combining the substrate and enzyme in solution in a test tube, and then measuring either the decrease in the amount of substrate or the increase in the amount of product as the reaction occurs; the more active the enzyme, the more substrate that will be converted to product. How can we measure the changing concentration of substrate or product? If either the substrate or product molecules absorb specific wavelengths of visible light (i.e., they are pigments), then we can measure the concentration of these pigment molecules by measuring how much light is absorbed as the reaction occurs using an instrument called a spectrophotometer. In some cases, the substrate and products are colourless. In these cases, we can add indicator chemicals to the reaction that help to generate a coloured product that we can measure. Spectrophotometers measure how much light is absorbed by a solution relative to how much light passes through the solution (transmittance); they are among the most widely used research instruments in biology. By placing samples of the solution containing the substrate and enzyme in the spectrophotometer at different times as the reaction occurs, the relative amount of light absorbed over time will tell you how the concentration of the pigment molecule (either substrate or product) is changing. How a spectrophotometer works 1. Light is separated into its different wavelengths by a prism (Figure 2.2). 2. A slit allows only one specific wavelength of light, selected by the user, to shine on the sample. A specific pigment molecule will absorb some wavelengths better than others. By changing the position of the slit relative to the light passing through the prism within the spectrophotometer, the spectrophotometer can be set to use a wavelength strongly absorbed by the molecule under investigation. Figure 2.2 Measurement of light absorption by a spectrophotometer 3. Light of the selected wavelength is then passed through the sample solution. Some of the light will be absorbed by the solution, while the rest of the light will be transmitted through it. The higher the concentration of the coloured pigment molecule or indicator in solution (i.e., the darker the solution), the greater the absorbance. 4. The transmitted light strikes a photocell that converts the light energy to an electric current, which can be measured by a meter. Note: Spectrophotometers measure the absorbance of light by a molecule, NOT the absorbency of a molecule. 2 Using a standard curve to determine the concentration of pigment in solution The absorbance reading that a spectrophotometer displays tells you how much light of the selected wavelength a particular pigment molecule absorbs, relative to the amount of light that passes through the solution. If there are few molecules that absorb at the wavelength that you are testing, the value reported will be very low (e.g., 0.005). As the concentration of the molecules increases in the solution, you will see an increase in absorbance values. Notice that there are no units on absorbance values! Unfortunately, the spectrophotometer does not tell you the concentration of the pigment molecule in the solution, which is what we need to know to determine enzyme activity. How can we convert absorbance values to concentration? If we make solutions that contain known concentrations of the pigment molecule that we are measuring (i.e., the product molecule of a reaction) and then determine the absorbance of each solution, we can construct a graph known as a standard curve, which displays the relationship between absorbance and concentration for that pigment. To make this graph, you plot the absorbance you measure as a function of pigment concentration (i.e., plot absorbance on the y-axis and pigment concentration on the x-axis); you should obtain a straight line as shown in Figure 2.3. Figure 2.3 Example of a standard curve A standard curve allows you to convert absorbance values of a pigment molecule to its equivalent concentration. To convert an absorbance reading for a solution containing an unknown concentration of the pigment molecule, you could determine the equation of the line and use that equation to calculate the concentration. In Lab 2, you will use a quick and easy method in which you: Locate the point on the y-axis that corresponds to the measured absorbance value for that solution. Draw a straight line horizontally from that point over to the standard curve. Draw a straight line down to the x-axis. The value where your line intersects the x-axis is the concentration of pigment in your unknown solution. 3 Kinetic Study of the Enzyme b-Galactosidase Over the next two labs, you will investigate the activity and kinetics of an over-the-counter degradative enzyme, b-galactosidase, which is present in Lactaid™ and used to treat lactose intolerance. Lactose is a disaccharide found in milk and other dairy products. Each lactose molecule consists of glucose and galactose linked together by a b-(1,4) glycosidic bond. This bond can be broken by a class of enzymes called b-galactosidases. Lactase is a member of this class of enzymes and is responsible for breaking lactose into its constituent glucose and galactose monomers (Figure 2.4). Figure 2.4 Degradation of lactose by β-galactosidase Humans produce high levels of b-galactosidase at birth, enabling them to break down the lactose in breast milk, but the ability to digest lactose as adults depends on ethnic background. Most people of Northern European heritage produce adequate levels of b-galactosidase as adults, but people of Asian or indigenous origin often produce less of this enzyme as they grow older and so cannot digest lactose and are said to be lactose-intolerant. In humans, lactose is degraded in the small intestine; when lactose-intolerant individuals consume milk or other dairy products, lactose accumulates in the small intestine, where it is fermented by bacteria, resulting in diarrhea, gas formation, and the related symptoms of bloating and cramping. However, commercial products such as Lactaid™ allow lactose-intolerant people to consume dairy products without suffering these side effects. Lactaid™ consists of b-galactosidase combined with various other ingredients in tablet form. In Lab 2, you will work with other students at your bench to measure the enzymatic activity of b- galactosidase from Lactaid™. In Lab 3, you will investigate the effect of different inhibitors. 4  Measuring the activity of b-galactosidase One challenge in measuring the activity of b-galactosidase is that neither the natural substrate (lactose) nor the products (glucose and galactose) absorb light. However, there are artificial substrates for b-galactosidase that do produce a pigmented molecule as a product. For example, we can use o-nitrophenyl-b-D-galactopyranoside (ONPG) as a substrate for this enzyme. ONPG consists of galactose and o-nitrophenol joined together by a b-(1,4) linkage like that found in lactose. Breakage of this bond by b-galactosidase produces both galactose, which is not pigmented, but also o-nitrophenol, which is pale yellow in colour. To produce a more intensely coloured end-product that is easier to measure, we can expose o-nitrophenol to alkaline pH (>8), converting it to o-nitrophenolate, which has an intense yellow colour, as summarized in Figure 2.5. O T E S !-galactosidase CH2OH H2O CH2OH O O OH pH > 8 HO HO O + HO O OH OH NO2 NO2 NO2 OH OH ONPG Galactose o-Nitrophenol o-Nitrophenolate (colorless) (colorless) (yellow) Figure 2.5 ONPG hydrolysis catalyzed by β-galactosidase Figure 2.5. ONPG hydrolysis catalyzed by β-galactosidase. To measure b-galactosidase To measure activity, a solution β-galactosidase containing activity, the enzyme a solution is mixed with containing thea buffer enzymedesigned is to keep the pH of the solution constant. The substrate ONPG is then added, and the solution is mixed with a buffer designed to keep the pH of the solution constant. The incubated at a specific temperature. substrate ONPG is then added, and the solution is incubated at a specific temperature. As β-galactosidase As β-galactosidase breaks breaks down down ONPG, and o-nitrophenol ONPG is formed, and o-nitrophenol a pale-yellow colour develops. At thatispoint, an alkaline formed, a pale solution yellow (1.0colour M sodium carbonate)At develops. is added, which an that point, inactivates alkalinethesolu- enzyme and stops the hydrolysis of ONPG. The increase in pH also causes the o-nitrophenol that was formed by tion (1.0 M sodium carbonate) is added, which inactivates the enzyme the hydrolysis of ONPG to be converted to o-nitrophenolate, which is dark yellow. and stops the hydrolysis of ONPG. The increase in pH also causes the The exact time interval between o-nitrophenol that was when the substrate formed by the was added to theofenzyme-buffer hydrolysis ONPG to be solution and when converted the reaction was stopped by the addition of the alkaline solution must be noted to calculate the rate of to o-nitrophenolate, which is dark yellow. The exact time interval between enzyme activity. The amount of yellow product formed depends on enzyme kinetics, i.e., how fast the enzymewhen theONPG converts substrate was added to galactose to the enzyme-buffer solution and when and o-nitrophenol. the reaction was stopped by the addition of the alkaline solution must be noted in order to calculate the rate of enzyme activity. B. INVESTIGATING THE !-GALACTOSIDASE ACTIVITY FROM BEANO® Beans such as chickpeas and lentils are excellent sources of fiber and protein and are an important component of a healthy diet. However, eat- ing beans can have unfortunate side effects: the oligosaccharides in the beans can be difficult to digest if you aren’t used to eating them, meaning 5 that you lack adequate levels of the digestive enzymes needed to break down these carbohydrates. These undigested sugars accumulate in the Dilutions Dilutions are used in a variety of procedures, such as making a standard curve and preparing specific concentrations of a stock solution. Dilutions are made by pipetting a volume of an undiluted or stock solution into a volume of liquid, known as the diluent. Dilution (D) is the ratio of the volume of the original solution to the total volume of the diluted sample: D = volume of original solution / (volume of original solution + volume of diluent) For example, if you add 1.0 mL of your original solution to 9.0 mL of diluent, the total dilution will be a 1 in 10 dilution, which is expressed as 0.1 or 10−1. Dilution = 1.0 mL / (1.0 mL + 9.0 mL) = 1.0 / 10 = 0.1 or 10−1 Once you know the dilution, you can determine the concentration of the diluted sample (Cd) using the following mathematical formula: Cd = Cu × D Cd = concentration of diluted sample Cu = concentration of undiluted (original) sample D = dilution Some courses, such as chemistry, ask you to calculate the dilution factor, which is the total volume of your solution (i.e., the volume of original solution + volume of diluent), or the denominator of the equation above. The dilution factor for this example would be 10. Preparing Fixed Volumes of Specific Concentrations from Stock Solutions For some labs in this course, you will need to make a specific volume of known concentration from a stock solution. The formula below is a quick approach to calculating such dilutions where: V 1C 1 = V 2C 2 V1 = volume of the original or stock solution C1 = concentration of stock solution V2 = volume of the new solution C2 = concentration of the new solution For example: Suppose you have a stock solution of 100 mg mL−1 glucose (C1). You want to make 200 µL (V2) of a solution with a concentration of 25 mg mL−1 (C2). Your unknown is V1, or the volume of the stock solution that you will need to prepare this diluted solution. V1C1 = V2C2 , so, V1 = V2C2 / C1 V1 = (0.2 mL × 25 mg mL−1) / 100 mg mL−1 V1 = 0.05 mL, or 50 μl 6  LABORATORY 7 Harvesting the Eutrophication Experiment Objectives After completing this lab, you should be able to: Collect and analyze data relating to the effect of fertilizer on various components of terrestrial and aquatic microcosms (terrestrial and aquatic plants, water chemistry, and aquatic microorganisms). Summarize changes in water quality and primary productivity that occurred because of fertilizer treatment in microcosms; draw conclusions about your hypotheses. Readings Fenton et al. (2022): Section 28.2 (Nutrient Cycling in Ecosystem), pages 764-771. Chislock MF, Doster E, Zitomer RA and Wilson AE. 2013. Eutrophication: causes, consequences, and controls in aquatic ecosystems. Nature Education Knowledge. 4(4):10 Smith VH, Schindler DW. 2009. Eutrophication science: where do we go from here? Trends in Ecology and Evolution. 24: 201–207. Smith VH, Tilman GD, Nekola JC. 1999. Eutrophication: impacts of excess nutrient inputs on freshwater, marine and terrestrial ecosystems. Environmental Pollution. 100: 179–196. I. PROCEDURES FOR HARVESTING MICROCOSMS Your group will harvest your microcosm: working in pairs, you will measure the height and chlorophyll content of the terrestrial plant, the final mass of the aquatic plant, the final cell density of the zooplankton, and the Nitrogen levels, Phosphorus levels, and chlorophyll content of the “pond” water (as an estimate of cell density of phytoplankton). Record all data in both the appropriate tables on page 6 below and in the class data tables. Carefully remove the tape from your microcosm and separate the top and bottom sections. Discard the tape in the garbage. 1 A. HARVESTING THE AQUATIC PLANTS 1. Carefully remove each plant by gently pulling up on the base of each plant to uproot it from gravel. 2. Gently blot the plants dry (only blot them enough to remove excess water from the plant’s surface; the plants do not need to be completely dry). 3. Determine the total biomass of the plants and record this value in Table 7.1 and in the class data table. 4. Dispose of the plants in the autoclave bag at the front of the lab. B. MEASURING CHLOROPHYLL CONCENTRATION OF THE “POND” WATER Total phytoplankton biomass can be estimated by determining the total chlorophyll content of the phytoplankton community in the lower microcosm chamber. 1. Mix the “pond” water thoroughly with a stir stick, and then transfer 100 mL to a graduated cylinder. 2. Place one piece of Whatman filter paper into the Büchner funnel at your bench. Turn on the vacuum. Pour 10 mL of distilled water onto the filter paper to seal it to the funnel. 3. Slowly pour the 100-mL sample of pond water onto the filter paper. When all the pond water has been filtered, turn off the vacuum. You will use the filtered water for Parts D and E 4. Remove the filter paper using forceps and place the filter in a centrifuge tube. Add 6 mL of 95% ethanol. Vortex to mix well, wrap the tube in tinfoil, and let sit for 30 minutes. 5. During those 30 minutes, continue with Steps D and E. 6. After 30 minutes, give the tube to your TA, who will centrifuge the tube at 3000 rpm for 5 minutes. 7. While you are waiting for your sample to be centrifuged, set your spectrophotometer to 649 nm. Fill a cuvette with ethanol to use as the blank. 8. Carefully pipette the supernatant from the centrifuge tube into a cuvette. Blank the spectrophotometer and read the absorbance of your sample. A649 = ____________ 9. Reset your spectrophotometer to read absorbance at 665 nm and blank the spectrophotometer. Read absorbance of your sample. A665 = ____________ 10. Use the following equation to calculate the amount of chlorophyll a (µg mL–1) in your sample: Chlorophyll a (µg mL–1) = 13.95(A665) – 6.88(A649) 11. Record this value in Table 7.2. Enter this value in the class data table. 2 C. DETERMINING FINAL POPULATION SIZE (CELL DENSITY) OF ZOOPLANKTON 1. Mix the “pond” water thoroughly with a stir stick. Remove a 500 µL aliquot of the pond water to a 1.5-mL microcentrifuge tube. 2. Add 500 µL of Lugol’s iodine to the microcentrifuge tube, close the lid, and invert tube several times to mix the contents. Lugol’s iodine will kill the cells, enabling you to count the number of cells accurately. 3. Obtain a Palmer-Maloney counting slide; add a cover slip and aliquot 100 µL from the microcentrifuge tube to one of the side ports on the slide. 4. Work with a partner to use a light microscope to count the number of cells, as you did when you set up the microcosm. Use the 10X objective lens and sequentially select 5 fields of view, moving from left to right across the slide. One person should count the number of zooplankton cells visible in each field of view while the other person records these values and keeps track of the number of fields of view that you examine. Then calculate the final cell density per mL of “pond” water using the following equation: (𝑣𝑜𝑙𝑢𝑚𝑒 𝑜𝑓 “𝑝𝑜𝑛𝑑 𝑤𝑎𝑡𝑒𝑟” 𝑎𝑑𝑑𝑒𝑑) 𝑛𝑢𝑚 𝑜𝑓 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑠𝑙𝑖𝑑𝑒 = 𝑐𝑒𝑙𝑙𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 × , 𝑎𝑟𝑒𝑎 𝑜𝑓 𝑓𝑖𝑒𝑙𝑑 𝑜𝑓 𝑣𝑖𝑒𝑤 𝑎𝑡 10𝑋 𝑝𝑜𝑤𝑒𝑟 × 𝑑𝑒𝑝𝑡ℎ 𝑜𝑓 𝑠𝑙𝑖𝑑𝑒 𝑟𝑒𝑠𝑒𝑟𝑣𝑜𝑖𝑟 ? 8 > × 𝑛𝑢𝑚 𝑜𝑓 𝑓𝑖𝑒𝑙𝑑𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 For our slides, this equation can be simplified to: 100 𝑚𝑚! 𝑛𝑢𝑚 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑠𝑙𝑖𝑑𝑒 = # 𝑐𝑒𝑙𝑙𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 ∗ 0.02 𝑚𝑚" 𝑋 0.4 𝑚𝑚 𝑋 5 5. The # cells/slide is the # cells in 100 µL of solution from the microcentrifuge tube. We want to express cell density as # cells per mL of “pond” water. To convert our calculated value to # cells per mL, we first multiply # cells/slide X 2 because you diluted your sample by ½ when you added the iodine and then multiply by 10 to convert the number of cells in 0.1 mL to the number of cells in 1 mL. 𝑖. 𝑒. , # 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑚𝐿 𝑜𝑓 “𝑝𝑜𝑛𝑑 𝑤𝑎𝑡𝑒𝑟” = 𝑛𝑢𝑚 𝑜𝑓 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑠𝑙𝑖𝑑𝑒 𝑋 20 Record the final cell density (# cells per mL of “pond water”) of zooplankton in Table 7.3 and enter this value in the class data table. 3 D. MEASURING AMMONIUM CONCENTRATION Gently swirl the filtered pond water you collected in Step B. Follow the instructions on the side bench for measuring ammonia concentration. Record this concentration in Table 7.4 and in the class data table. E. MEASURING PHOSPHATE CONCENTRATION Gently swirl the filtered “pond” water you collected in Step B. Follow the instructions on the side bench for measuring phosphate concentration. Record this concentration in Table 7.4. and in the class data sheet. F. MEASURING HEIGHT OF TERRESTRIAL PLANTS 1. Cut off each plant stem at the soil surface and lay the plants on a damp paper towel. 2. Carefully lay each plant along a ruler, being careful not to break the stem. Measure the height of the main stem in mm. Record the heights in your notebook. 3. Calculate the average shoot height for the plants from your microcosm; enter this value in Table 7.5 and in the class data table. 4. You will use the terrestrial plants for Step G. Empty the soil mixture from the top section into the garbage can. Rinse the top section with distilled water and invert on a paper towel to dry. G. TOTAL CHLOROPHYLL CONTENT OF TERRESTRIAL PLANT LEAVES 1. Choose the greenest leaf from the terrestrial plants in your microcosm and use the borer to cut a circular disc from the leaf, avoiding the midline vein. 2. Transfer the leaf disc to a mortar and pestle and grind it with 2 mL of ethanol until dissolved. 3. Carefully pour the contents of your mortar into a labeled centrifuge tube. Add ethanol to bring the volume up to 4 mL. 4. Give the centrifuge tube to your TA who will centrifuge it at 3000 rpm for 5 minutes. 5. While you are waiting for your sample to be centrifuged, set your spectrophotometer to 649 nm (the absorption maximum for chlorophyll b). Fill a cuvette with ethanol to use as the blank. 6. Carefully pipette the supernatant from the centrifuge tube into a cuvette. Blank the spectrophotometer and read the absorbance of your sample. A649 = __________ 7. Reset your spectrophotometer to 665 nm (the absorption maximum for chlorophyll a) and blank the spectrophotometer. Read absorbance of your sample. A665 = __________ 4 8. Use the following two equations to calculate the amount of chlorophyll a and b (in µg mL–1) found in your leaf sample: Chlorophyll a (µg mL–1) = 13.95(A665) – 6.88(A649) Chlorophyll b (µg mL–1) = 24.96(A649) – 7.32(A665) Total chlorophyll = chlorophyll a + chlorophyll b = _______ µg mL–1 Record these values in Table 7.5 and in the class data table. 9. Dispose of terrestrial plants in the autoclave bag at the front of the room. H. CLEANING UP THE LOWER CHAMBER 1. Pour the remainder of the “pond” water into the designated waste container, being careful not to let any gravel go into the waste container. 2. Remove the gravel and place it in the designated bucket on the side bench to be reused. 3. Rinse the bottom chamber well with distilled water and invert it on a paper towel to drain. Remove any remaining tape or labels from the microcosm. 5 Table 7.1 Final mass of aquatic plants after three weeks of treatment. Final Mass of Microcosm Aquatic Plants (g) (Total of 4 plants) Table 7.2 Chlorophyll a content of “pond” water in lower microcosm chambers following three weeks of treatment. Chlorophyll a Microcosm (µg mL–1) Table 7.3 Cell density of zooplankton in “pond” water in lower microcosm chambers and following three weeks of treatment. Final Cell Density Microcosm (# cells per mL of pond) Table 7.4 Concentrations of ammonia/ammonium and phosphate in “pond” water in lower microcosm chambers following three weeks of treatment. Ammonia/ Ammonium Phosphate Concentration Microcosm Concentration (µg mL–1) (µg mL–1) Table 7.5 Shoot height and leaf chlorophyll content of terrestrial plants in upper chambers of microcosms after three weeks of treatment. Total Shoot Height (mm) Chlorophyll a Chlorophyll b Microcosm Chlorophyll (Average of 5 plants) (µg mL–1) (µg mL–1)