Spectrophotometric Assay for Lipase Activity
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Published: Fri, 25 May 2018
Decomposition of human and animals bodies depends on numbers of factors. One of these factors is the presence of bacteria, both endogenous and exogenous of the body. They use the environmental factors to drive the decomposition of the tissues in the body. The various tissues are degraded at different rates by different bacterial cells. As it was seen in the model burial of a pig that is the early stages of decomposition Gram negative bacterial were mostly present in the decaying body. But after 6 – 7 weeks later the Gram negative bacteria started to decrease as the number of Gram positive bacteria present in the decaying body started to increase.
The bacteria produce enzymes which break down any tissue in the body. In the adipose tissue bacteria produces lipases which is secreted in to he tissue and slowly starts to break down the fat.
Lipases producing bacterial has been collected from a model burial environment without any environmental factors to see if there is a difference in the activity of the lipase enzyme which are produced by different bacteria species. These bacteria were used in two of the spectrophotometric assay that has been described in the literature. The turbidity assay shows how quickly the lipase enzyme can break down the lipid in the emulsion solution. On the other hand the BALB (dimercaprol Tributyrate) – DTNB (5, 5′- dithiobis (2-nitrobenzoic acid)) method shows the increase in the product that is produced by the lipase.
Lipases are found naturally as it is produced by plants, animals and micro-organisms. In the last few decades, the micro-organism production of lipases has been studied for commercial use, which leads to bacterial lipases being studied a great deal. Lipase enzymes breakdown and mobilize lipids which are present within the cell of the organism and the breakdown of lipid is also present in the environment. However there are many questions still unanswered. For example, is the activity of the lipases different when they are produced by different strains or species of bacteria? Hopefully in this research paper, this question will be answered.
When bacteria is grown in a surrounding of hydrophobic media, the bacterial cell releases lipase for the breakdown of fats in the environment for a source of energy. Bacteria produce lipases during the late phases of log phases and in the stationary phases. Lipases are hydrolases which hydrolyzes triacylglycerols in aqueous conditions to form fatty acids and glycerol. The reaction releases energy which is used for growth of the bacteria which is why the bacterium produces lipases within these phases. The substrates of the lipases are triacylglycerols which are hydrophobic and the reaction occurs in aqueous condition and this leads to the reaction occurring in lipid-water interface. Some lipases can also catalyze the synthesis of long chain fatty acids.
Lipases contains α/β fold, which has eight β sheets in the middle which are parallel except for the second β sheet and the sheets are surrounded by α helices. This fold offers a scaffold for the active site in the lipase molecule. The active site or binding site of the lipase molecule is where the interface occurs. This is where the chains of the enzyme are subdivided; at the bottom of the active site is where the ester bond binds to which means this region is hydrophilic. Towards the surface of the enzyme is where the molecule binds to the fatty acids and therefore this region is hydrophobic. Within the β-sheets there is an area which is highly conserved which is made up of the triad which is a nucleophile and histidine. The nucleophile is made up several amino acids, which are Serine, Cysteine or aspartic acid. The nucleophile is present on β5 and the histidine is present on β7. The histidine is the only highly conserved area of the active site/enzyme that differs in shape and structure from one type of lipase enzyme to another. Another area of the active site that is important but only present in some type of lipases is the lid. This area is what gives the lipase enzyme the structural explanation of the interface property. When the substrate comes into contract with the lid, it opens the lipid – water interface where the substrate binds to for the reaction to occur. Some lipase molecules are only active in the presence of Ca2+ and this is due to the subdivisions of the active site being bound together by the Ca2+ion. The hydrophobic region of the active site leads to less inhibitors that can bind to and inactivate the enzyme.
Since lipases are extracellular enzymes, the secretion/production of these enzymes is affected by a number of factors:
- Nutritional – enzymes are produced when the bacteria is in the presence of a lipid environment such as oil, tweens, hydrolyzable esters and triacylglycerols. These are the main sources of lipid but many bacteria can produce lipases in the presence of various sources of substrates. For example Pseudomonas aeruginosa produce lipase in the presence of long chain fatty acids such as oleic and linoleic acid.
- Temperature – the temperature at which maximum production of lipase can occur depends on the optimum temperature for growth of bacteria. The temperature normally ranges from 30 – 60°C, but some can survive at colder or warmer temperatures. Therefore it depends on the type of bacteria in question.
- pH – normally bacterial lipases are active in neutral pH or alkaline pH. However there are a few exceptions like Pseudomonas fluorescens’ lipase has an optimum pH of 4.8, whereas most bacterial species possess stability over a broad range of pH of 4 – 10.
- Effect of ion – one type of lipase which is produced by Pseudomonas species is activated by the presence of Ca2+ ion in the environment.
- Growth of bacteria – if the bacterial cell is present in the log phase then the production of lipase is decreased in the bacterial cell.
Inhibitors – inhibition of lipases does not affect the production or the secretion of the enzyme but affects the activity of the enzyme. There are two types of inhibitors; irreversible or reversible. The reversible inhibitors are split into two types. The first of which are non specific as they bind to the enzyme but not at the active site. When the inhibitor binds to the enzyme, the active site changes and therefore prevents the lipases from binding to the substrate as the structure of the active site has been changed. An example of this type of inhibitor is bile salts. However bile salts can activate some lipases such as the lipase produced by the pancreas. The second type of reversible inhibitors is specific inhibitors as they bind to the active site of the lipase enzyme. They can also be irreversible as the interaction between the inhibitor and the enzyme is so strong that it cannot be broken. An example of this type of inhibitor is boronic acid which can bind to the active site for a long time but can still be removes leaving the active site unchanged. These types of inhibitors bind to the triad of the active site, which means that when they bind to the triad, the interaction is irreversible.
There are three major types of microbial lipases depending on the substrate they bind to.
- Nonspecific – these enzymes act randomly on the lipid substrate molecules which then completely breakdown the molecule. For example with the triglyceride molecule, the enzyme will break the ester in random fashion until the molecule is complete broken down to fatty acids and glycerol.
- Regiospecific – these enzymes only hydrolyze the primary ester bond, these are the C1 and C3 bonds in the triglyceride molecule , which means that when hydrolyzing triglycerides the final products are free fatty acids, 1, 2(2,3)-diacylglyceride and 2-monoacylglyceride.
- Fatty acid-specific – there are some bacteria that only produce this type of lipase and they bind to fatty acids which are then broken down by the lipase. One type of bacteria that can produce lipases that only bind fatty acids is the Achromobacterium lipolyticum. Other bacteria that produce this type of enzyme are Bacillus species which mostly bind to long chained fatty acids. However other bacteria like Pseudomonas species produce lipases that can bind to short or medium length of fatty acids. Staphylococcus aureus can produce a lipase molecule that can bind to unsaturated fatty acids.
Lipase in Decomposition
The bacteria that are going to be used in the research project are bacteria that were purified from a model burial environment. The bacteria that were present in the model burial environment must have been already been present in the pig’s body, which means that all the bacteria that are going to be used are endogenous bacteria that are part of the pig’s microflora. The bacteria sample had been taken out of the fluid from the decaying organism in a steel box which was free from all external environmental factors except from oxygen. The sample of bacteria was taken two times a week and then towards the end it was reduced to once a week. It was discovered that at the beginning of the decaying process the bacteria that were present were Gram negative bacteria. However after week 9 the bacteria that were growing in the decaying pig changed from Gram negative to Gram positive. These bacterial cells can release lipases which can break down fats in the body which leads to the formation of adipocere. Adipocere is made up from a mixture of saturated fatty acids which have been produced during decomposition of the adipose tissue in the body. These adipoceres are formed straight away after death by lipases which are present inside the body. These lipases are mostly produced by the bacteria in the body of the pig which breaks down triglycerides to free fatty acids. If in a suitable environment, bacteria release lipases for hydrogenation of unsaturated fatty acids to its saturated form.
There are two assays that will be performed to find out the activity of the lipase which are present in the solution. The first is based on BALB – DTNB method and it uses dimercaprol tributyrate (BALB) and 5, 5′ – dithiobis (2-nitrobenzoic acid) (DNTB). The lipase enzyme binds to BALB and cleaves it to form an SH group which then binds to DNTB. The product then forms a yellow product which then increases the absorbance which can be measured using a spectrophotometer. The colour intensity is measured at 412 nm; the colour change is proportional to the activity to lipase at to 1:1 ratio.
The second assay also uses the spectrophotometer but this time it measures the optical density of the solution instead of measuring the amount of product that is formed. Tributyrin and olive oil is emulsified in the solution which gives a turbid appearance. As the lipase breaks down the lipid in the assay solution, the optical density of the solution decreases which can be measured. The optical density of the solute ion can be measured at 450nm. Both assays measure the activity of the lipase but in two different ways. The first measures the amount of product that is formed while the second measures the breakdown of the substrate.
AIMS AND OBJECTIVES
Decomposition of human or animal bodies is dependent upon a number of factors. Bacteria which are endogenous (in the body) and exogenous (in the environment) are the key components of decomposition. Different tissues in the body degrade at different rates and are degraded by different bacteria.
Previously it has been shown that bacteria in the model burial environment can produce lipases which breakdown the lipids found within the tissues of the body. However it does not tell you if there are different lipases that are secreted by different bacterial cells. Lipase production was demonstrated by using plate assay when lipase breaks down tween 20. Therefore it does not compare the different lipases produced and the activity of different bacterial species.
There have been different spectrophotometric assays that have been described in the literature to calculate the activity of lipase enzymes, but only two of these will be used. The bacteria that is going to be used in the assay has been purified from fluid from a decaying pig in a steel box which is free from all external environmental factors expect oxygen.
Two assays are going to be preformed to find the activity of lipase, the first one similar to the BALB – DTNB method. Lipase forms a SH group on BALB which then binds to DTNM to give a yellow product. The amount of product that is formed in a solution is related to the activity of lipase in a 1:1 reacting ratio which is a direct measurement of the activity. The colour change is measured at 420 nm.
The second assay is also measure the change in the solution but this time it measures the decrease of the substrate that is left in the solution. It measures the density of the solution, as the substrate (olive oil) is denser than the product. The density is measured 450 nm. The decreased of the substrate is related to the activity of lipase.
At first before anything can be done we need to see if the bacteria cells produced lipase is by growing them in a plate which contains Tween 80. If the Tween is broken down then the bacterial cell produces lipase.
MATERIALS AND METHODS
The bacterial strains that were given to me were extracted from fluid from a pig that was decaying in a steel box which had a controlled environment that was free from all external environment factors expect fresh air.
The bacterial strains were grown in half nutrient agar which was made from 2.6g of nutrient broth (OXOID, Basingstoke, England) and 4.8g of Agar bacteriological (OXIOD) in 400ml of water which was autoclaved and then poured in to 20ml Petri dish. The bacterial strains were plated and left in a 30°C incubator overnight. After the bacteria were grown on just half nutrient agar, they were then grown on half nutrient agar with 4ml of sterile Tween 80 (SIGMA ALDRICH, UK) and 400µl of 10% of CaCl2 (scientific equipment, Loughborough, England). Again the plates were placed in a 30°C incubator.
The bacterial strains were also grown in minimal medium agar which contained 2.8g of Potassium Hydrogen Orthophosphate (BDH Laboratory Supplies, Poole, England), 1.2g Sodium Dihydrogen Orthophosphate (BDH LS) and 0.04g of Magnesium Sulphate (BDH LS) in 200ml of sterile water and 2.4g of Agar bacteriological. After the solution came out of the autoclave, 2ml of Tween 80 was added and 200µl of 10 % CaCl2.
For the bacterial strains to be used in spectrophotometric assay, the strains had to be grown in liquid media. The bacterial strains were grown in two different types of media, Tryptic Soy Broth and Minimal Medium.
The Tryptic Soy Broth (TBS) was made from 30g/L Tryptone Soya Broth (OXIOD) which was autoclaved. After the bacteria were added to the media, the bottle was placed in a shaking incubator at 37°C over night.
The Minimal Medium contained 14g/L of potassium hydrogen orthophosphate, 6g/L sodium dihydrogen orthophosphate and 0.2g/L of magnesium sulphate. 100µl of Tributyrate (SIGMA ALDRICH) was added to 10ml of the Minimal Media. The bacteria were added to the media and then placed in a shaking incubator at 37°C over night.
After the bacteria are left to grow, the media is used to make up three different samples of bacteria to use in both of the assays. The first sample is purified bacterial strain from the media and this was obtained when 1ml of the media was placed in a sterile eppendorf tube which was then centrifuged at full speed for 2 minutes. The supernatant was replaced with 500µl of 150mM of CaCl2 and 500µl of 200mM of Tris buffer (12.11g of Trizma base in 150ml of water and then 0.1M of HCl was added to make the pH of the solution 8, this to make 0.5M Tris Buffer which was then diluted to make 200mM solution) (SIGAM ALDICH).
The second sample was done in the same manner but instead of adding Tris buffer and CaCl2 to the pellet, PBS (Phosphate buffered saline) solution is utilized to re-suspend the pellet and 2ml of the media solution is used. Each suspension was transferred in to a different Bijou Bottle which is kept on ice. The suspension in the Bijou Bottle is sonicated twice for 30 seconds at 30W.
The last sample was made when the media solution is filtered with the use of a sterile syringe and sterile 0.2µm pore syringe filter and placing the filtered solution into a sterile universal bottle. 3ml of the media was only filtered.
The samples were ready for the assay and two different that were used. They both measured the absorbance of the solution at different wavelengths. One measured the turbidity of the solution while the other looked at the change in the absorbance of the solution.
For the turbidity assay an emulsion solution is made and it is made from 100mM of Tris buffer (4.975ml), 50mM of CaCl2 (4.975ml) and 50ml of lipid source (either olive oil or Tributyrate or both). The solution was sonicated for 3 minutes at 40W. The solution is left in a water bath until it is used for the assay. The emulsion solution is used in three different ways as the assay was performed in a cuvette, Petri dish or 96 well plate. When done in a cuvette, 40mg of low melting point agarose (SIGMA ALDRICH) is added and the boiled before sonication. The agarose stabilises the emulsion. If the assay was done in a 96 well plate, then no agarose is necessary. The last test that is performed is in 20ml plates; 20ml of the emulsion solution is made up with 80mg of agarose to made a solid media (INVITROGEN, Paisley, UK) which is then boiled before and after sonication.
For the 96 wells plate, 200µl of the emulsion solution was placed in each well and then 20µl of the sample solution was added. As soon as the sample was added the absorbance is measured at 450nm to measure the optical density of the solution. The absorbance was then measured every 15 minutes up to 60 minutes. Here the samples that were used were grown in the Minimal Medium. The lipid source in this part of the assay was 25µl of olive oil and 25µl of Tributyrate in 10ml of the emulsion solution.
For the assay that was done in the cuvette 1L of the emulsion solution was added to a micro cuvette and 100µl of the sample solution. The absorbance was also measured at 450nm as soon as the sample is been added and then every 5 minutes up to 45 minutes. The lipid source is 50µl of olive oil in 10ml of emulsion solution.
For the plate assay after the solution was boiled for the second time, the solution was poured in to a plate for the agarose to set. After the agarose was set, wells were made in the agarose using a hollow punch about 8mm in diameter which was filled with 10µl of the sample solution and the plate was left at room temperature over night. In 20ml of the emulsion solution the lipid source was 50µl of each olive oil and Tributyrate.
Colour Assay (BALB – DNTB Method)
The second assay measures the absorbance change in the working solution. The working solution is made from BALB (SIGMA ALDRICH) and DNTB (SIGMA ALDRICH) and Tris buffer solution. The working solution was made from 1 ml of BALB is added to 17.5ml of 0.5M of Tris Buffer at pH 8.5 and 625mg of DNTB. 150µl of the working solution is added to the well after adding 150µl of water. To this 10µl of the sample was added. When the assay was done in 96 well plate the absorbance was measured after the sample was added at 405nm and then every 10 minutes for 30 minutes.
When the assay was done in a cuvette, at first 400µl of water was placed in the cuvette then 380µl of the working solution was added to the water. Then the 20µl of the working sample was added into the cuvette. The absorbance was the measured at 412 nm for the 20 minutes. The reason why there is a difference in the wavelength in which the absorbance is measured is due to the plate reader not being able to read the absorbance at 412nm. For this assay the samples that were used were prepared from the bacteria that were grown in TSB.
When the bacteria colonies were grown on the agar plate which had Tween 80 and CaCl2, around the colonies there was the presence of halos or the colonies has a halo this can be seen in figure 1a. The arrow shows the halo colonies of the bacteria species. The bacteria colonies that were placed on other plates was not as clear as 16C but the halo can only be seen when the plates are held up by the light (result not shown).
The first assay that was done was the turbidity assay in a cuvette, the optical density of the solution did not increase or decrease, and it just stayed the same. But when the assay was done in the 96 well plate the optical density increased when the bacteria were added to the well, and then decrease and keep decreasing even after 60 minutes (figure 2a).
Then the filtered media was added to the emulsion solution in the 96 well plate, the optical density again decreased. However not all the bacteria were filtered to see if there was a decrease in the optical density (figure 3). Only some of the bacteria were used to see if it was an enzyme that was decreasing the optical density and not the bacterial cells. However the general result showed a decrease in the optical density except for 2 bacterial strains (1A and 4A) which showed an increase in the optical density after 30 minutes and then it optical density again.
Then the bacteria cell free lysates were added to the welled plate and the same result appeared as the optical density levels decreased once again. The bacteria that were used were the same bacteria that were used in the filtered part of the assay (figure 4). After 45 minutes the optical density is starting to level off. The gradient of the line for all the bacteria strains are the same as they all decrease at the same rate expect for bacteria strain 5 which has flatter gradient than the rest.
For the plate test in the turbidity assay, the bacterial solution in the well was not present and no zone of clearance was noticeable in any of the plates (figure 1b). Only one of the plates is shown in the figure and the rest of the plates looked the same as no zone could be seen.
Colour Assay (BALB – DNTB Method)
In the BALB-DNTB method, the absorbance increases when bacteria strain 6 was added to the working solution in a cuvette and measured for 20 minutes. The increase was slow for the first 10 minutes and then increased at a faster rate for the next 10 minutes, figure 5.
When the assay was done in the welled plate, the absorbance increases for all the strains but some increase more than others. For example strain 5 increased from 4.204 to 4.412 while strain 1 only increased from 4.241 to 4.265. This is shown in a table in figure 2b.
When only the media in which the bacteria grew in was added as the sample, the absorbance also increased for most of the bacterial strains but not as much as when the bacterial cells were added. For some of the strains the absorbance decreased. For example in strain 1 there was a decrease from 4.241 to 4.235, figure 2c. The same happened when the content of the bacterial cell was added to the working solution. But when the absorbance increased, the increase was bigger than the increase when media was added (figure 2d). However there were still some strains in which the absorbance still decreased in 20 minutes but the absorbance increased from 0 to 10 minutes and then decreased from 10 to 20 minutes.
Figure 1, (a) the plate has been plated with strain 16C (left) and 16B (right); the halo can be seen clear by the arrow which is the colonies of bacteria 16C. However the halo can not be seen clearly in the colonies of bacteria. (b), the plate contain solid emulsion solution with well which contain lipases from different bacteria, and there is no presence of zone of clearance from any of the well. There were 3 plates in total and all look the same (only one is shown) but the well had different lipases from different bacteria.
Figure 2, A is a table that shows the optical density change when bacterial was added to emulsion solution for the turbidity assay. The optical density decreases when the bacterial cells were added to the emulsion solution. The next 3 tables are showing the absorbance change when the strains were added to the working solution for the colour BALB-DNTB method, (B) has bacterial cells added to the working solution; (C) has only filtered media, which had bacteria growing in, was added and lastly (D) had bacterial cells free lysates added. In the colour assay the absorbance increased in all three cases.
Bacteria produce lipases that can break down or hydrolyse lipid molecules such as fats and oils. They produce lipases in the log phase of growth when there is a high level of lipid source for energy. There are different lipases which can break down different lipid molecules. The bacterium produces lipases to break down lipid for energy as adequate amount energy is present in lipids. As most of the lipids cannot cross the cell membrane, the lipid has to be catabolised into smaller lipid molecules which can then enter the cell where it is broken down further. Lipases from bacteria are studied for industrial uses. Here it was studied to see if the lipases that were produced from different bacteria are different and if there was any variation in the activity of the lipases.
When the bacterial cells were grown on agar plate without any Tween 80 the bacterial colonies do not have any halos or precipitate around the colonies. But when some of the bacteria were grown in agar that contained Tween 80 and CaCl2 the colonies had halo colonies 3 to 8 days after they were inoculated. In the past Tween has been used for lipase activity to see if the bacteria produce lipase. If lipases are produced then it binds to the Tween and breaks the Tween down to fatty acids. The fatty acids then bind to the Ca in the media which forms crystals. These crystals then become soluble in the media which can then be seen by eye as halos. Some of the colonies had halos which meant that the cell produced lipases.
Figure 6, the turbidity plate assay should have looked like this but what the figure 1b shows. There the one of clearance can be seen very clearly where as in the plate in figure 1b there are no clearing at all what meant the assay did not work at all.
The turbidity assay that was done is the plate which showed no zone of clearance, it should have had zone of clearance around the well which contained the sample of bacteria. The bacteria in the wells should have diffused out of the well and in to the agarose media in which the bacteria should have released lipases to break down the olive oil and Tributyrate. When the lipids were broken down the media would have become clear. The plate should have look like figure 6 from, the zone of clearance is shown very clearly.
The other assay that did not work was the same assay that was done with the cuvette. This is when the absorbance levels did not decrease but just stayed the same. The absorbance levels should have decreased and the reason in why this did not occur is not known. It might have been due to the stability of the solution as the agarose must have been concentrated which meant that the bacteria solution was not able to diffuse through the media.
The concentration of agarose might be the problem because when agarose was not added like in the 96 well plate part of the assay, the absorbance of the emulsion solution decreased. This was due to the emulsion solution being turbid by lipid in the solution when sonicated, when the bacteria sample was added the optical density increased slightly as the bacteria cell scatter the light which leads to the increase in the optical density absorbance levels. The bacteria cell then releases lipase in the solution or lipase that are inside the cell break down the lipid in the emulsion solution which then leads to the decrease in the level of lipid in the emulsion solution which then means that less light is scattered.
The well plate assay was done to 3 different type of sample solution, one of which contained bacteria cell, one of which contained the filtered media solution and the last contained the bacteria cell free lysates. The bacterial cells were used to see if the bacterial cell produced lipases. The filtered media was used to see if the bacterial cell released lipase in to the media and if it was in fact the lipase that was decreasing the absorbance and not anything else. The bacteria content was used after the bacteria cell were sonicated for one minute, to use all the lipases that had been produced by the bacterial cell but not secreted. As not all the bacteria cells release the lipase in to the media and sometime the lipid molecule is too big to cross the cell membrane and wall of the bacteria.
To see if there are any differences in the activity of the different lipases which are produced by different bacterial cells, cannot be done by adding the sample to the emulsion solution as different concentration of lipase must have been in the sample for each of the strains. In order to make the test fair, the amount of bacterial cell and the lipase concentration must be the same for each of the bacterial strain. But still it might be a fair test as some of the bacterial cells can still divide inside the emulsion solution and then increase the concentration of lipases. The lipases produced by the bacteria are produced in the log phase.
The same can be said for the BALB-DNTB method. This assay is not like the other assay because the absorbance does not decrease but increase. This is due to the lipase bind to the BALB in which is cleaved to form a SH group. The SH group then binds to DNTB which is in excess in the working solution, to form a yellow substance. The complex then absorbs light hence increasing the level of absorbance. The bind of the BALB with the new SH group binds to the DNTB in a one to one reacting ratio, this means that increases is absorbance is proportional to the reacting activity of the lipase.
When bacterial cells were mixed to the working solution the absorbance for most of them increase. This meant that lipases that were present in the well were cleaved BALB. The same thing also occurred when filtered media was added to the working solution but the increase were small and this must be due to the fact that not a lot of lipases were released by the bacterial cells in to the media solution. However, when the bacterial cell free lysates is added not all of the absorbance levels increase but in fact some of them decrease and then increase. It may mean that the lipases need time to start working since they had been on ice before the experiment. To see if this was true, the test needs to be done again but for a longer period of time.
In the cuvette test, only one strain, it was used when the first assay was done it had the largest change in absorbance. It was used to see a general increase of the solution over 20 minutes and the absorbance was measured every minute to see the turning point when the rate of enzymatic activity change from being slow to a steady normal rate. The graph in figure 5 shows that the rate was slow during the first 10 minute this meant the bacteria cell needed to adapt to the new environment before the activity of the enzyme can to back to normal. If the test was done longer then the graph would start to level due to the substrate concentration starting to decrease.
From the results, there is not enough evidence to conclude that there any differences in the activity of the different strains of lipase. To see if it is true then the both of the a
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