How the Concentration of Enzymes Affects the Breakdown of Starch

How the Concentration of Enzymes Affects the Breakdown of Starch
In the cleaning business, it?s important to get a maximum cleaning effect at a minimum cost. This is especially applicable to the washing of clothes (both commercially, before an item of clothing goes on the market, or at home). This means trying to wash clothes at the lowest possible temperature, to keep the amount of electricity used at a minimum, yet trying to make and maintain a low-priced washing product that cleans effectively. This is why many washing powders use enzymes: enzymes are biological catalysts that speed up the breakdown of certain substances (in this case the molecules in food stain). Enzymes have a certain optimum temperature (a temperature at which the enzymes function at its best). Optimum temperatures are different for every enzyme, but they tend to be around 45Â?C. This means that if enzymes are to be used in washing powders, the temperature at which the clothes are washed will have to be at the optimum temperature, in order to achieve maximum enzymatic effect.
This optimum temperature, in the case of the enzymes concerned (the enzymes that break down protein, fats and starch in food stains on clothes), is lower than the normal washing temperature of clothes, 60�C, which means the use of enzymes in washing powders will reduce the washing temperature, thereby making the wash more cost effective. The enzymes break down the food stain deposits much better and much more efficiently than a hot wash with normal soap. Over a longer period of time, using normal hot washings to wash the clothes will wear down the material the clothes are made of, and damage them, making them visibly less attractive. Enzymes on the other hand, do not damage the material. This means that the clothes themselves will be cleaner, less damaged, and washed at lower temperature (not as money spent on electricity), which has a very positive result on the use of enzymes in washing powders. This is the main reason why enzymes are used in washing powders. The enzyme I will be focusing on is a-amylase, which is an enzyme present in biological washing powders that breaks down starch deposits in food stains into maltose, which is then further broken down into glucose sub-units. I however, am only looking at the initial breakdown of the starch by a-amylase into maltose. Aim: The aim of my investigation is to investigate the effect that changing enzyme concentration has on the breakdown of starch (i.e. rate of breakdown) by a-amylase. Does concentration affect the rate at which a-amylase breaks down starch? If so, what effect would this have on the concentration of a-amylase used in washing powders? What percentage concentration is most suitable for the breakdown of starch, to be used in washing powders to clean clothes? Hopefully the following investigation will help me in finding the answers to these questions. Hypothesis: As I increase the concentration solutions of the enzymes, the rate at which the enzyme breaks down the starch in the agar plate will increase (i.e. the rate of breakdown will increase with an increase of enzyme concentration solutions). The investigation below will help to determine this, or to disprove this prediction. Enzymes are biological catalysts. They differ to inorganic catalysts in that they are specific in that they catalyse if not a few only one specific breakdown/reaction. Enzymes are large globular proteins. This means that they will have a very specific tertiary structure. This also means that the active site of an enzyme (the area on an enzyme where the reaction occurs) will have a very specified structure. This means that the substrate molecule will have to be of that specific shape to be able to fit into the active site of the enzyme. This is why enzymes are seen as so specific; there are usually only one substrate (or very few substrate) molecules that have this specific shape. This also helps to explain why enzymes are so susceptible to factors such as heat: a slight alteration of their overall tertiary shape means a slight change to the shape of the active site. Substrate molecules will therefore not be able to fit into the active site as well, which means the enzyme will not be able to catalyse as efficiently in comparison to their activity at the optimum temperature, which means there is a decrease in the rate of breakdown. When an enzyme?s tertiary shape has been so heavily altered that the substrate molecule will not fit into the active site at all, the enzyme molecule is seen to be ?denatured?. Enzymes are thought to work on the idea of a ?lock-and-key? mechanism. The idea behind this mechanism is that the substrate molecule fits into the rigid shape of the active site of the enzyme. This helps to explain the specificity of enzymes: a slight alteration in the shape of the active site means a mismatch in the shape of the substrate and the active site. However, this idea is rather illogical, as the idea depends on the random movement of substrate molecules into the active site of the enzyme. This analogy seems to be rather vague for a molecule so important as enzymes. A better theory to explain for the activity of enzymes is what is known as the ?induced fit? mechanism. In this mechanism, when a substrate molecule comes into close proximity with an enzyme molecule, the enzyme molecule enfolds the substrate and changes its shape accordingly, so that the enzyme takes up its most effective shape once the substrate has bonded with the active site of the enzyme. Just as the shape of a plastic bag is affected by what is held inside it, the shape of an enzyme is affected by the substrate that it bonds with. Enzyme solutions are a mixture of enzyme and other substances (in this case, it is enzyme and distilled water). Due to this mixture, a ratio is created of enzyme-water molecules. In a pure sample of enzyme, no water is present, and therefore only enzyme molecules are present. If water is added, however, there will be a certain number of water molecules for every enzyme molecule. As this ratio increases, the concentration of the enzyme solution decreases, as there are more and more water molecules for each enzyme molecule. Increasing the concentration of enzyme means increasing the number of enzyme molecules in the solution. This has the same type of effect upon considering the addition of substrate into the solution. Taking for example a pure enzyme solution, adding a substrate into this solution means the ratio between the enzyme molecules and the substrate molecules is fairly small, which means there are a small number of substrate molecules for each enzyme molecule. This means that there is a quick breakdown of substrate, as each enzyme molecule has less substrate to break down. However, decreasing the concentration of the enzyme solution means decreasing the number of enzyme molecules present, which means the ratio between the number of enzyme and substrate molecules gets bigger and bigger. This means that there are an increasing number of substrate molecules for each enzyme molecule, which implies that each enzyme molecule gets an increased workload, and will take longer to break down the substrate molecules. This suggests that if the enzyme solution concentration decreases, the period of time needed to break down the substrate increases (i.e. gets longer). Chosen techniques: At the beginning of this experiment, there were a number of different ways I could go about conducting the experiment. One way of doing this would be mixing enzyme solutions to starch suspension in test tubes and adding the iodine solution afterwards to test for the breakdown of starch. In a number of ways, this is a good method of conducting the method. As the experiment is done in test tubes, the test tubes could then be placed in a water bath, in order to keep the solutions at a constant 37�C, which is the optimum temperature. This would mean the enzyme would be working at its most efficient point, which would be very applicable to the idea of using enzymes in washing powder (37�C being close to the normal washing temperature using biological washing powders). In a number of ways this is also a bad way of conducting the experiment. It would take a great deal of time to set all of the equipment up (i.e. test tubes, water baths). Once the equipment has been set up, the actual obtaining of the results would be quite time consuming as well. Firstly, only one concentration can be done at one time. This means in order to test all of the enzyme concentration solutions and to then repeat them twice to get a reliable set of result would take an extremely large period of time. This method would also not very realistic; comparing the time needed to conduct the experiment this way to the time given to conduct the experiment. Secondly, once testing for the breakdown of starch, what would be done is the iodine solution would be added into the test tubes, to show how far the starch has been broken down. The results obtained from this, however, are only observations by the human eye. There is nothing there that can actually be measured using equipment of any sort. This means there is a lack of solid evidence to prove or disprove my hypothesis. A second way of conducting this experiment would be to use starch agar plates, and to make wells in the agar plates using a cork bored in which the enzyme concentrations are put. The agar plates would then be put in an incubator/oven, and left for a certain period of time. When taken out of the incubator/oven, iodine solution would be added to the starch agar to show how far around the wells the a-amylase has broken down the starch, which would be measure using Vernier calipers. In some ways this would be a very good method to conduct the experiment. Firstly, many different concentrations of enzyme solution can be tested at once, as more than one well can be bored in one agar plate (the enzyme will not react so far as that one concentration will take up one agar plate). This means a fairly small number of agar plates will be need to conduct the experiment, let alone do two repeats of the experiment. This means that a lot less time will be spent on doing the experiment. As the plates are left in an incubator/oven for a certain period of time, most of the time will be spent preparing the plates to go into the incubator/oven and obtaining results (which is comparatively still a very small period of time, if compared to the first method). Secondly, upon obtaining the results, when adding the iodine solution, a circle of faded colour will be seen where the starch has been broken down. This circle can be measured in diameter, and this measurement can be used as values for the rate of breakdown. This means there is solid concrete evidence on which I can base my analysis and with which I can prove or disprove my hypothesis. In some ways, however, this would be a bad method for conducting the experiment. The optimum temperature of a-amylase is 37�C, yet the highest temperature at which a successful experiment can be carried out on the agar plates at is 26�C. This means the temperature at which the experiment is carried out is a considerably lower temperature than the optimum temperature of the enzyme. This means that in order to obtain the same kind of results when conducting the experiment at the optimum temperature, the agar plates will need to be left in the incubator/oven for a much longer period of time. Comparing the two methods, I feel that the second method would be a much better option for conducting this experiment, as the results obtained are much more concrete and accurate than the results obtained in the first method. Taking time into account, the second method would be a much more logical option, as this would leave me more time in the emergency if something did happen to go wrong during the investigation, and the investigation would have to be repeated all over again. The second method would also be much easier to repeat. Apparatus: � Termamyl� 60T a-amylase granules � Distilled water (used to make concentration solutions) � Beaker (250cm³) � Measuring cylinder, 50cm³ (accurate to 1cm³, �0.5cm³) � Balance (accurate to 2d.p.) � Starch agar plates, pre-prepared (filled to a depth of 6mm, using 1% starch agar) � Cork borer, � 5mm � Vernier calipers (accurate to 0.1mm, �0.05mm error) � Stirring rod � Iodine solution of concentration 0.01M (used to test for rate of breakdown of starch) � Pipette Brief method: Concentration solutions at concentrations shown in ?Variables? section below will be made up. On one agar plate, three equidistantly placed holes of �5mm will be bored into the starch agar plates, using a cork borer. Two starch agar plates will be produced like this, to provide enough holes for the concentration solutions. Using a pipette, drop 4/5 drops of each concentration solution into the wells. The agar plates are then placed in an incubator/oven at 26�C for a period of 24 hrs. When the plates are taken out again, the plates are flooded with iodine solution, to show where the enzyme has broken the starch agar down. Iodine solution in the presence of starch is blue/black; iodine solution itself is orange. This means where the enzyme has broken the starch down, there will be a circle of orange inside a circle of blue/black. The diameter of this circle is measured using Vernier calipers, and written down on a results table. The experiment is repeated twice. Constants: The volume of distilled water used to make up the solutions for the enzyme concentrations will be kept the same throughout the whole experiment: 100cm³. If this volume of water is not kept the same throughout the experiment, there will be an imbalance between the concentrations. Some concentrations will be more dilute than necessary, as more water is being used, and some concentrations will be more concentrated, which would result in an unfair testing of the concentrations, and the results obtained would be false. The volume of distilled water used to make up the concentration solutions is therefore kept a constant. The depth at which the agar plates are filled will be kept the same: 6mm. If the agar plates are not filled to an equal depth, some agar plates will contain a larger volume of starch agar that others. This means that comparing one concentration on two different agar plates, on one agar plate the enzyme will have much more starch agar to break down than on the other plate. This means the two results obtained for the same concentration of enzyme will be very varied, making the results false and unusable. The depth to which the agar plates are filled with starch agar is therefore kept a constant. The same cork borer will be used to bore all the wells in the agar plates in this investigation, to ensure the holes are the same size. If the wells bored with the cork borer to store the enzyme concentration solutions are different sizes, different surface areas over which the enzyme reacts are provided, which means the test will not be a fair test, leaving the results obtained false. The diameter of the wells is therefore kept a constant. The temperature at which the starch agar plates are stored will be kept at a constant. This will be done by putting the agar plates into an oven/incubator, and leaving the incubator on at a constant 26�C. This, however, is still very far away from the optimum temperature of the a-amylase. It is not possible to store the agar plates with the enzyme in them at the optimum temperature of 37�C, because at temperatures slightly higher than 28-30�C, the agar jelly itself would melt, rendering the investigation useless, as nothing can be done with liquid agar. This is why the plates can only be kept at a constant 26�C. This means that the experiment itself with take a significantly longer period of time to do, as the enzyme will be reacting with the starch much slower than at the optimum temperature. Ideally, the pH of the starch agar should be kept at a constant, optimum pH of 5.6 (optimum pH for a-amylase). This will, however, create quite a problem, as in order to keep a constant pH a buffer would need to be introduced into the starch agar. Even if the pH buffer would be introduced into the starch agar, it cannot be ensured of the even division of starch suspension and pH buffer, which means in some areas of the starch agar plate, the pH would be higher than other areas. This means the starch broken down around one well by the a-amylase would be at a slightly higher or lower pH than the well next to it. In any case, considering the pH, there will be an eventual disagreement in the level of pH of the starch agar, which means the results given will not be totally accurate. However, as there is no other possible method considerable for pH equality in the starch agar, this factor will have to be ignored in the final results, which means the results themselves will not be as fully accurate as was first hoped. The concentration of the starch agar will be kept at a constant. A starch suspension of concentration 1% will be used to fill the agar plates to a depth of 6mm. If this were not kept at a constant, there would not be a continuous level in the concentration, which ultimately means that there is no continuous level in the number of substrate molecules held in the starch agar. This means that in each starch agar plate, a different number of substrate molecules will be held, which means each enzyme concentration will not have an equal number of substrate molecules to break down. An unequal number of substrate molecules mean results that cannot be compared with each other, as they have not been carried out with the same number of substrate molecules, which means the results will be false. Variables: The one variable that I will change is the concentration of the enzyme solution. A control will be set up at 0% concentration (i.e. no enzyme present at all), in order to be able to compare the effect with enzyme and without enzyme. To make a 1% suspension of the a-amylase, 1g of a-amylase granules needs to be dissolved in 100cm³ of distilled water. However, the recommended concentration for a-amylase used in washing powders is 0.1%. This means that I will need to add 0.1g of the a-amylase granules to 100cm³ of distilled water. I will then proceed to go down in concentration by 0.02%. This will leave me with concentrations of: Ø 0.1% ? 0.1g a-amylase granules in 100cm³ distilled water Ø 0.08% ? 0.08g a-amylase granules in 100cm³ distilled water Ø 0.06% ? 0.06g a-amylase granules in 100cm³ distilled water Ø 0.04% ? 0.04g a-amylase granules in 100cm³ distilled water Ø 0.02% ? 0.02g a-amylase granules in 100cm³ distilled water Ø 0% ? no a-amylase granules, just 100cm³ distilled water These will be my six concentrations that I will use in my experiment. Safety: Avoid all contact of the concentration solutions of a-amylase with skin or eyes. Avoid ingestion or inhalation of the enzyme granules. In order to avoid all contact with eyes or skin, gloves, lab coat, and safety glasses will be used at all times while handling the enzyme concentration solutions. Spilled preparation should be removed immediately. Avoid formation of dust; in order to avoid the inhalation of the enzyme. Take up by mechanical means, preferably a vacuum cleaner equipped with a high efficiency filter. Flush the remainder carefully with water. Avoid splashing and high-pressure washing (i.e. avoid formations or aerosols). Ensure sufficient ventilation. Wash contaminated clothing. The cork borers being used to bore the holes in the agar are sharp. Therefore extra care must be taken when handling these cork borers, to ensure the safety of others around me, as well as myself. An iodine solution of 0.01M will be used to test for the presence of starch. If spilt on clothes or skin, brush off solid immediately. Flood the affected are with water immediately, or bathe with sodium thiosulphate solution. If blistering occurs on affected area, seek medical attention. If spilt in the laboratory, ventilate area of spill. Scoop as much solid as possible into sodium thiosulphate solution. Spread sodium thiosulphate solution over area of spill. Leave for an hour, and mop up and rinse the area of spill. If iodine solution comes into contact with eyes, flood the eye with running tap water for 10 minutes, and seek medical attention. To avoid getting the iodine solution on hands or in contact with the skin on the hands, gloves will be worn at all times. To avoid getting the iodine solution into contact with clothes, a lab coat will be worn when handling the iodine solution. To avoid getting iodine solution in eyes, safety glasses will be worn at all times. Diagrams: Pilot Test -???- Numerous pilot tests were carried out in order to test whether the constants and variables chosen above worked or not. The following will show the results of these pilot tests, and will show any changes I have made to my plan as a whole. From the very beginning, a very crucial problem had occurred. As I tried to make the enzyme concentration solution for my 0.1% solution, the enzyme was nowhere near fully dissolved in the 100cm³ of distilled water. This means that if I was to make up all of the solutions with this enzyme, not all the enzyme will have dissolved, which means I cannot ensure that the concentration solutions that have been made up are of the actual concentrations they are meant to be at. To overcome this problem, I decided to change my enzyme, as I thought this factor would affect my result too much for me to be able to get a decent set of results. I decided to change from detergent a-amylase Termamyl� 60T in granule form to non-detergent a-amylase Termamyl� 120L a liquid form of the a-amylase enzyme, which means no problems will occur while dissolving. This means, however, that I will not be able to relate my results to their effect on the practical uses of enzyme concentration in biological washing powders. Firstly, a test was done to find out which diameter of cork borer was best suitable to obtain results for the real experiment. Different sized wells have been bored using varying diameters of cork borers. In each of these wells, an a-amylase solution of concentration 0.1% (i.e. 0.1g in 100cm³ distilled water) has added, using a graduated pipette. The diameter of the cork borers used was measured using Vernier calipers. The starch agar plate was left in an incubator/oven at a temperature of 26�C over a period of 24 hours. When taken out, iodine solution was added to the agar plate, to give the clearance rings. The diameter of these clearance rings was also measured using Vernier calipers: Diameter of hole bored (measured using Vernier calipers) (mm) Diameter clearance of ring with 0.1% Termamyl� 60T (mm) 14.2 27.2 11.5 23.9 10.2 22.2 7.1 19.0 4.5 12.8 From these results I can conclude that the 14.2mm cork borer would be best suited for this experiment, as it will give me the largest difference in between results, which means they will be more easily compared. (Note: this experiment was still done before the enzyme change was decided.) Upon using a graduated pipette to fill the wells in this experiment, I also needed to find out the volume of concentration solution each different size well held. These volumes were measured using a graduated pipette. These results are shown below: Diameter of well (mm) Volume of enzyme solution held (cm³) 14.2 0.40 11.5 0.28 10.2 0.16 7.1 0.12 4.5 0.06 These results show that if I were to use the cork borer with the diameter of 14.2mm, I would need to fill each well with a 0.40cm³ volume of enzyme concentration solution, using a graduated pipette. A change in enzyme also meant that a suitable change in concentrations was necessary. In the following table, an experiment has been done to show the effect of different types of concentration using the a-amylase Termamyl� 120L enzyme. Different extremes of concentrations have been made up, to see what effect each had on the breakdown of the starch agar, in order to decide on final concentrations of the enzyme solutions. Concentration of 10%, 5%, 1%, 0.1% and 0% were used. These were made by the method shown below: Ø 10% ? 2 cm³ a-amylase into 18 cm³ distilled water Ø 5% ? 1 cm³ a-amylase into 19 cm³ distilled water Ø 1% ? 0.2 cm³ a-amylase into 19.8 cm³ distilled water Ø 0.1% ? 0.02 cm³ a-amylase into 19.98 cm³ distilled water The agar plates again have been left in the incubator/oven for a period of 24 hours at the same temperature of 26�C. A well diameter of 14.2mm was maintained for all the holes. The results are shown in the table below: Concentration of the enzyme solution (%) Diameter of clearance rings (mm) 10 39.2 5 35.6 1 30.0 0.1 27.7 0 0.0 These results show that if a good range of percentage concentrations is used, ranging from 10% to 0%, I will get a nice range of results. To ensure that there is a balanced set of results, I will go from 0% to 10% concentrations by going up in 2% jumps. This means that I will need to make new concentration solutions for each of these concentrations. The following shows how I will go about making these concentration solutions: Ø 10% ? 5 cm³ a-amylase in 45 cm³ distilled water Ø 8% ? 4 cm³ a-amylase in 46 cm³ distilled water Ø 6% ? 3 cm³ a-amylase in 47 cm³ distilled water Ø 4% ? 2 cm³ a-amylase in 48 cm³ distilled water Ø 2% ? 1 cm³ a-amylase in 49 cm³ distilled water Ø 0% ? 0 cm³ a-amylase, 50 cm³ distilled water A pilot test using these concentrations has been done, to test whether these concentrations would be suitable for the real experiment. This time, however, the agar plates were only left in the incubator/oven for 3:30 hours, as there was not sufficient time for me to, at that particular point in time, leave the agar plates in the incubator/oven for the full 24 hours. This is, however, not a major problem, as this experiment is just so that an idea is achieved of whether the concentrations are suitable or not. The table below shows the results obtained from this pilot test: Concentration of enzyme solution (%) Diameter of clearance rings (mm) 10 27.4 8 23.0 6 21.4 4 21.0 2 20.2 0 0.0 These results show that a good increase in the results is shows, as expected. However, to make the results for each concentration more distinct, I will leave the agar plates in the incubator/oven for a period of 24 hours for the real experiment. Updated apparatus: � Termamyl� 120L a-amylase, pure liquid form � Distilled water (used to make concentration solutions) � Beaker 100cm³ � Measuring cylinder, 50cm³ (accurate to 1cm³, �0.5cm³) � Starch agar plates, pre-prepared (filled to a depth of 6mm, using 1% starch agar) � Cork borer, � 14.2mm � Vernier calipers (accurate to 0.1mm, �0.05mm error) � Stirring rod � Iodine solution of concentration 0.01M (used to test for rate of breakdown of starch) � Graduated pipette 2cm³ (accurate to 0.02cm³, �0.01cm³) � Mounted needle (used to remove the centres of the holes bored in the starch agar plate) [image] Constants (including the ones mentioned before): The same graduated pipette will be used, to ensure the same measuring error is included on all volumes measured. If different pipettes were to be used, there may be a slightly different error of measurement on the pipette, which would give slightly different volumes of enzyme concentration solutions in each well. This inequality in volumes would mean the experiment is not wholly fair, which could cause some concern as to whether the results can be seen as true results or not. This volume will no doubt be so small that it could be considered as ineffective towards the final results. Final method: Measure out 49cm³ of water using a measuring cylinder, and transfer to a dry beaker of volume 100cm³. Measure 1cm³ a-amylase using a graduated pipette, and add these to the beaker with water. Use a stirring rod to stir the mixture until the enzyme and the distilled water are evenly distributed. This mixture will give a 2% enzyme concentration, as shown at the top of page 10. Repeat these steps, making the concentration solutions for each of the concentrations in the table at the top of page 10. Take the lid of a clean starch agar plate. On this lid draw with permanent marker pen three lines, so as the lid is evenly spaced into 6 equal sections. Place the lid on the bottom of the starch agar plate. Use a cork borer of diameter 14.2mm to bore three holes, one on every other line correlating to the lid grid (the lines drawn on the lid of the plate earlier on). Use a mounted needle to remove the centres of the holes bored. If done properly, what should be left is an agar plate with three wells equidistantly spaced apart on them. Repeat this step, using the same lid grid, to make a second starch agar plate, in order to make enough holes for all my concentration solutions. Use a graduated pipette and fill each well with 0.40cm³ of the corresponding enzyme concentration solution (as shown in the diagram below), so that there is one concentration of the enzyme solutions in each of the wells in the agar plates made. Place the lids of the plates back on to the dishes, and place the dishes in an oven/incubator at 26�C for a period of 24 hours. If the plates were left in a laboratory to react for 24 hours, the temperature would not be constant, and the temperature would be lower than 26�C. This temperature is slightly closer to the optimum temperature of the enzyme, which is 37�C, which means the enzyme will function better this way. The temperature will also remain at a constant. Once the 24 hours have passed, remove the agar plate from the oven/incubator. Add enough 0.01M iodine solution to the agar plate so that the agar has a thin layer of iodine solution. Wait until the iodine solution has reacted with the starch. In the presence of starch, iodine solution turns blue/black. However, if no starch is present, the iodine solution takes on a different colour. This means that around the well, where the enzyme has broken down the starch, there will be a ring of different coloured iodine solution, followed by the blue/black ring of iodine on the starch agar that has not been broken down yet. This ring is called the clearance ring. Measure the clearance ring made by the broken down starch agar with Vernier calipers. The diameter of the rings is measured, and these will then be used as a means of a value for the breakdown of starch by a-amylase. Note down these diameters in a result table such as the one below. Repeat the above steps two times. If measuring mistakes are made in the second run of the experiment, the results of the third run will be able to confirm whether results for the first run are correct or not. This is why two repeats are necessary: to make sure the results are reliable. Results: The results obtained from the experiment will need to be noted down in a result table, shown below: Experiment number Concentration of enzyme solution (%) Diameter of clearance ring (mm) Experiment 1 10 8 6 4 2 0 Experiment 2 10 8 6 4 2 0 Experiment 3 10 8 6 4 2 0 The possibility of a second table with the averages of the experiment could be made. A graph to present the results will also be drawn out (by hand), where the x-axis is the diameter of the clearance rings, and the y-axis is the increasing percentage concentrations. Analysis of Results Obtained After having carried out my practical, I obtained the results shown in the table below: Experiment number Concentration of enzyme solution (%) Diameter of clearance ring (mm) Experiment 1 10 37.7 8 32.4 * 6 32.8 * 4 34.4 * 2 30.3 0 0.0 Experiment 2 10 37.5 8 35.7 6 33.8 4 35.0 * 2 30.4 0 0 .0 Experiment 3 10 37.8 8 36.1 6 34.5 4 31.3 2 30.1 0 0.0 Experiment 4 10 39.2 8 37.5 6 35.9 4 34.8 2 32.3 0 0.0 Average of all results (excluding anomalous results) 10 38.2 8 36.5 6 34.7 4 33.1 2 31.0 0 0.0
= Anomalous results
As can be seen from the table, Experiment 1 obtains many anomalous results. This may be due to a very slight error in measurement, as it is very hard to measure out exactly 0.4cm³ of enzyme concentration solution, using a pipette and pipette fillers. Pipette fillers are used by turning a knob on the side of the filler. This knob has grooves in it, and works as a gear: when the knob is turned, a long cylindrical shaft, also with grooves in it, moves up, pulling in the air below it (as there is an air tight seal around the base of the pipette filler, where the pipette enters the filler). This causes the concentration solution to enter the pipette. However, in some cases in order to get a measurement of exactly 0.4 cm³, the knob of the pipette filler needed to be positioned so that one groove of the knob was at the midpoint of fitting exactly into a groove either side of it. This meant that the groove would automatically slip into either of the grooves next to it, which means that it was very hard to keep the bottom of the meniscus of the concentration solution in the pipette at the 0.4 cm³ mark. This may help to explain why several anomalous results were obtained in these experiments. As there were so many anomalous results in experiment 1, I decided that it would be better just to ignore these results all together, as they would significantly change my final average results. Another repeat, experiment 4, was done to provide a third set of results to help confirm the reliability of my results. Experiment 4 was done under the exact same conditions as the other three, using the same concentration solutions made up at the beginning of the experiment (the solutions were kept for several days. This may possibly have affected the enzyme?s ability to catalyse. The graph (Graph 1) however, does not suggest this at all, as the line for experiment 4 is the highest in comparison to the other two, and coincidentally also appeared to have the least amount of anomalous results), using agar plates that were from the same batch that was made at the beginning of the experiment (which means the agar inside the plates will be the same as the agar plates used in the first three experiments.), using the same cork borer that was used in the other three experiments (to make sure that the holes are of equivalent sizes), and incubating the plates at the same temperature, using the same incubator and incubating for the same period of time. Experiment 4 did give results that coincided with the results obtained in the other experiments. Notice that there is another anomalous result in experiment 2, for the 4% concentration. This will probably have been caused by the same reason as the other anomalous results, explained above. All anomalous results obtained in these experiments will be ignored in the final average of the results. For the three valid experiments I have constructed a graph (see Graph 1) with each experiment plotted as a separate graph. Experiment 2: There is a very clear anomalous result at 4% concentration as can bee seen from the large bulge in the graph on the graph (the gradient of the line increases steeply after 2%, and then decreases again after 4% before again maintaining a steady increase. This may well be caused by an error in measurement, as explained above. Neglecting the anomalous result, there is what appears to be a steady increase in the diameter of the clearance rings, as the concentration increases. This suggests that there is a fairly constant increase in the rate of breakdown of starch when increasing the concentration. A pattern is seen in this set of results, looking at the last the concentrations going from 6% to 8%, there is an increase of 1.9mm in the diameter of the clearance rings, and going from 8% to 10% there is a 1.8mm increase in the diameter of the clearance rings. This suggests there is an overall fairly constant increase in diameter of the clearance rings. I cannot, however, go as far as to say that due the fact that from 6% to 8% there is a 1.9mm increase in diameter and from 8% to 10% there is a 1.8mm increase in diameter the graph will level off, and that there would be a certain point where the concentration of the enzyme solution does not become the limiting factor anymore, mainly because due to the anomalous result I cannot tell whether the increase is leveling off or not, or whether it is a fluctuating increase. This would however seem to be a fairly logical conclusion to draw. An increase in the number of enzyme molecules means that comparatively, each enzyme molecule will have less starch molecules to break down. For example, say if initially the ration of enzyme:starch molecules was 1:20. Doubling the concentration of the enzyme solution means doubling the amount of enzyme molecules. This means the enzyme:starch molecule ratio has now become 2:20, or rather 1:10. The ratio has been halved. This means that now, each enzyme molecule will only be needing to break down half the number of starch molecules they were originally breaking down. Technically, this means that the rate at which these starch molecules are broken down will have doubled as well. This works the other way as well. If a starch:enzyme molecule ratio of 1:20 is established, halving the concentration of the enzyme means that now every enzyme molecule is breaking down twice as many starch molecules. This gives a starch:enzyme ratio of ½:20, or rather 1:40. Now, technically the rate at which these starch molecules are broken down will have halved. In other words, the enzyme molecules will take twice as long to break down the starch molecules, in comparison to the initial ratio. As I was increasing my concentration, the effect of this would basically be that the ratio is getting larger and larger, until the ratio enzyme:starch molecules ratio is at 1:1, and beyond that?s, at 2:1, 3:1 etc. This means that now, not every enzyme molecule actually has a starch molecule to break down. The effect of this is that if the enzyme:starch molecules ratio is 1:1, increasing the concentration past this stage will not actually change the rate at which the enzyme molecules are breaking down the starch molecules anymore, as each starch molecule will now have more than one enzyme molecule trying to break it down. As only one enzyme molecule will eventually break down the starch molecule, the rate of reaction will not change, and the concentration of the enzyme has now not become the limiting factor anymore. This relates closely to the Vmax theory. Vmax relates to increasing the concentration of the substrate, as opposed to the enzyme concentration. It states that since the rate of reaction depends on the rate at which enzyme-substrate complexes form, increasing the concentration will increase the rate of reaction, only up to a certain point, where all the enzyme molecules are in use, and no more starch molecules can be broken down at one time. This means the rate of reaction reaches a maximum velocity, and remains constant, and is known as Vmax. The same could be said for increasing the enzyme concentration. Increasing the enzyme concentration is effectively decreasing the substrate concentration. There will be a certain point in time where each enzyme molecule is breaking down a starch molecule, and increasing the enzyme concentration more will have no more effect. Each starch molecule is being broken down by one enzyme molecule, so therefore all the starch molecules are occupied. This means the rate of reaction will have reached its maximum velocity, and increasing the enzyme concentration more that this will not have any affect. This is effectively also a form of Vmax, except the other way around, with the starch molecules in abundance, instead of the enzyme molecules in abundance. Experiment 3: Looking at experiment 3, there is what appears to be another anomalous point. At the point at 4% concentration, the point looks like it is slightly lower than is expected, because if the point was slightly higher than it is now, one could draw what looked like a fairly straight line through all the points on the graph for experiment 3. This would then suggest that there is a steady increase in the diameter of the clearance rings as I increase the concentration of the enzyme solution. Ignoring the anomaly, and looking again at the last three points, going from 6% to 8% there is a 1.6mm increase in the diameter of the clearance rings, and going from 8% to 10%, there is a 1.7% increase in the diameter of the clearance rings. I could continue on from there and draw the conclusion that by taking into account the gradient of the graph at each point (which for this graph seems fairly constant, possible even slightly increasing), the graph would continue on at a constant increase as I increase the concentration of the enzyme solution. This would mean that (judging solely from my results) unlike experiment 2, there would be no point where the enzyme molecules do not become the limitors anymore, and that the increase in diameter of the clearance rings would continue until the concentration solution reaches 100%, when the concentration cannot go any higher. This, however, seems very unlikely, as like I said before; there must be a certain point in time when the number of substrate molecules becomes the limitors. Experiment 4: Looking at the results obtained in experiment 4, there do not appear to be any real anomalous results (though possibly the last point, at 10% concentration, as this point has a slightly larger gradient than that of the other points). Looking at the results in general, they appear to be slightly higher than the other two experiments. This may be due to the fact that experiment 4 was done on a different day than experiment 2 and 3. This means that the temperature of the lab may have been slightly different. The time period of incubation may not have been exactly the same. Technically, the agar plates themselves should not have been one of the problems, as the plates were from the same batch of starch agar plates as the plates used in experiment 2 and 3. Looking at the distance by which the clearance rings have increased in size: Concentration difference (%) Distance by which clearance rings have increased (mm) 2 to 4 1.7 4 to 6 1.6 6 to 8 1.1 8 to 10 2.5 The reason why I have not included the jump from 0% to 2% is that as this jump starts at 0% concentration, the breakdown of starch will react more to a smaller concentration change, and it is only after 2% that the diameter of the clearance rings start to increase proportionately to the increase in concentration. As can be seen from the table above, there is a fairly fluctuating increase in the diameter of the clearance rings. This makes it impossible for me to determine whether after 10% the diameter of the clearance rings will continue going up, making the logical assumption that the enzyme molecules stop being the limiting factor false, or whether the diameter of the clearance rings will level off, supporting the idea the enzyme molecules do not become the limiting factor anymore. There is a very high increase in the diameter of the clearance rings as the concentration solution is initially introduced (i.e. going from 0% to 2%). This means a small change in the concentration would have a large effect on the diameter of the clearance rings, (i.e. the breakdown of the starch. Comparing this to concentration changes after 2%, there is a very large difference in the gradients of the lines from 0% to 2% and 2% to 10%. This suggests that the initial change of concentration has a much larger effect on the overall starch broken down that the later change of concentrations (i.e. 2% onwards). It is hard to say whether the diameter of the clearance rings and the concentration of the enzyme solution increase proportionally. Going from 0% to 2%, the increase is not proportional at all, and the diameter of the clearance rings rises much faster than the concentration. After 2% concentration, however, the rise of the diameter of the clearance rings drastically slows down, and the increase in concentration is now slightly faster than the rate at which the diameter of the clearance rings increases. Average Results Adding all of the values for that concentration together, and dividing it by the number of values in each concentration obtained an average result of each concentration. These averages are also included in the large results table above (page 13). I have drawn a graph to show the average results of the investigation. This graph contains range bars for each result, to show the accuracy of the result. A range bar is made by plotting the highest value for that concentration and the lowest value for that concentration used in one average. These two points are then joined up by a straight line. This range bar will literally do what it says: show the range of results obtained for one concentration. This means that if the range bar is short, the average result is fairly accurate, ass all the values obtained are close together. If the range bar is long, however, the values obtained of that concentration are far apart, which means they will not be as accurate. Taking this information, and looking at my graph, all except the 4% value seem to have fairly small range bars. This is a good this, as a small range bar means there is not much difference between all the results, thereby making the average value slightly more accurate. The only concentration on my graph that has a larger range bar in comparison to the other range bars is the 4% concentration. This is due to the fact that one of the results for this concentration was an anomalous result, and was left out when calculating the average result. If there are fewer values to calculate an average result from, the average will not be as accurate. This is therefore why my 4% concentration average is not as accurate as the other results. A large range bar is not good, as this means that there is a large discrepancy in between different results. The average result will therefore not be as accurate as the averages with the shorter range bars. Looking at the difference in diameter of the clearance rings for each concentration: Concentration difference (%) Distance by which clearance rings have increased (mm) 2 to 4 1.7 4 to 6 1.8 6 to 8 1.6 8 to 10 2.1 As you can see from these results, the average results are also fluctuating, which means it is hard to say which way it is increasing (decreasing increase or increasing increase). By the looks of the results in the table, it appears as though the values are fluctuating, but getting closer and closer together. This means that there is a possibility that, judging from my average results, there is a certain concentration where the graph will become level, and the concentration of the enzyme solution will, as I have said before, not be the limiting factor. I do not, however, know this as a fact, so I cannot come to any valid conclusion on whether the graph levels off or whether the graph keeps on increasing. For the value of increase going from 0% to 2% concentration, this value cannot be used as a value for the increase. This is because I do not know where on the scale my value for the 1% concentration will be, and therefore cannot make a justified guess as to how fast the rate of breakdown is increasing. Looking at the increase from 2% to 4%, and 4% to 8%, if I double the concentration, the value for the increase in diameter of the clearance rings going from 2% to 4%, the value of increase going from 4% to 8% roughly doubles. Looking from the 2% value onwards, it is quite evident that there is an almost perfect line going though the points. This suggests that there is a fairly steady increase, which means that there must be a fairly constant increase in the rate of breakdown of the starch molecules. Notice that the points plotted on the graph are joined up using a straight line. There is a very logical reason for this. This reason is that I simply cannot guess what the diameter of the clearance rings is going be at concentrations in between 0% and 2%, 2% and 4%, 4% and 6%, 6% and 8%, and 8% and 10%. This means that I will therefore not be able to make an accurate judgment as to what the gradient of the curve is at points in-between the tested concentrations. I would therefore be wrong in drawing a curve to join up the plotted points. Also, I do not know whether the graph keeps on increasing, or whether the graph levels off. I would therefore be in the wrong place to make an assumption like that, and therefore would also not be able to draw a good curve to suit the experiment. This is why I have joined up the points with a straight line. Looking back at my hypothesis now: "As I increase the concentration solutions of the enzymes, the rate at which the enzyme breaks down the starch in the agar plate will increase", I can say that this hypothesis is supported by my results, and can therefore come to certain conclusions about the hypothesis (this is only true within the context of what I have investigated. Further investigations to try to further prove this hypothesis are discussed in the evaluation). The rate at which the starch in the agar plate is broken down does increase as the concentration of the enzyme solution is increased. This is due to the fact that as I increase the concentration of the enzyme solution, I increase the number of enzyme molecules in the solution. This means that there are more enzyme molecules to react with a constant number of substrate molecules, which means that the substrate starch molecules will get broken down much more quickly when there more enzyme molecules than when there are less enzyme molecules. This is why as the concentration of the enzyme solution is increased, the rate at which the starch agar in the plates is broken down increases too. Note: due to the fact that I had to change my enzyme from detergent a-amylase to non-detergent a-amylase, I will not actually be able to do what I intended at the beginning, which was to apply what I had done in this practical experiment to the washing powder industry. I would now have been in the right place to say that as the concentration of the enzyme is increased, the rate at which the starch is broken down also increases. This means that more enzymes in the washing powder would technically mean a better wash. however, I cannot yet determine whether there is a limit to the mass of enzyme that can be put in the washing powder, based on the idea that by increasing the number of enzyme molecules but keeping the number of starch molecules the same, there would be a point where increasing the concentration would no longer affect the rate at which the starch is broken down. This however, would require further practical research, which will be discussed later in the evaluation. Evaluation As the experiment has been done and repeated twice, there is experimentally justified evidence with which I can support my conclusion. This, however, does not mean that my results are completely accurate. There are some areas in this practical in which some errors leading to slightly modified results may have occurred. The majority of these factors were beyond my control, and nothing could have realistically been done to keep these factors at a constant. Looking at the starch agar plates firstly, there are already sever factors that may have affected my final results. Firstly, the cork borer that was used did not have a totally smooth cutting edge. Due to this the circular holes bored will have had jagged sides. These jagged sides ?protrusions? contained in the jagged surface will increase the overall area of the reacting surface, which would cause the starch to break down slightly more rapidly than expected. Even though the same cork borer was used, each hole will not have had an equal number of ?protrusions? in their reacting surface, which means there will have been a disagreement in the total reacting surface of each hole. Hence not all the concentration solutions will have had an equal surface area over which to react, implying already that the experiment may not have fully been a fair test, and possibly causing some of my results to be lower than expected, and some to be higher than expected. Focusing still on the holes bored in the starch agar plates, there is another area in which errors may have been present. The holes may not have been bored with the borer coming down at an exactly equal angle. Some of the holes may have been bored slightly more slanted than others. A hole bored slightly slanted, in comparison with a hole bored perpendicularly to the plate (the borer entering the agar at a 90Â? angle to the plate), will give a slightly larger reacting surface, again providing a disagreement in the surface are over which the enzyme is reacting, which. This may have caused errors in the final results. Theoretically, the total area of the reacting surface will be 267.66mm², using C=2Ï?r to find the length around the bottom of the circle (the circumference of the hole) and multiplying this by its height, to give: Total reacting surface area of single well=2Ï?rh Knowing that the diameter of the hole in 14.2, dividing this by 2 will give the radius of the circle. With this we can calculate the total reacting surface area, by putting this into the equation shown (the height of the well will be 6mm, as the agar is at a 6mm depth). This is assuming, however, that the reacting surface area is totally smooth, and assuming that the height of the agar remains the same all along the agar plate. As this is not the case, there will again be a slight contradiction to the fair testing that was intended. This may have been overcome by standing directly above the plate and boring the holes, though this would still not be a very accurate method of determining all the holes are bored at the same angle. Taking a look now at the actual plates, there are also a few sources of error that may have affected my final results. The plates were filled with a 1% starch suspension. Starch, at a concentration of 1% (1g of starch in 100ml of pure water) is sparingly soluble, and anything above that is not soluble at all. As I was using starch with a concentration of 1%, not all of the starch molecules will have been dissolved in the pure water (starch molecules are very large and thus very hard to dissolve). It would therefore be wrong to call the starch in pure water a starch solution, as it is not fully dissolved. It would more correctly be called a starch suspension. There will therefore always be some starch settling at the bottom of my beaker in which the starch agar is initially prepared. The same is true for the starch agar plates. As the starch suspension is in a jelly form, the starch agar will solidify after a certain period of time. Before this solidification occurs, however, the undissolved starch molecules will have settled at the bottom of the plate. This will cause an uneven distribution of the starch molecules around the agar plate, with more starch molecules being at the bottom of the plate. This will cause the starch molecules in the upper layers of the plate to break down more quickly than the starch molecules in the bottom of the plate. When the potassium iodide indicator is added to the agar jelly, there will therefore not be a single clearance ring where the enzyme will have stopped breaking down the starch. This introduces a complication in the measuring of the clearance rings. There will be a smaller clearance ring in the bottom of the agar jelly, and a larger ring in the upper layer of the agar jelly, making the overall clearance ring look blurred. I have measured the clearance rings using the ring in the upper layer of the agar jelly. Also, with some of the starch molecules being in solution, there is no guarantee that the starch molecules will be distributed around the agar jelly equally. There may therefore be a slightly larger number of starch molecules present in one area of the plate, and a slightly smaller number of starch molecules present in another area of the agar jelly. Depending on where the hole is bored in the agar jelly, some concentration solutions may have more starch molecules to break down than others, or vice versa, if comparing relatively to one another (i.e. how many starch molecules for each enzyme molecule in each concentration.). This leads to the conclusion that one concentration will relatively have a longer reaction time than others, which again shows fair testing may not have been carried out. There would, however, be no way to control this factor, as there is no physically way possible to control how many starch molecules go in each mm² of the starch agar jelly. The bottom surface of the actual plated did not appear to be flat. The base of the plates appeared to be curving upwards slightly. This causes a slight complication in the measurement of the depth of the agar that fills the plates, as due to the curved surface, the plate will not be filled at an even 6mm depth. The plates were filled to a 6mm mark on the side of the plate. But due to the curved base, the depth of the agar in the middle of the plate will not be equal to the depth of the plate around the sides of the plate. The plate is not actually filled to a depth of 6mm, but to an average of something just under 6mm. This creates another problem: as the starch is not equally leveled, some areas of the plate will have a different depth than others (i.e. around the outside, the depths will be deeper, and towards the centre, the depths will be shallower). Some of the holes bored will therefore have a slightly reduced reacting surface area than other holes, depending on where the hole is bored. This suggests that the reaction time will be slightly slower. However, a decrease in depth also means there are less starch molecules to break down. This suggests the overall reaction time will be faster. The two opposite factors may cancel each other out, and a valid result may be obtained. One variable may have more effect than the other. I am not in the right place to say which is true (as I do not know the total number of starch molecules in the shallow depths or deep depths, or whether the number of starch molecules is proportional to its depth, or whether the starch molecules are distributed evenly around the agar). I cannot come to a conclusion about which is true. However, I can make the point that in one way or another, it will have an effect on my final results. Taking this into consideration, a liability of unfair testing is present. As I mentioned earlier, some problems were encountered whilst measuring the volume of enzyme concentration solutions that were to be transferred to the holes bored in the agar plates. This was due to the fact that I had used a pipette filler to fill the graduated pipette, but then kept the pipette filler on the end of the pipette to transfer 0.40cm³ of the enzyme concentration solution into the bored holes. This made it very difficult to transfer an exact measurement of 0.40cm³. A much more accurate method of measuring out the 0.40cm³ would be to fill the graduated pipette with the pipette filler, past the 0.0cm³ mark. Quickly take the pipette filler off and place you right index finger on the top end of the pipette, making sure no solution is leaking out of the bottom end (due to lack of pressure). As there is much more control in the right index finger than the pipette filler, it will be much easier to control the volume of solution transferred (it is easier to let out very small volumes at a time using your finger than using the pipette filler). Enough solution would be run out of the pipette, so that the bottom of the meniscus of the fluid lies on the 0.0cm³ mark of the pipette. Hold the bottom end of the pipette above the allocated well, and very slowly let out 0.40cm³ of solution (by very lightly lifting up the right index finger so that the solution comes out very slowly), so that the bottom of the meniscus of the fluid now lies on the 0.40cm³ mark of the pipette. Repeat this for all the concentration solutions, using a clean pipette for each different concentration. If this method were to have been followed, there would have been a much smaller discrepancy in the measurement errors of the concentration solutions, resulting in much more accurate results. Keeping the number of enzyme molecules in mind, the same idea applies to both the number of starch molecules in the agar jelly and the number of enzyme molecule. Having enzyme concentration solutions, means having a certain number of enzyme molecules in solution. One may think that upon stirring the solution for a long period of time the enzyme molecules will get evenly distributed

How the Concentration of Enzymes Affects the Breakdown of Starch 8.6 of 10 on the basis of 1250 Review.