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How the Concentration of Enzymes Affects the Breakdown of Starch

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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 45°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, Æ 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 Æ5mm 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, Æ 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 90°
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=2πr 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 throughout the solution.
We cannot, however, be sure of this, or assume that this is the case.
If comparing volume to number of enzyme molecules not every area of
the solution will have the same number of enzyme molecules. If a small
volume of enzyme concentration solution is pipetted out, it may
therefore have a generally larger number of enzyme molecules in
comparison to the solution made up in the beaker (if comparing by
enzyme molecules/mm³). Or on the contrary, it may have less enzyme
molecules in comparison to the solution in the beaker. When a small
volume of enzyme concentration is pipetted out of the pipette and into
the well of the agar plates, again the same thing applies. The number
of enzyme molecules in the sample pipetted out of the pipette and into
the wells may be comparatively larger that the number of enzyme
molecules in the pipette. The number of enzyme molecules in the sample
of concentration solution pipetted out of the pipette may also be
comparatively smaler than the number of enzyme molecules in the
pipetter (if measuring by enzyme molecules/mm³). Yet again unfair
testing is a possibility. Similar to controling the number of starch
molecules in the starch agar, there is no way in which to control this
factor, so nothing can be done about it.

There is a possibity that not all the measurments made were fully
accurate. This is to be expected, as there is only a certain degree of
accuracy to which these experiments can be done. A possibility that
some, or many of my results are not completely correct arises. The
first accuracy problem occured when measuring out the small volumes of
enzyme needed to make the enzyme concentration solution. The type of
graduated pipette used was of capacity 2cm³, and was accurate to
0.02cm³. There was thus a ±0.01cm³ error in the readings that have
been taken. As I took two readings while using the pipette (one at the
beginning of measuring, and one at the end of measuring) there is a
total of ±0.02cm³ (±0.01 x 2 = ±0.02cm³) error in the readings made by
the graduated pipette.

Another accuracy problem was met when measuring with the measuring
cylinder used to measure out the volume of distilled water with which
to mix hte enzyme on order to make the concentration solutions. The
measuring cylinder was of cpacity 50cm³, and was accurate to the
nearest 1cm³, so there was an error of ±0.05cm³. As only one reading
was taken with the measuring cylinder, the total error in measurment
remains ±0.05cm³.

A third area where measuring errors may have occured was when
measuring with the digital Vernier calipers. The Vernier calipers read
accurate to 0.1mm, giving a measuring error of ±0.05mm. Again, as only
one measurement was taken, the error stays at ±0.05. As I have
explained before, the actual measurement of the diameters of the
clearance rings was difficult. The fact that the line where the
breaking down of the strach by the enzyme had stopped was very vague,
and there were multiple clearance rings layered on top of each other,
made it very hard to try to get accurate readings, as it was hard to
see where to start measuring from. This may also have contributed to
the errors in measurments and accuracy.

As mentioned before, there were numerous anomalous results obtained in
the experiments. There are a large number of sources in this
experiment where errors may have occured during the carrying out of
the practical, or where unfair testing may have occured, which helps
to explain why the anomalous results have been obtained. However, this
makes it very hard to limit the error to one or two particular sources
(this in itself is not so important, as the errors mentioned above are
all fairly logical, and are to be expected. They all help to give
reasons why the anomalous results were obtained in the first place).

I think it would be a wrong assumption to say that my results are
completely accurate, as there are so many anomalous results, and as
there is a fairly significant discrepancy between the repeats and the
original (in order for me to be able to say the results were accurate,
the graphs on Graph 1 needed to have been a lot closer together, have
obtained less anomalous results, and the range bars on Graph 2 would
have been significantly smaller than what they are now) and again,
also the fact that there are so many areas in which error may have
occurred. However, I am able to make a reasonable assumption as to the
reliability of my results. Looking at the graph containing the
separate graphs (Graph 1), each separate graph shows a generally
increasing trend. Comparing the three graphs, and ignoring any
anomalous results, they are comparatively very similar. They all show
the same trend, which suggests that if the results for each separate
experiment are comparatively very similar, the average results will be
fairly reliable. As far as the range is concerned, the graphs are
slightly spread apart, which suggests error/accuracy has played a part
in affecting these results. Taking everything into consideration, the
graphs are very similar, and I can therefore come to the conclusion
that my results are reliable.

There were a few limitations that hindered the investigation slightly.
The first one considered is pH: a-amylase functions best at a slightly
acidic pH, with an optimum pH of 5.6. I was not, however, able to
maintain this slightly acidic condition. This is due to the fact that
if I wanted to make the starch agar jelly slightly acidic, the
introduction of a pH buffer would have needed to be taken into
account. Upon adding a buffer, however, the complexity of the
investigation will be increased. It would have been very hard to
maintain a pH of 5.6, as the agar jelly would not have solidified as
well if the pH buffer was introduced. Also, it would have been hard to
introduce the buffer, yet getting a decent distribution of starch
molecules around the plate for the enzyme molecules to break down, to
give comparable results. I therefore did not introduce a pH buffer in
this experiment.

The next limitation to be considered is the temperature at which the
investigation was carried out. The optimum temperature of a-amylase is
37°C, yet the temperature at which the experiment was conducted at was
26°C in the incubator/oven. This is a fairly large gap in between the
optimum temperature and the temperature at which the experiment was
carried out. The reaction time at 26°C will therefore be much longer
than the reaction time at 37°C (this is why in order to get valid
results, the agar plates have to be left in the incubator/over for a
period of 24 hours). There is a very logical reason for this: at any
temperature above 28°C-30°C the agar jelly will melt, leaving the
experiment impossible to carry out at temperatures above 28°C. The
temperature inside the incubator/oven was a constant 26°C. The result
of this is that the agar plates need to be left in the incubator/oven
for a longer period of time, in order to achieve the same effect as
carrying out the experiment at the optimum temperature.

As the enzymes are left in the incubator/oven for such a long period
of time, there is also the time factor that needs to be taken into
consideration. Is it possible that enzymes denature with time? If so,
as the experiment progresses, the enzyme will become less and less
effective, therefore breaking down less and less of the starch
present. This implies an increase in the reaction time (i.e. more time
needed to break down an equal number of starch molecules). Also,
taking into consideration the fact that as the starch gets broken down
the enzyme needs to diffuse further through the agar jelly to get to
the starch that has not been broken down yet, again, as the experiment
progresses, the enzyme will get less and less effective (as less and
less of the enzyme molecules will diffuse though the agar jelly to get
to the starch molecules that have not been broken down).

One of the main features of this investigation that requires further
investigation is to obtain evidence to prove whether the graphs in the
graph level off or not (to find out whether there is a point in time
where the concentration of the enzyme will have no more effect on the
rate of breakdown of the starch molecules, and it does not become the
limiting factor of the experiment anymore). In order to continue this,
I will need to continue the procedure as has been done before, making
sure everything remains a fair test, and use higher enzyme
concentration solutions to determine whether the rate of breakdown of
the starch increases or decreases.

An altered method of this investigation would possibly have been more
suitable, and would have given slightly more accurate results. This is
a 'test tube' experiment. A small known volume of the enzyme
concentration solutions is placed in a boiling tube. A pH buffer of
pH5.6 is added, so as to maintain the enzymes optimum temperature.
Boiling tubes like this would be prepared for the concentrations used
in the previous experiment (i.e. 0%, 2%, 4%, 6%, 8%, 10%), and the
boiling tubes would be labeled with their corresponding
concentrations. These boiling tubes would be placed in a boiling tube
rack, and the test tube rack would be placed in a water bath at 37°C,
in order to maintain the optimum temperature of the enzyme. Upon
placing the rack of boiling tubes in the water bath, they will be left
submerged in the water bath for 10 minutes (keep track of time using a
stop clock or a stopwatch), in order to allow the enzyme to
equilibrate. Then, a known volume of starch suspension is added to
each of the boiling tubes, along with several drops of potassium
iodide indicator. Upon adding these two, the time is noted down at
which they were added to the boiling tubes (using the stopwatch or
stop clock). Once the indicator has turned colourless (which means all
of the starch in the boiling tubes has been broken down) the timer is
stopped, and the time taken to break down the starch is noted down in
a results table.

In several ways, this method is better that the method I had
previously chosen to do the experiment with. This method allows me to
keep both the pH and the temperature at an optimum (as the enzyme will
be at its optimum temperature and pH, the reaction time will be much
faster, in comparison to using the method with the agar plates, making
the process less time consuming as well.

However, each concentration will need to be done separately, and each
concentration will need at least two repeats (to make sure results are
similar), making the process still fairly time consuming. Also, the
end point at which all the starch has broken down is not very clear.
The end point will also be decided by the human eye, and human
interpretation, which is not a very accurate way of deciding the end
point. This will make the end point of the experiment much less
accurate that that of the experiment using the agar plates. In this
experiment, the time taken for a colour to disappear is being
measured, which is not an accurate measurement, whereas with the agar
plates, a certain distance is being measure, which is much easier to
measure to a good degree of accuracy, using measuring equipment (in
this case the Vernier calipers were used).

Bibliography:

1. Advanced Biology, Micheal Kent; OXFORD University Press

2. Nelson Advanced Science: Molecules and Cells, John Adds, Erica
Larkcom, Ruth Miller; Nelson

How to Cite this Page

MLA Citation:
"How the Concentration of Enzymes Affects the Breakdown of Starch." 123HelpMe.com. 18 Apr 2014
    <http://www.123HelpMe.com/view.asp?id=122465>.




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