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Will amylase inhibitors affect the colorigenic reaction between starch and iodine?

Will amylase inhibitors affect the colorigenic reaction between starch and iodine?


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I'm doing an experiment for my IB bio EE involving colorimetry. I'm not experienced at all with colorimetry, so I'm having some trouble planning it. The experiment is on enzyme kinetics, and I'm testing the effect of an inhibitor on the rate of digestion of starch by alpha-amylase. Currently, my plan is just to use a starch solution with an iodine indicator, add the amylase and inhibitor and measure the change in absorbance of a certain wavelength over time with the colorimeter, however, I'm worried that when I add the amylase and the inhibitor it'll affect the colour of the solution quite a lot. Will this be an issue, and if so is there a better method I could use to control for it?

Any help would be appreciated, this is my first time actually planning an experiment and I'm finding it difficult :)


To measure your amylase activity, you will monitor the disappearance of amylase's substrate, starch. Starch reacts with iodine (which is yellow) to form a blue compound (Amax 620 nm). This reaction is the basis of a colorimetric assay for amylase activity. You should be very careful about the volume of the final solutions so it won't effect your results. For example you will add inhibitor to one of them and you won't add that to the other but you should add water instead of the inhibitor so dansity won't be a problem. (final volumes should be same since starch amount at the beginning will be same) You can use some acid like HCL to stop enzyme activity so it won't change the colour and won't effect your results. After the acid stops the enzymatic reaction and the iodine reacts with the starch to produce the blue color. Any starch that has not yet been hydrolyzed by the amylase will turn blue color being proportional to the amount of starch remaining. The intensity of the blue color can be by measured by its absorbance at 620 nm. The greater the change in absorbance between a sample containing the initial amount of starch (without enzyme) and the hydrolyzed mixture containing the enzyme, the greater the amount of starch degraded by the enzyme, therefore the greater the activity of the enzyme being measured. Make sure you give the enzyme enough time and heat to catalyse the reaction. Sorry for grammar errors. I hope this was helpful!

Edit:you asked if inhibitor effects the experiment. I did say it above. If you use a colorless substance to denature or inhibit the enzyme it won't effect your results. If you can't do this you may add the inhibitor after the reaction is over so it won't effect the reaction but you may eliminate possible differences in color


Competitive and product inhibition-based α-amylase activity analysis method

To propose a methodology for analyzing salivary α-amylase activity (sAMY) for a hand-held device that can be used easily and quickly for evaluating human psychological effects.

Methods

An improved method for the analysis of sAMY is proposed using competitive and product inhibition in a dry-chemistry system with a reagent paper containing 2-chloro-4-nitrophenyl-4-O-β- d -galactopyranosylmaltoside.

Results

Not only the competitive (maltopentaose) inhibitor, but also the product (maltotriose) inhibitor significantly reduce the reaction speed of sAMY. There is an independent effect between the two inhibitors. The dynamic linear range of the analysis can be enhanced by 2.5 times larger by adding the competitive and product inhibitors simultaneously and by preparing their concentrations appropriately.

Conclusions

It is considered that the application of competitive and product inhibition can be effective from the viewpoint of enhancing analysis range and effectively reducing costs.


Factors Affecting the Activity of Catalase and Amylase Lab Answers

Proteins (or polypeptides) are organic macromolecules consisting of amino acid subunits (monomers) that are arranged in a specific order and folded into a specific shape. These monomers of amino acids are arranged in a specific sequence known as the primary structure which is determined by the DNA of the gene coding that protein.

Amino Acids consist of a centralized C atom attached to a unique “R” functional group which dictates which of the twenty possible amino acids found in the human body it is. All amino acids contain an amino terminal and carboxyl terminal, which play a significant role its ability to form peptide bonds with adjacent amino acids.

Amino acids can form peptide bonds with neighbouring amino acids through their amino and carboxyl terminals the condensation will remove a water molecule and construct stable polypeptide chains [1](see figure 2).

A protein’s shape and function are not only dictated by its primary structure of amino acid bonds, but also the weak intermolecular forces between the hydrogen atoms of the main amino chain the carbonyl groups (C=O) known as the secondary structure. Hydrogen bonding (an intermolecular force between H and O,N,F) results in two possible secondary structures alpha helix or beta sheet [2].

The alpha helix, right-handed coiled or spiral conformation will be springy and flexible whereas the beta-sheet will have high tensile strength due to hydrogen bonding. These segments of the protein can further fold and supercoil into tertiary structures, which are controlled by intermolecular forces (covalent, ionic, van der Waal) between the R-groups/ side chains of the amino acids.

Proteins often have molecular weights in the thousands to 100 thousand[3] larger proteins might be made up of several polypeptides. The interaction of two or more polypeptides to form a functional group is known as a quaternary structure forming a detailed globular shape for a very specific activity.

Proteins have many functions throughout the body such as:

  • Transport molecules (hemoglobin transports oxygen)
  • Enzymatic catalysts
  • Storage molecules (Ex: Iron is stored in the liver as a complex with the protein: Ferritin)
  • Movement (Ex: Proteins are the major component of muscles)
  • Mechanical support (Ex: Skin and bone contain collagen-a fibrous protein)
  • Mediating cell responses (Ex: Rhodopsin is a protein in the eye which is used for vision)
  • Antibody proteins are needed for immune protection
  • Cell differentiation uses proteins (Hormones)[4]

However, proteins can be affected by certain factors such as temperature and pH, known as denaturisation. This can destroy the primary structure of the protein rendering it unable to perform its original task and proving extremely detrimental to the cell.

A specific type of protein, known as an enzyme is responsible for catalyzing biological reactions by a factor of 10^6. They allow complex biological reactions to happen to safe temperatures to prevent protein denaturisation and water vaporization.

It lowers the amount of energy required for the reactants to form products (ex. correct angle, sufficient force) and facilitates a successful reaction. The alternate EA transition point the enzyme provides is known as the enzyme-substrate complex.[5]

An enzyme has a specific 3-D shape which is dictated by its primary structure and how the available “R” groups interact with each other. However, the enzyme dynamically changes its shape to better accommodate the substrate which is known as the induced-fit model. The specific area in which an enzyme reacts with its designated substrate is known as the active site.

The active site for an enzyme is specific to a substrate to ensure maximum efficiency in catalyzing it toward its products however, the enzyme is never consumed during the reaction. [6] The diagram below (figure 6) shows the steps in which the substrate binds to the active site, forms an enzyme-substrate complex, forms the product(s) and releases its.

The three-part experiment in this lab investigates the activity of two enzymes catalase and amylase. The first and second experiments will qualitatively/ quantitatively evaluate the activity of these enzymes. Hydrogen peroxide is naturally produced in organisms as a by-product of oxygen metabolism and needs to be broken down because high levels of it are extremely toxic. [7]

Amylase, a carbohydrase, catalyzes the hydrolysis of starch. The reaction can be written as: (C6H10O5)n + (n-1) H2O > nC6H12O6

The third reaction will analyze the enzyme activity rate through the decomposition of once again H22 into it products however this time, quantitatively.

The purpose of this experiment is to analyze the effect of multiple variables to determine if they have an effect on the rate at which catalase catalyzes H22 decomposition.

Such factors include the re-using of the enzyme, increasing its surface area, the effect of temperature or its concentration. These sources of catalase (i.e. liver and potato) will be compared to an inorganic catalyst (mangansese dioxide) to see the effect of each variable alteration.

The purpose of this experiment is to analyze the effect of pH on the rate in which amylase catalyzes the decomposition of starch into simpler monosaccharides. Through the use of a control and several buffers, this experiment will analyze quantitatively via Lugol Iodine indicator the effect of pH on enzyme structure and the optimum conditions for enzyme activity.

The purpose of this experiment is to analyze the effect of substrate concentration on the activity of catalase from yeast. Through the use of control and several dilutions of H22 solutions it will quantitatively determine the correlation between [ H22] and the rate at which catalase catalyzes the substrate.

Stagger time:

1) Devise an appropriate stagger time of about 30 seconds, so two buffers can be tested at the same time. (pH 3,5 and pH 7,9) two stop watches will be required.

2) Place the amylase into the first starch test tube and stir for several seconds once amylase is placed into the test tube, immediately begin the stop watch.

3) At the 30 seconds mark, place the amylase into the second test tube and stir for several seconds immediately begin the stop watch, once amylase is placed into the test tube.

4) Proceed with the experiment and dispense the solutions into their appropriate micro wells at each of their 1 minute intervals.

5) Repeat for the second pair of pH buffers being tested.

1) Obtain all necessary materials (3% H22 solutions, H22 dilutions, stop watch, filter paper, hole punch, 50mL beaker, 25mL graduated cylinder, yeast suspension)

2) Place filter paper in yeast suspension for 2 minutes.

3) Take filter paper and place to it at the bottom of the 25mL H22 solution.

4) Observe the time it takes for the paper to rise to the top of the 25mL H22 solution.

7) Repeat steps 2-6 for the 1:5, 1:10, and 1:50 dilutions.

To make the dilutions for each test, a simple calculation can be performed to determine the H22 solution need:

1:5 dilution: (25mL/ 5) = 5mL of 3% H22 solution and 20mL of water

1:10 dilution: (25mL/10) = 2.5mL of 3% H22 solution and 22.5mL of water

1:50 dilution: (25mL/ 50) = 0.5mL of 3% H22 solution and 24.5mL of water

Observations

Table 1.0: Qualitative Descriptions of the Relative reactions Rates of Catalase

Table 2.0: Starch Test Results with Lugol’s Iodine of Amylase at various pH (3,5,7,9)

pHTime (min)
123456789101112131415
3blue
5bluebrown
7bluebrown
9bluebrown

Table 3.0: Amylase Test Results with filtered paper in yeast suspension placed into H22 solutions

Results Summary

Error Analysis

Error: Part 1

Inaccurate liver sample sizes and finger contact

The first error that was problematic in ascertaining accurate results in part 1 was the inability to cut identical 1cm 3 pieces of liver. Despite the use of a ruler, the slippery texture and irregular shape of the liver sample made it impossible to cut matching pieces, therefore certain pieces were slightly larger or smaller than others.

This proved to be a dilemma because the control variable for several parts of the experiment was altered and therefore provided inaccurate data. Having a larger piece of liver would have given the liver greater surface area in which to react with the H22 solution and therefore could have created a larger column of oxygen bubbles having a smaller piece would have resulted in vice versa.

Also, any finger contact between the liver sample and the experimenters could have caused a transfer of oils from their fingers to the liver piece. The insoluble oils could have delayed or completely hindered the touched surfaces from catalyzing the decomposition of the H22 solution and once again provided inaccurate data.

Error: Part 2

Stagger time slightly off

This result is an entirely human error but is a problem because of the difficultly of properly managing the stagger time for this experiment. The stagger time selected by most groups was 30 seconds, giving ample time to mix the starch with amylase and then place it into its approximate micro well. However, if a group was off by 5-10 seconds, due to human inefficiency, this could have resulted in inaccurate data.

Though unlikely, being off by perhaps 10 seconds during your first stagger time would cause the timer to be off by 10 seconds each time they reach the 1 minute interval after 6 rounds of observations the time would be off by an entire minute. This may not be significant for smaller time variations (such as 1-3 seconds) but could be greatly problematic for yielding accurate values for when the amylase is functioning properly at perhaps pH 7 or 5.

Error: Part 3

Touching the paper is once again entirely human error but could have easily transpired because the tweezers made it extremely difficult to separate the filter paper into single sheets. Touching the filter paper would have resulted in the transfer of oils from the experimenters’ hands to the filter paper and coat it with a non-water soluble layer.

This could have delayed or completely hindered the touched surfaces from catalyzing the decomposition of the H22 solution, taking more time for the filter paper to raise to the top of the solution providing inaccurate results.

This was a very simple error, which could be easily corrected. However not stirring the yeast suspension, would have resulted in the yeast particles sinking to the bottom and this could alter the concentration of the yeast on the filter paper. By not stirring it, the filter paper was submerged in a lower concentration and by stirring it it would increase the potential concentration that could be absorbed by the filter paper.

This could create an erroneous control and provide inaccurate data. A lower concentration of the yeast would take longer to rise to the top less oxygen bubbles would be provided from catalyzing the decomposition of the H22 solution.

For the most part, the results for this experiment transpired as predicted. In the first test, with the inorganic catalyst (MnO2) and the sand, there was only a minimal reaction observed with the MnO2. The sand was inert and produced no reaction because it lacked any component to cause the H22 solution to decompose into its products the MnO2 produced only 0.5cm (table 1) of oxygen bubbles because though it is a catalyst for this reaction, it has minimum efficiency.

In the second test, both the potato and liver reacted with visible results due to them both possessing catalase. However the reaction with the liver produced a more vigorous reaction which can be attributed to it containing more catalase than the comparative potato slice. For test three, the addition of more liver to an already catalyzed reaction of H22 yielded no results, whereas the addition of more H22 solution did.

This is because adding more catalase to the reaction but not more substrate gives the enzyme nothing to react with and therefore no reaction is produced. However the addition of more H22 solution does produce a reaction because the catalase is never consumed by the reaction it catalyzes and therefore can continue to catalyze substrate as more is added.

Crushing both the liver and potato produced a faster reaction with longer oxygen columns than the cubic pieces because this increased the surface area of the catalase. This increase in surface area allowed a greater percentage of catalase to react with the H22 solution and produce a much more vigorous result. As for the variable of temperature, boiling the liver caused it to yield no reaction because it was denatured.

The excessive heat caused the catalase’s peptide bonds to break or re-arrange, altering its active site and rendering it unable to decompose the H22 solution. The liver at 37C had an optimum reaction because this is the average temperature of our body and thus the optimum temperature for this enzyme. The liver at 0C still produced visible results however the oxygen column they produced was smaller than the one produced when the catalase was at its optimal temperature.

Finally the increasing increments of the liver subsequently produced double the amount of oxygen bubbles for each sequential reaction. .5cm 3 yielded 4cm (table 1) of bubbles, doubling to 1cm 3 produced twice as many bubbles (8cm) (table 1) and 2.0cm 3 produced +14cm (table 1) of bubbles. By doubling the size of liver per reaction, it doubles the amount of catalase available to react, producing twice as many oxygen bubbles rapidly.

For this experiment, several different outcomes were recorded for the catalyzed decomposition of starch into simpler monosaccharides via amylase. At a pH of 3, there was no visible reaction observed in the micro wells, which is due to the acidity of the test tube environment. The functioning pH range for the enzyme alpha-amylase (which is found in saliva and the pancreas) is between 6.7- 7.0 with an optimum pH of 6.8 (slightly acidic).

At pH 3 (more than 1000 times more acidic) than its functioning pH range, the amylase denatured and was no longer able to bind with the substrate at its active site to catalyze the reaction. This is exactly what was observed, because during the full 15 minute observation, there was no colour change with the addition of Lugol- Iodine solution.

At the pH of 5 and 7, there were visible colour changes that transpired within the first 5 minutes (table 2). The pH 7 yielded visible results a full minute faster than the pH 5 because it was closer to the optimum pH of the enzyme. As seen in figure 14, the effectiveness of an enzyme is the greatest at its optimum pH and slowly degenerates as the surrounding pH shifts either lower or higher.

At pH 5, it was not acidic enough to completely denature the amylase structure but it did impede its ability to catalyze the reaction and therefore resulted in it taking a full minute longer to produce the light brown colour. At pH 7, it was obviously much closer to the optimum pH and therefore catalyzed the reaction at the fastest rate out of all 4 tests.

At the pH of 9 (basic) it took the amylase 9 minutes to produce the indication of starch being converted into simpler monosaccharides. It was expected that at this pH the amylase would have denatured, however a change in [OH] may have reduced the enzyme’s activity (figure 14) but it didn’t completely denature it and prevent it from catalyzing the reaction.

During part 3, the reaction transpired in accordance with the increase or reduction of H22 concentration within the 25mL beaker. As seen in the (graphic 1), the greater the dilution of H22 solution, the more time it took for the filter paper to reach the top of the beaker. At the controlled reaction of pure 3% H22 solution it took an average of 4.83 seconds to transpire, which was the fastest of all 4 solutions.

This can be expected because the amylase located on the filter paper from the yeast suspension will be able to catalyze the greatest % concentration of H22 solution. This was made evident with the experimental data because there was more substrate present for the amylase to catalyze and therefore more oxygen bubbles produced.

As the dilution factor increased, (1:5) (1:10) (1:50) so did the amount of time it took for the filter paper to reach the top of the suspension. As seen with the experimental data from table 2 or graphic 1, each dilution, subsequently reduced the amount concentration of H22 solution per 25mL and provided less substrate in which the amylase could catalyze.

This resulted in less H22 being catalyzed to its products and less oxygen bubbles being created to raise it to the surface.

[1] Transgalactic Ltd, Initials. (2005). What is Protein?. Retrieved from http://www.bionewsonline.com/5/what_is_protein.htm

[2] Campbell, NA and Reece JB (2005) Biology. Seventh Edition. Pearson Education, Inc

[3]G., Deleage. (1994). Lecture 1: secondary structure of proteins. Retrieved from http://www.chembio.uoguelph.ca/educmat/phy456/456lec01.htm

[4] Port, T. (2003). What is an Enzyme catalyst?. Retrieved from http://biochemistry.suite101.com/article.cfm/what_is_an_enzyme

[5] Di giuiseppe , M. (2003 ). Biology 12. Toronto, Ontartio: Nelson.

[6] Di giuiseppe , M. (2003 ). Biology 12. Toronto, Ontartio: Nelson.

[7] Williams, D. (2003, July 17). The many Benefits of hydrogen peroxide. Retrieved from http://educate-yourself.org/cancer/benefitsofhydrogenperozide17jul03.shtml

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Bacillus EXPRESSION: A GRAM-POSITIVE MODEL

Eugenio Ferrari , Brian Miller , in Gene Expression Systems , 1999

Amylase Promoter

The amylase promoter represents another class of growth phase and nutrition-regulated promoters of Bacillus. The amylase promoter of B. subtilis is turned on at the end of exponential growth and its expression is repressed by glucose ( de Vos et al., 1997 ). The amylase promoter is useful in the industrial expression of secreted proteins during the long stationary production phase ( Arbige et al., 1993 ). A number of vectors using the amylase promoter and amylase secretion signals have been described ( Behnke, 1992 ). Consideration must be given to the requirements for secretion from Bacillus, such as signal sequence structure and the proteolytic activities of the host Bacillus strain ( Wong, 1995 ). Proteins produced from amylase promoters in Bacillus have included various types derived from humans and other mammals ( Behnke, 1992 , Nakazawa et al., 1991 Novikov et al., 1990 ).


The New Standard Gallery

Boiling tube Safety hazards / precautions Enzymes – All enzymes are potential allergens so contact an inhalation should be reduced to a minimum. Can cause asthma, cause irritation to nose and eyes do not wallow also if spilt on skin wash off immediately . So gloves and goggles are necessary. And when disposed of it must be diluted in 10 liters of water.

Iodine solution – Vapor is harmful to eyes and when inhaled. Use in a fume cupboard and wear eye goggles. Should again be diluted before disposed of. Avoid contact Witt skin as stains can occur.Lead salts – toxic, do not swallow, avoid inhalation of dusts, use eye protection, dispose of by heavy dilution Justifications for method The equipment used is accurate enough for the experiment by way of measuring and rope adding. I choose the amounts and concentrations specifically so the numbers become easier to work with for example when working out the concentrations also it makes the test more fair if the changes in concentrations change equally each time. I added the Lead nitrate solution and the amylase at the same time so not giving extra time for the lead ions to work (if they have an inhibiting effect).

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And also to avoid starch being reacted before the lead ions are added so it is a fair test. The reason for boiling tube 6 and test tubes F and IF are purely for a controlled experiment to see the pure reaction of Starch solution with the Enzyme Amylase so comparisons can be made. The reason for use of Iodine solution is because of iodine solutions reactions when it comes in contact with starch as it turns Blue / Black. So we can see by darkness of the colour change how much Starch is left in the solution Prediction To make a prediction we need to know a little more about substances we are dealing with.So we can base a prediction on such information. Now we can look at the enzyme it self so we can get an idea of the enzyme structure.

Amylase is a protein, which acts a catalyst, and so is called an Enzyme. Enzymes act as catalysts for many different reactions throughout the body, this is because the heat or energy needed to perform the reaction is too high for the body to sustain so a catalyst is used to lower this energy which is called the activation energy. Amylase is a starch-degrading enzyme. Starch is a mixture of two types of glucose polymers.We can only absorb small molecules so the action of amylase during digestion is essential for survival. All organisms make amylase, which breaks the glucose chain in the centre *1 . We can also use this source to give a thorough explanation of the structure of Amylase *2 lo– The enzyme is a globetrotting. Its single polypeptide chain tot about 4 96 amino acid residues has two SSH (Slothfully) groups and four disulphide bonds and contains a tight bound Ca+.

. 20– Amylase, like many proteins and enzymes, mostly contains wide arrangements of a-helices and b-sheets.The secondary structures are held tightly together by hydrogen bonds. Hydrogen bonds between amino acid residues are especially stable in the hydrophobic interior of the protein, where water molecules do not enter ND therefore cannot compete for hydrogen bonding. 30– Amylase’s tertiary structure consists of one domain of connected a-helices and b- sheets. The sheets and helices are connected by an array of loops and turns of hydrophilic amino acid residues, mostly valise, praline, and glycogen because they produce an abrupt change in direction of the polypeptide chain.By the way of denaturing them changing their shape there fore the substrate will not be able to fit in to the active site.

What causes the Denaturing is the action of the heavy metal ions on specific groups of the protein enzyme itself which is described below Heavy metals, such as lead (BP) and mercury (Hug) exert their poisoning effect by binding to instable or slothfully groups of proteins, including vital enzymes. Once hey bind to an essential functional protein, such as an enzyme, they denature and/ or inhibit it. 8 The sulfured groups are mentioned in the primary structure of the Enzyme Amylase. The action of these bonding to the ions causes the denaturing of the enzyme. From all of this information we can quite accurately make a prediction that the lead ions will more than likely have an inhibitory effect on the Amylase and the effect will be irreversible. As a result from the test I then believe that the higher the concentration of the lead nitrate solution the longer it takes for the enzymes to fully exact the starch solution.I also believe that the detect tot the lead ions (time taken tort starch to react) is proportional to the concentration I. E.

If we double the concentration the time taken will approximately double also. So I would expect a graph of time taken for starch to fully react plotted against concentration of lead ions to look like this: c It would not start from zero because even without an inhibitor it will take some time to react the starch. Also we are able to say from the sources that it will be non-competitive I.


Assay of Salivary Amylase activity

To determine activity of Amylase enzyme in Saliva

Principle:

Amylase is the hydrolytic enzyme that breaks down many polysaccharides like Starch, Amylose, dextrins, and yields a disaccharide i.e., Maltose.

Reagents:

  1. Substrate (Starch): Mix 1 gm of soluble starch in 200ml of 0.1M Phosphate buffer (pH 6.8) boil for 3 minutes and cool to room temperature and filter it necessary.
  2. Enzyme: Saliva is the best and easily available source of amylase. Collect some saliva in a beaker and dilute it to 1:20 dilution with distilled water.
  3. 1% Sodium chloride: It is necessary for enzyme activity
  4. DNS (Dinitro Salicylic acid): Dissolve 1.6 gm of NaOH in 20ml of distilled water. Take 1gm of 3,5 DNS in NaOH solution. In other beaker take 30gm of Sodium potassium tartrate. Dissolve in 50ml of distilled water. Mix this DNS solution and finally make the volume up to 100ml with distilled water.
  5. Standard solution of Maltose: It is prepared by dissolving 200mg Maltose in 100ml of water (2mg / 1ml).

Procedure:

  • Take 0.5ml of substrate and 0.2ml of 1% NaCl in a test tube and pre-incubated at 37 0 C for 10 minutes then add 0.3ml of diluted saliva and incubate for 15 minutes at 37 0 C.
  • Stop the reaction by addition of 1 ml of DNS reagent mix well and keep the test tubes in a boiling water bath for 10 minutes.
  • Cool and dilute with 10ml of distilled water.
  • Read the color developed at 520 nm. Simultaneously setup the color developed at 520nm.
  • Simultaneously setup the blank as per the test by adding DNS prior to the addition of enzyme simultaneously.
  • Set up the standards of different test tubes and repeat the experiment as per the test and measure the color developed at 520nm absorbance.

Preparation of Phosphate buffer:

  • Dissolve 0.2M (2.7218 grams) of KH2PO4 in 100ml of distilled water to this solution add 0.5M (2.8053 grams) KOH drop by drop till the pH is set to 6.8.
  • Then make it to 200ml with distilled water. So the final concentration is 0.1M of 200ml Phosphate buffer.

Result:

The amount of Maltose in the given unknown sample is _________ grams of Maltose formed per 100ml of enzyme per one hour.

Calculation:

  • 1.5 mg of Maltose formed / 0.3. ml / 15 minutes
  • 1.5 X 4 mg of Maltose formed / 0.3 ml of Enzyme / 1 hour
  • 1.5 X 4 X 3.3 mg of Maltose formed / 1ml of Enzyme / 1 hour
  • 1.5 X 4 X 3.3 X 100 mg of Maltose formed / 100ml of Enzyme / 1 hour

Get this protocol in PDF format. Just download this “Color Reactions of Carbohydrates” file and make a print and distribute to the students. It helps you to protect your students from spelling mistakes and volumetric errors. All the best


Beyond the Enzyme and Substrate: Interactions of Starch and Non-α-Amylase Macromolecules

Proteins

Protein alone does not contribute considerably to flour paste viscosity. However, proteins can affect flour paste viscosity through their interactions with starch, and such interactions are related to both storage proteins (that is, gluten) and starch granule associated proteins. Wheat contains 10% to 12% (w/dry wt) storage proteins, which can form a gluten matrix through the interchain disulfide bonds between glutenin and gliadin (Shewry & Halford, 2002 Shewry, Tatham, Barro, Barcelo, & Lazzeri, 1995 ). The gluten matrix becomes a physical barrier and limits the accessibility of starch to water, which is a critical need for starch gelatinization. Consequently, the existence of storage proteins results in a delay of starch gelatinization and the time it takes to reach the highest paste viscosity (Chen, Deng, Wu, Tian, & Xie, 2010 Jekle, Muhlberger, & Becker, 2016 ). Chen et al. ( 2010 ) also reported that mixing gluten with starch decreased the peak viscosity of the reconstituted flour. Wheat starch granules contain about 0.2 % (w/w) protein, which is referred to as starch granule associated protein, and it can be located in the interior or on the surface of starch granules (Rahman et al., 1995 ). The existence of starch granule associated proteins negatively correlates to starch paste viscosity. Li et al. ( 2016 ) removed starch granule associated protein, using an alkaline solution that greatly increased the peak, setback, and final viscosity of the starch paste and decreased pasting temperature and peak time during the RVA measurement. Removing those starch granule associated proteins facilitates amylose leaching from starch granules and promotes starch granule swelling, which leads to a higher pasting viscosity (Li et al., 2016 ).

Nonstarch polysaccharides

Wheat flour contains 2% to 3% (w/dry wt) nonstarch polysaccharides, which can affect the leaching of starch molecules from starch granules during gelatinization thus, nonstarch polysaccharides can have an effect on starch granule swelling and starch paste viscosity (Sasaki, Yasui, & Matsuki, 2000 Shi & Bemiller, 2002 ). Arabinoxylan is the major nonstarch polysaccharide in wheat flour, and 1 gram of dry water-soluble arabinoxylan can hold ten grams of water (Kweon, Slade, Levine, & Gannon, 2014 ). Adding rye arabinoxylan to wheat flour shortens the pasting time (Harasztos et al., 2016 ), and adding a high concentration (3% w/w) of nonstarch wheat polysaccharides to wheat starch decreases starch peak viscosity and increases breakdown (Sasaki et al., 2000 ). Wheat arabinoxylan structural characteristics, including molecular weight, the ratio of xylose to arabinose, and side-chain substitution and distribution, vary among cultivars (Izydorczyk & Biliaderis, 1993 Saulnier, Sado, Branlard, Charmet, & Guillon, 2007 ) and are affected by the planting temperature. The arabinoxylan in winter wheat can prevent ice formation in cold weather, and a decrease in the ratio of xylose to arabinose increases solubility (Kindel, Liao, Liske, & Olien, 1989 ). The effect of arabinoxylan on FN has not been reported. In our previous report, it has been observed that the quantity of water-soluble arabinoxylan increased in some low FN wheat (Shao, 2018 ). More studies are needed to elucidate the role of arabinoxylan in wholemeal paste viscosity.

Lipids


RESULTS AND LABORATORY REPORTS

Previous Tests of Starch Concentration Using Saliva Samples

The starch concentration was tested in advance by the instructors to establish the optimal conditions for the qualitative analysis of saliva samples. Saliva obtained from the instructors (1–2 mL) was used. Three starch concentrations (0.5, 1, and 1.5%) were checked. The plates were incubated at 4, 40, or 60°C for 48 h. The results in Fig. 3 a show the appearance of larger starch-degradation halos in the 0.5% starch plate compared to the other plates (1 and 1.5%). In addition, in the 0.5% starch plate, the halos were observed at lower saliva dilutions than in the other two plates.

(a) Lugol test using saliva samples to determine the most suitable starch concentration. (b) Representation of the amylase activity (determined using the Phadebas test) in saliva at different temperatures.

These preliminary results suggest that the starch concentration is an important parameter for the sensitivity level of the qualitative assay in terms of the AAMY detection capacity. We hypothesize that this could be related to the level of agar fluidity due to the starch concentration, making the 0.5% starch media more suitable in terms of the optimal dispersion of the enzyme through the plate and the minimal substrate concentration required.

Quantitative AAMY Analysis

Three currently available laundry detergents were studied, and only detergents A and C had AAMY activity (Fig. 4 a). Detergent C exhibited higher AAMY activity than detergent A. Of the temperatures tested, the optimal temperature for AAMY activity in detergent A was 60°C, but the best temperature for detergent C was 40°C.

(a) Representation of the amylase activity (determined using the Phadebas test) in different detergents at different temperatures. (b) Results of the Lugol test (qualitative test) at different temperatures and incubation times.

Although detergent C has its highest activity at 40°C, it also has significant activity at 60°C, reinforcing the concept that industrial enzymes are thermostable.

We also studied the saliva AAMY activity quantitatively (Fig. 3 b). We observed that the optimal temperature for ptyalin was 40°C, and no activity was observed at either 4 or 60°C. It is noteworthy that the absolute amylase activity was nearly 18 times higher in the saliva samples (ptyalin at 40°C) than in the detergents (detergent C at 40°C), but the human enzyme was less resistant to significant temperature changes.

Qualitative AAMY Analysis

The qualitative analysis (Lugol test) was performed using the same three detergents (A, B, and C) and three temperatures (4, 40, and 60°C) that were used in the quantitative analysis. Although we had already determined that the optimal temperature for ptyalin was 40°C, saliva samples were used as positive controls in all of the assays. This control allowed us to confirm the specificity of the negative and positive results for the detergent amylases.

We tested the qualitative assay using two different incubation times (24 and 48 h). Using this methodology, we were able to obtain AAMY activity halos only with detergent C after 48 h of incubation and, surprisingly, only at 60°C. This activity was seen as specific degradation halos that decreased during detergent dilution detergents A and B had no AAMY activity at any temperature (Fig. 4 b).

The qualitative test was also performed on the saliva samples. It was clear that, of the tested temperatures, the optimal temperature for the AAMY in saliva samples was 40°C, which is consistent with the physiological context of the enzyme (Fig. 3 a).

Correlation Between Qualitative and Quantitative Analysis

The quantitative test (Phadebas) appears to be more sensitive than the qualitative test (Lugol). We were able to observe AAMY activity at 40 and 60°C in detergent A with the quantitative method but not with the qualitative method. In addition, detergent C showed activity at both 40 and 60°C using the Phadebas test however, when the Lugol test was used, we observed clear starch-degradation halos only at 60°C (with slightly transparent halos at 40°C). Initially, we believed this result to be incorrect (e.g. due to improper manipulation or poor assay development), but we (students and teachers) obtained the same final result in all of the replicates performed. As a consequence, a result initially believed to be an error actually provided hypothetical information about the final structures of two enzymes with the same activity. As later discussed, this observation could be due to a different enzyme presentation or configuration of the amylase from detergent compared with that of ptyalin (which was clearly positive in this assay).

However, this effect (i.e. no restriction due to hypothetical rigidity changes) was not evident in the quantitative assay, possibly because the liquid medium does not present the same hypothetical accessibility restrictions as the solid medium. The results of the quantitative and qualitative tests of the saliva samples were absolutely concordant.

PH Influence on the Activity of AAMY

We studied the influence of pH on the activity of AAMY in detergent C at both 40 and 60°C because we observed significant enzymatic activity and, unexpectedly, better starch-degradation halos at 60°C in the qualitative test. This observation facilitated the correlation between the quantitative and qualitative assays.

Assays of the saliva samples were incubated at 40°C, which was the optimal temperature of those tested.

In the quantitative assay of detergent C (Figs. 5 a and 5 d), a similar effect of pH was observed at both tested temperatures. The amylase activity was higher at pH 7, and there was a slight decrease in activity when the pH was increased (i.e. when the solution was made more alkaline). When a more acidic pH was used (pH 5), the activity was significantly affected compared with that at pH 7. This observation reinforces the concept that laundry detergent enzymes must be adapted to washing conditions with high alkalinity. The results of the qualitative assay for detergent C followed the same profile as previously observed: there were no clear starch-degradation halos at 40°C (no evaluable results), but apparent degradation halos were observed at 60°C. At this temperature, the clear differences obtained using the quantitative assay were less evident in the qualitative test, in which only slight differences in the halo diameters were observed between different pHs (Figs. 5 b– 5 c and 5 e– 5 f).

(a–c) Representation of detergent C amylase activity in solutions of different pHs at 40°C. (a) Results of the quantitative test. (b) Image of the qualitative test. (c) The measurement of halos in the qualitative test was not possible. (d–f) Representation of detergent C amylase activity in solutions of different pHs at 60°C. (d) Results of the quantitative test. (e) Image of the qualitative test. (f) Measurement of halos in the qualitative test. (g–i) Representation of saliva amylase activity in solutions of different pHs at 40°C. (g) Results of the quantitative test. (h) Image of the qualitative test. (i) Measurement of halos in the qualitative test. (j–l) Representation of detergent C amylase activity in solutions of different oxidant/surfactant agents at 60°C. (j) Results of quantitative test. (k) Image of the qualitative test. (l) Measurement of halos in the qualitative test.

In contrast, the quantitative test showed that the optimal pH for ptyalin activity at 40°C (Fig. 5 g) was between 7 and 8. The results of the qualitative test suggested that, of the pH values tested, the optimal pH was between 5 and 7, and a pH of 9 slightly affected ptyalin starch degradation activity (Figs. 5 h– 5 i). Although the differences obtained from the quantitative test were generally less evident, these data, when taken together, demonstrate that the optimal pH for ptyalin is approximately 7.

It is important to remember that the absolute activity of the detergent enzyme is nearly 18 times lower than that of the saliva enzyme. This fact is also evident in the qualitative assays, in which clearer degradation halos were obtained when saliva was used as the source of AAMY. This fact could explain the poor correlation between the quantitative and qualitative AAMY detergent assays in the pH influence study and could suggest that, in this specific case, a higher amylase activity would provide more comparable results with those obtained using the qualitative test.

Influence of Surfactants and Oxidizing Agents on the Activity of Detergent AAMY

The stability of the amylase enzyme from detergent C was assessed by exposing this enzyme to different surfactants and oxidizing agents. This assay also used both quantitative and qualitative methodologies.

Triton X-100 (1 and 5%) did not seem to affect AAMY activity. In contrast, the higher concentration (1%) of SDS decreased the enzymatic activity in both the quantitative and the qualitative assays. The oxidizing agent hydrogen peroxide strongly affected the AAMY activity we observed a low activity in the qualitative analysis, and we were not able to see halos in the qualitative analysis.

In this case, we were able to obtain a good correlation between the quantitative and qualitative tests (Figs. 5 j– 5 l).


Abstract

Glycemic indexes of bread made from mixtures of wheat flour and buckwheat flour are lower than those made from wheat flour. To discuss the mechanism of the buckwheat flour-dependent decrease in glycemic indexes, the formation of a starch−iodine complex and amylase-catalyzed digestion of starch were studied using buckwheat flour itself and buckwheat flour from which fatty acids, rutin, and proanthocyanidins including flavan-3-ols had been extracted. Absorbance due to the formation of a starch−iodine complex was larger in extracted than control flour, and starch in extracted flour was more susceptible to pancreatin-induced digestion than starch in control flour. Fatty acids, which were found in the buckwheat flour extract, bound to amylose in the extracted flour, inhibiting its digestion by pancreatin. Rutin and epicatechin-dimethylgallate, which were also found in the extract, bound to both amylose and amylopectin in the extracted flour, inhibiting their digestion induced by pancreatin. We discussed from these results that the lower glycemic indexes of bread made from mixtures of wheat flour and buckwheat flour were due to binding of fatty acids, rutin, and epicatechin-dimethylgallate, which were contained in buckwheat flour, to wheat flour starch.


Will amylase inhibitors affect the colorigenic reaction between starch and iodine? - Biology

a Describe the effect of pH on this enzyme.
b Explain why pH affects the activity of the enzyme.

7 The graph below shows the effect of temperature on the rate of reaction of an enzyme.

a What is indicated by X?
b What temperature would X be for a mammalian enzyme?
c Explain what is happening in region A.
d Explain what is happening in region B.
e Enzymes are effective because they lower the activation energy of the reactions they catalyse. Explain what is meant by 'activation energy'.

8 The reaction below occurs during aerobic respiration. The reaction is catalysed by the enzyme succinic dehydrogenase.

c Heavy metals such as lead and mercury bind permanently to -SH groups of amino acids present in enzymes. These -SH groups could be in the active site or elsewhere in the enzyme.

i Name the amino acid which contains -SH groups.[1]
ii Explain the function of -SH groups in proteins and why binding of heavy metals to these groups would inhibit the activity of an enzyme.[4]
iii What type of inhibition would be caused by the heavy metals?[1]

9. You are provided with three solutions: A, B and C. One solution contains the enzyme amylase, one contains starch andone contains glucose. Starch is the substrate of the enzyme. The product is the sugar maltose. You are provided withonly one reagent, Benedict's solution, and the usual laboratory apparatus.
a Outline the procedure you would follow to identify the three solutions.[6]
b What type of reaction is catalysed by the enzyme? [1]

10 The activity of the enzyme amylase can be measured at a particular temperature by placing a sample into a Petri dish containing starch-agar ('a starch-agar plate'). Starch-agar is a jelly containing starch. One or more 'wells' (small holes) arecut in the agar jelly with a cork borer, and a sample of the enzyme is placed in each well. The enzyme molecules then diffuse through the agar and gradually digest any starch in their path. At the end of the experiment, iodine in potassium iodide solution is poured over the plate. Most of the plate will turn blue-black as iodine reacts with starch, but a clear 'halo' (circle) will be seen around the well where starch has been digested. Measuring the size of the halo cangive an indication of the activity of the enzyme.

A student decided to investigate the rate at which a mammalian amylase is denatured at 60°C. She heated different amples of the enzyme in a water bath at 60°C for 0, 1, 5, 10 and 30 minutes. She then allowed the samples to cool downto room temperature and placed samples of equal volume in the wells of five starch-agar plates, one plate for eachheating period. She then incubated the plates in an oven at 40°C for 24 hours.

The results of the student's experiment are shown on the next page. A diagram of one dish is shown, and the real size of one halo from each dish is also shown.

a Why did the student cut four wells in each dish rather than just one? [1]
b One dish contained samples from amylase which was not heated (time zero). This is a control dish. Explain the purpose of this control. [1]
c Explain why the starch-agar plates were incubated at 40°C and not room temperature. [1]
d Describe what was happening in the dishes during the 24 hours of incubation [4]
e Why was it important to add the same volume of amylase solution to each well? [1]
f Measure the diameter in mm of the representative halo from each dish. Record the results in a suitable table. [4]
g Only one halo from each dish is shown in the diagrams. In practice there was some variation in the diameters of the four halos in each dish. How would you allow for this when processing your data? [1]
h Plot a graph to show the effect of length of time at 60°C on the activity of the enzyme. [5]
i Describe and explain your results. [4]
j Another student discovered that amylases from fungi and bacteria are more resistant to high temperatures than mmalian amylases. Using starch-agar plates as a method for measuring the activity of an amylase at 40°C, outline an experiment that the student could perform to discover which amylase is most resistant to heat. Note that temperatures up to 120 °C can be obtained by using an autoclave (pressure cooker).
k Enzymes are used in many industrial processes where resistance to high temperatures is an advantage.[5]
State three other variables apart from temperature which should be controlled in an industrial process involving enzymes.[3]

Answer for end-of-chapter

1 B
2 D
3 D
4 C
5 straight line drawn from origin at zero to show steepest gradient of curve
6 a maximum activity/optimum pH, is pH 5 activity gradually increases between pH 2 and
pH 5, and decreases from pH 5 to pH 10 activity very low at pH 2 and pH 10
b pH is a measure of the hydrogen ion concentration hydrogen ions are positively charged
hydrogen ions can interact with the R groups of amino acids affects ionic bonding/affects ionisation of R groups affects tertiary structure/affects 3D shape of enzyme therefore substrate may not fit active site (as precisely)

7 a optimum temperature
b 37 °C accept 40 °C
c as temperature increases the kinetic energy of the molecules increases the rate of collision between substrate and, enzyme/active site, increases rate of reaction increases
d the enzyme is gradually being denatured when the rate is zero the enzyme is completely
denatured enzyme loses tertiary structure substrate no longer fits into active site/active site
loses its (specific) shape so substrate does not fit hydrogen bonds broken/increased vibration of
enzyme molecule
e the extra energy which must be given to the substrate before it can be converted into the product

Exam-style questions


10 a replication increases reliability AW [1]
b to act as a reference to show what happens if there is no denaturation AW [1]
c 40 °C is the optimum temperature for a mammalian enzyme [1]
d enzyme/amylase (molecules) diff use(s) from wells into the agar
enzyme/amylase digests the starch
to maltose
forms rings/halos, of digested starch around the wells
amount of digestion/rate of digestion, is related to degree of denaturation of enzyme/amylase[max. 4]
e the more enzyme/amylase added, the greater the amount of digestion of starch
or
want results to be due to diff erences in preheating times, not to diff erences in amount of amylase/enzyme AW [1]

x-axis (horizontal axis) is labelled ‘Time (heated) at 60 °C’, y-axis (vertical axis) is labelled ‘Diameter’
units given on axes, min/minutes and mm
regular intervals on both axes (check that 0, 1, 5, 10, 30 are not regularly spaced on x-axis)
points plotted accurately
points joined with straight lines or smooth curve [5]
i enzyme was completely denatured after 30 minutes
rate of denaturation was rapid at first and then gradually slowed down
data quoted
enzyme loses tertiary structure
substrate no longer fi ts into active site/active site loses its (specific) shape so substrate does not fit
AVP e.g. hydrogen bonds broken/increased vibration of enzyme molecule [max. 4]
j heat samples of mammalian, fungal and bacterial amylases at diff erent temperatures
suitable range, e.g. between 40 °C and 120 °C

40 °C is a control (for reference to fi nd out size of halo with no denaturation)
at least five temperatures, e.g. 40, 60, 80, 100, 120 °C
heat for suitable length of time (e.g. one hour, at least ten min)
cool to room temp/40 °C, add equal volumes to wells in starch–agar plates, replicate wells in each
plate (e.g. four), leave 24 hours, test for starch, measure diameters of halos [max. 5]

Background information: amylase enzymes from the bacterium Bacillus licheniformis and
the fungus Aspergillus have been developed by biotechnology companies for use in industrial
processes. For example, a bacterial amylase that functions in the range 90� °C has been
developed and is used in beer brewing and other processes, and a fungal amylase that operates in
the range 50󈞨 °C is used for pastry baking and maltose syrup production.

k pH
substrate concentration
enzyme concentration [3]


Watch the video: Iodine Clock. Υλικά και περιγραφή.!! (February 2023).