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What is the expected effect of pH on the activity of a fungal pectinase?

What is the expected effect of pH on the activity of a fungal pectinase?


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I am working on an enzyme assay for a fungal pectinase.I assayed the enzyme in different buffers from pH 1-12.5 However,the enzyme has good activities starting from pH1-10.5.

Is it possible to have enzyme activity over such a broad range of pH?


I actually doubt that the pectinase has such a broad pH range in which it works optimally. Searching the web I found two figures which support my doubts:

The first is from an article ("Immobilization of pectinase by adsorption on an alginate-coated chitin support") which compares the activity of native and immobilized pectinase under different conditions. The second is from the datasheet of a company which sells the enzyme. Both show an activity over 50% roughly between pH 3.5 and 5.5 but not at higher or lower pH values.


What is the expected effect of pH on the activity of a fungal pectinase? - Biology

Catalase

Hydrogen peroxide (H2O2) is a common by-product of metabolic reactions. In high concentration it is toxic therefore, its accumulation in cells would be harmful. Most tissues, however, contain the enzyme catalase, which catalyzes the breakdown of peroxide to water and oxygen as follows: The reaction is extremely rapid. The action of the enzyme can be demonstrated easily by the evolution of oxygen in the form of gas bubbles when an extract of a tissue containing the enzyme is added to a dilute solution of hydrogen peroxide. We will use homogenized (ground-up) chicken or beef liver as a source of catalase. Catalase

  • Catalase - general information Classrooms of 21st century
  • Catalase - An Extraordinary Enzyme
  • Heat denature enzyme
  • Structure of Catalase image and graph
  • Links -

Amylase

    Amylase References:
    • Amylase - Independent study by Amy Caruana (Student at Kean Univ)
    • Amylase - results from U. North Dakota
    • Enzymes amylase, temp, pH, substrate concentration
    • Benedicts test for reducing sugar - image of result
    • Amylase How does it work?
    • Amylase - starch hydrolysis
    • Salivary amylase - pH
    Pepsin is a protease that begins digestion of proteins, breaking them into peptides and amino acids. Pepsinogen, is secreted by gastric glands of the stomach into the stomach. There, in the acid environment of the stomach, pepsinogen is converted into pepsin.

    Although both pepsin and trypsin are proteases, they require quite different conditions of acidity and alkalinity for their action.

      Pepsin References:
      • Pepsin -
      • Pepsin - molecule of the month
      • pH optimum -
      • Pepsin - mentions pH optimum
      Trypsin is a protease secreted into the small intestine by the pancreas. As pepsin, trypsin digests proteins into peptides and amino acids and is made and secreted in an inactive form, trypsinogen.

      Although both pepsin and trypsin are proteases, they require quite different conditions of acidity and alkalinity for their action.

        Trypsin References:
        • Trypsin Encyclopedia description
        • Illustration search for trypsin within this large file
        • Images Google search engine

        An Analogy

        Suppose you are interested in purchasing a Pizza store and wish to investigate how productive the store is without the present owner knowing because, you fear the owner will raise the price. So, instead of going into the store and watching what happens and asking to examine the books that record expenses and profits, you decide to watch the store from outside.

        You observe how often trucks arrive with pizza dough, pizza toppings (cheese, pepporoni, etc.), and other supplies. You also observe how often workers leave the store to deliver pizzas to customers.

        In this analogy, the pizza supplies are the reactants and the boxed pizzas that are delivered to customers are the end products. The workers within the store that shape the dough, add the toppings and place the pizzas in ovens and finally in boxes are the equivalent of the enzymes.

        Although we don't actually see the workers doing their job, we can infer that if the store is using large quantities of reactants (dough and toppings) and / or making large numbers of end products (pizzas) that the workers (enzymes) must be very active.


        You might also Like

        Beta amylase isn't found in human tissue. irontoenail January 8, 2013

        I was actually reading the other day about how there are some natural chemicals that can trick you into thinking something is sweet when it isn't. It happens when you eat an artichoke, for example. Afterwards lots of things taste sweet (like pure water, for example).

        They are trying to figure out ways to use it as a low calorie sweetener. I wonder if it works with enzymes the way amylase does. Although, with amylase I guess it is actually converting starch to sugar, where the chemicals just make things taste like they have sugar in them. croydon January 8, 2013

        @Iluviaporos - Actually, I don't think it's because of digestion that people are told to chew more. I mean, it might be true for some people, who have problems with their digestion in general, and it might be slightly true for foods that don't digest easily, like meat. I don't know about that.

        But I was always told that people should chew their food more because it slows down the whole process of eating and gives the body time to tell you that it's satisfied. I always thought people just stop eating when their stomach is full, but when I was diagnosed with insulin resistance my doctor explained that the body actually decides that it is full when it has enough blood sugar.

        So, I guess chewing a lot could also help with that, as it would speed up digestion and allow blood sugar to hit faster. lluviaporos January 7, 2013

        Amylase is the reason that they say you should try to chew your food thoroughly before you swallow it. Saliva isn't just there to keep your mouth from drying out, it also starts the digestive process before the food even hits your stomach.

        They used to recommend that people chew each mouthful about 40 times before swallowing it. I don't know if it's the real amount you should do it for, but most people definitely don't do it enough.


        Results

        Screening of isolates for pectinase activity

        Pectinase has many applications in various industrial processes ([16] #1170 [17] #1311). To screen excellent strains for pectinase production, some isolates from the primary screening were spot inoculated on pectin agar plates (PAPs) in triplicate. An additional image was shown this in more detail [see Additional file 1: Figure S1]/(see Additional file 1). The Hc value of 33 for this isolate was significantly higher than the values for other isolates (8.87 ± 0.15, Fig. 1). This bacterium was labeled as isolate B. tequilensis CAS-MEI-2-33, and further characterization and identification were carried out.

        The Hc values of screened strains. Values are given as the means ± standard deviation (n = 3). Different letters indicate significant differences at 5%

        Morphological characteristics of B. tequilensis CAS-MEI-2-33

        B. tequilensis CAS-MEI-2-33 was a gram-positive bacterium that contained spores, and the shape of the bacterial cells was clubbed. When grown at 37 °C for 8–12 h in LB agar, colonies of B. tequilensis CAS-MEI-2-33 were round, smooth, and cream colored, and the margin was entire.

        Biochemical characterization of B. tequilensis CAS-MEI-2-33

        The biochemical characterization of B. tequilensis CAS-MEI-2-33 was performed using a variety of tests, including the Voges–Proskauer, nitrate reduction, glucose utilization, catalase, motility, lysozyme tolerance, phenylalanine, gelatin, starch, lactose, casein, and mannitol tests. The results of the phenylalanine, starch, and mannitol tests were negative, while the results of the remaining tests were positive (Table 1).

        Identification based on phylogenetic analysis

        The phylogenetic tree generated using the 16S rRNA gene sequence of the bacterial isolate showed the highest homology (100%) with B. tequilensis. Furthermore, the biochemical characterization of the ability of CAS-MEI-2-33 to metabolize starch and mannitol was different from that of Bacillus subtilis but similar to that of B. tequilensis. According to Bergey’s Manual of Determinative Bacteriology, this strain was named B. tequilensis CAS-MEI-2-33 ([18] #1309). The constructed phylogenetic tree indicated that this strain had the closest genetic relationship with the B. tequilensis strain P12Pb (Fig. 2). The tree was inferred by the neighbor-joining method using MEGA 7.0 software. The numbers at the nodes of the tree are indications of the levels of bootstrap support based on neighbor-joining analysis of 1000 inferred replications.

        Phylogenetic tree based on 16S ribosomal RNA sequence analysis showing the position of the strain CAS-MEI-2-33 using MEGA 7.0 software. Numbers at branching points refer to bootstrap values (1000 resamplings) with 0.50 as the sequence divergence

        Toxicity of nicotine to B. tequilensis CAS-MEI-2-33

        TS, contains nicotine, which is harmful to many bacteria, and serves as a distinct source of nutrition. The average nicotine content in TS is as high as 1900–3800 mg/kg ([1] #1136), which is equal to 76–152 mg/L in the fermentation medium. In this study, 500–2000 mg/L nicotine in the medium was tested. The highest concentration of nicotine in the test medium was approximately 13 times that in the TS fermentation medium. However, the growth of B. tequilensis CAS-MEI-2-33 was less affected under the experimental conditions (Fig. 3).

        The growth of B. tequilensis CAS-MEI-2-33 under different concentrations of exogenous nicotine. Values are given as the means ± standard deviation (n = 3)

        Production of pectinase

        The production of high titers of pectinase by optimizing the growth parameters is of prime importance in industry ([19] #1145). In the present study, the one-factor-at-a-time method was implemented to optimize the components of the medium and conditions. The enzymatic activity was detected using 3,5-dinitrosalicylic acid (DNS) reagent (Beijing Leagene Biotech. Co., Ltd.) based on the production of D-galacturonic acid ([20] #1161). The curve for the production of pectinase during the fermentation of B. tequilensis CAS-MEI-2-33 was pivotal for indicating the optimal fermentation. The activity of pectinase increased gradually until 40 h, except for 32 h, and then began to decline (Fig. 4a). The highest pectinase activity for the process of fermentation using TS as the feedstock was 293 U/mL at 40 h. Parameters such as the pH of the fermentation medium have a substantial influence on the growth of strains and pectinase production. Pectinase activity (618 ± 9 U/mL, Fig. 4b) was significantly increased at pH 7.0 compared to that under other conditions.

        Pectinase activity of B. tequilensis CAS-MEI-2-33 during fermentation. a Pectinase activity during B. tequilensis CAS-MEI-2-33 growth. b Effect of the initial pH of fermentation medium on enzyme activity. c Effect of tobacco stalk concentration in the fermentation medium on enzyme activity. d Effect of the amount of inoculum on enzyme activity. Values are given as the means ± standard deviation (n = 3). Different letters indicate significant differences at 5%

        After the determination of pectinase activity with different concentrations of TS in the fermentation medium, it was found that at the concentration of 40 g/L TS and the optimized pH, pectinase activity was the highest (730 ± 38 U/mL, Fig. 4c), as supported by ANOVA (p = 0.05).

        The inoculum amount played a key role in the initial fermentation. Under the optimal pH and TS concentration, inoculum concentrations of 1, 3, 5, 7, and 9% were tested. The highest pectinase activity (1370 ± 126 U/mL, Fig. 4d) with 3% inoculum was not significantly different from that with 5% inoculum. At higher levels, such as 7 and 9%, enzyme production declined, which could be due to competition for nutrients among the population of bacteria, as has been observed in Thermomucor indicae-seudaticae ([21] #1230).

        Pectinase properties

        The results of the pectinase property analysis are shown in Fig. 5. The pH of the reaction system can affect pectinase activity. Pectinase activity increased when the pH of the reaction systems was increased from 6.0 to 10.0, but this activity was nearly undetectable at pH 11.0. Our results showed that 791 ± 42 U/mL pectinase activity at pH 10.0 was the highest recorded value (Fig. 5a). Subsequently, the influence of temperature on pectinase activity was investigated. Pectinase activity increased when the reaction temperature was increased from 30 °C to 40 °C, reached maximum activity at 40 °C, and then decreased rapidly as the temperature increased beyond 40 °C. The pectinase activity at 40 °C was higher than that at other temperatures (Fig. 5b). Thus, pectinase was more active at an alkaline pH and intermediate temperature. The effects of different metal ions on pectinase activity are shown in Fig. 5c. Ag + , Li + , Cu 2+ , Ca 2+ , Ba 2+ , and Mn 2+ ions increased enzyme activity in particular, Ag + ions increased pectinase activity by 193.95%, which was approximately 1.94 times higher than that of the control. K + , Co 2+ , Ni 2+ , Mg 2+ , Zn 2+ , Cd 2+ , and Fe 3+ ions, especially Zn 2+ , inhibited pectinase activity. Figure 5d shows the thermal stability of pectinase produced by B. tequilensis CAS-MEI-2-33. However, pectinase activity decreased when the cell-free supernatant was placed at 60 °C. Pectinase activity was stable when the cell-free supernatant was incubated at 40 °C however, pectinase could not tolerate the high temperature for a long time.

        Enzymatic properties of pectinase. a Effect of substrate pH on pectinase activity. b Effect of reaction temperature on pectinase activity. c Effect of metal ions on pectinase activity. d Temperature stability of pectinase. Values are given as the means ± standard deviation (n = 3). Different letters indicate significant differences at 5%

        Partial purification of pectinase from CAS-MEI-2-33

        Under the optimal fermentation conditions, 2.30 L supernatant was obtained by centrifuging the bacteria in a 3.0 L fermentation flask. The pectinase activity reached 1771 U/mL, and the total pectinase activity was 4,074,513 U (Table 2).

        According to the ammonium sulfate fractionation curve, suitable saturation was selected for salting out. The fractionation curve is shown in Fig. 6a. When ammonium sulfate was saturated to 70–80%, the pectinase activity of the precipitate increased, while that of the supernatant decreased significantly. When the saturation rate reached 80–90%, the pectinase activities (pH 10.0, Gly-NaOH) of the precipitate and supernatant did not change significantly. The target protein was dissolved with 30 mL buffer (pH 8.0, Tris-HCl) after salting out. Then, the target protein was further renatured with 55.0 mL buffer (pH 8.0, Tris-HCl) through a dialysis bag. The pectinase activity was 5166 U/mL, and the total activity was 284,144 U.

        The purification of alkaline pectinase from B. tequilensis CAS-MEI-2-33. a Ammonium sulfate fractionation curve b Elution curve of Mini Macro-Prep High-Q ion exchange chromatography c. Elution curve of Sephacryl S-100 column chromatography d. The partial purification of alkaline pectinase from B. tequilensis CAS-MEI-2-33 using TS. M: molecular weight makers 1. Sephacryl S-100 column chromatography with pH 7.2 2: Mini Macro-Prep High-Q ion exchange chromatography with pH 8.0 3. Concentrated ammonium sulfate salting out solution

        After dialysis, the supernatant was purified with a Mini Macro-Prep High-Q ion exchange column. The results are shown in Fig. 6b. The pectinase was eluted with pH 8.0 Tris-HCl buffer, and when the column was gradiently eluted with pH 8.0 Tris-HCl buffer containing 0–1 mol/L NaCl, peaks eluted. By detecting pectinase activity, it was found that the first elution peak was active. The first elution peak was collected with pectinase activity of 862,352 U/mL. The total activity was 77,611 U. Sephacryl S-100 was equilibrated with ultrapure water, sampled, and then eluted with pH 7.2 PBS buffer, with a flow rate of 0.8 mL/min, and the effluent was detected online by a UV detector at a wavelength of 280 nm to record the ultraviolet absorption peak curve (Fig. 6c). The components were collected and used for the determination of enzyme activity and protein content. The pectinase activity was 13,786 U/mL, and the total activity was 41,357 U.

        The molecular weight of the purified alkaline pectinase was detected with SDS-polyacrylamide gel electrophoresis (PAGE) as described by Mehrnoush et al. ([22] #1306). The molecular weight of the pectinase, which was approximately 45.4 kDa, is shown in Fig. 6d. Then, the protein band was cut from the SDS-PAGE gel, subjected to LC-MS/MS analysis by Shanghai Applied Protein Technology ([23] #1320 [24] #1321), and identified by searching the UniProt database. The additional figure about protein base peak was shown this in more detail [see Additional file 2: Figure S2]/(see Additional file 2 for protein peak). Finally, the protein was identified as pectate lyase, the sequence was K.ASSSNVYTVSNR.N (Fig. 7). The molecular weight was 45.4 kDa, which was consistent with the SDS-PAGE results. The results indicated that the enzyme was a good candidate for pectate lyase. Furthermore, this study was a new attempt to recycle and reuse TS in agricultural production.

        LC-MS/MS analysis of protein bands by SDS-PAGE


        Alkaliphiles

        At the other end of the spectrum are alkaliphiles, microorganisms that have pH optima between 8.0 and 11. Vibrio cholerae , the pathogenic agent of cholera , grows best at the slightly basic pH of 8.0 it can survive pH values of 11.0 but is inactivated by the acid of the stomach. When it comes to survival at high pH, the bright pink halophilic archaeon Natronobacterium , found in the soda lakes of the African Rift Valley, may hold the record at a pH of 10.5 (Fig. 9.37). Extreme alkaliphiles have adapted to their harsh environment through various evolutionary modifications. Alkaliphilic archaea have diether lipid membranes. The ether linkage is more resistant to chemical or thermal degradation compared to the ester-linked phospholipids. Given the paucity of protons in alkaline environments, maintaining a proton motive force is probably the most pressing challenge for alkaliphiles. One of the adaptations of alkaliphilic halophilic bacteria and archaea in soda lakes and other highly salty environments is the evolution of coupled transporters and flagella that exploit sodium motive force, thus conserving the PMF for oxidative and photophosphorylation by the ATP synthase. The cell surface of alkaliphiles has a high concentration of acidic (i.e. negatively charged) molecules and it has been suggested this acts as a “proton sponge”, allowing a more rapid lateral diffusion of protons from the ETS, to the ATP synthase, compared to the rate of diffusion into the surrounding waters [1] Finally, alkaliphiles may use Na + /H + antiport to create a sodium motive force. For example, the alkaliphile Bacillus firmus derives the energy for transport reactions and motility from SMF rather than a proton motive force. As with the acidophiles, the genes for secreted proteins of alkaliphiles have evolved to give enzymes that resist deprotonation/denaturation and chemical degradation at the high pH of their environment. These enzymes are also of interest to biotechnology companies. In fact, laundry detergents, which are alkaline in nature, contain alkaliphilic lipases and proteases to improve their stain-removing abilities.

        Figure 9.37. View from space of Lake Natron in Tanzania. The pink colour is due to the pigmentation of the extreme alkaliphilic and halophilic microbes that colonize the lake. [Credit: NASA]


        What is the expected effect of pH on the activity of a fungal pectinase? - Biology

        The Rate of Reaction on Liver, Apples, and Potatoes with Hydrogen Peroxide

        The purpose of this lab was to test enzyme function in various environments. Substrate concentration, temperature, and pH all affect the chemical reaction. In this lab, the enzyme catalase was used to break down hydrogen peroxide into less toxic water and oxygen gas. Using quantitative as well as qualitative observation the concept that enzymes remain after a reaction was confirmed from the first test. After testing liver, apple, and potato it was concluded that liver contained the most catalase. The final test focused on the reaction rate of liver in varying pH solutions. The relationship between catalase reaction rate and pH was shown to be parabolic with a peak near neutral.

        This lab covered the measuring of the effects of changes in temperature, pH, and enzyme concentration on reaction rates of an enzyme catalyzed reaction in a controlled experiment. The questions of how environmental factors affect the rate of enzyme-catalyzed reactions were answered in this lab. The reaction rates of enzymes were very much affected by changes in temperature, pH and enzyme concentration. The enzyme studied in this lab was catalase. Catalase breaks down hydrogen peroxide, which is toxic, into 2 safe substances- water and oxygen, by speeding up a reaction. Enzymes such as catalase are vital to our body, because if toxic substances such hydrogen peroxide were not broken down into harmless substances in our body, then they would poison our cells. The reaction that takes places is in this form: 2H2O2 ----> 2H2O + O2


        Methods
        This study was conducted at New Tech High @ Coppell under the facilitation of Mrs. Wootton on October, 8 2015. Throughout the three parts of the lab, the ability for enzymes to break down was studied. In Part A, we used 3 ml of 3% Hydrogen Peroxide on a piece of liver to observe the rate of reaction. Once this was done, we took the leftover liquids and used it on a new piece of meat to see if the enzyme was reusable. In Part B, we tested, observed, and recorded the rate of reaction with Hydrogen Peroxide on three substances: apple, liver, and potato. In Part C, we altered the pH of the solution by adding drops of diluted Hydrochloric Acid to see if it had an affect on the rate of reaction in catalyses on pieces of liver.

        When the substrate of hydrogen peroxide was added to the substances each performed a vastly different reaction. After dropping the Hydrogen Peroxide on the apple samples there was no reaction indicated therefore receiving a rating of 0. Once the same amount of the substrate was added to the potato there was a slight fizzy and bubbly reaction thus earning the rating of 2. The liver catalase was then tested and resulted in the largest reaction that was given a rating of 5 due to the speed and magnitude. After testing each substance the group then began to look into optimal conditions of pH for the reactions. After adding Hydrochloric Acid onto the liver the reaction rate seemed to diminish as the acidity increased.

        Substrate: Hydrogen Peroxide

        Looking at our data for the first experiment, we can infer that the liver contains the catalase enzyme while the hydrogen peroxide is the substrate. This is true because in the reaction, the hydrogen peroxide is observed to have gone through a chemical change turning into water molecules and dioxide molecules, whereas the liver stayed the same. Because the substrate already went through the chemical reaction, it could not go through another one with new catalase, but the old catalase still had functional enzymes and could therefore be reused. The reaction was very apparent and quick, which makes sense considering the liver’s function in an organism is to break down and purify cells.

        In the second part of the experiment, the intensity of the reaction in ascending order was apple, potato, and liver. The apple sample is simply carbohydrates meant as food for the apple seed and will have no enzymes in it, thus no reaction was observed. The potato will have a semi-intense reaction because only a small concentration of the sample of potato is enzymes whereas the rest is starch and organic compounds. The liver will have a large reaction because the majority of liver is proteins and enzymes which break down the Hydrogen Peroxide.

        In the third experiment, it was inferred that as the acidity increased in the substrate, the rate of reaction decreased. Our data shows a linear trend, however, research shows that the trend is in fact a parabola opening downwards with a maximum at a pH of 5, the acidity of Hydrogen Peroxide alone. Because we were only able to stimulate an environment of a lower pH, we were only able to get 1 direction of slope, showing linear results. This shows that enzymes only work in a certain pH level. If it’s too high or too low, it will reduce the efficiency and slow down the rate of the reactions.

        Through our experimentation the reaction between the enzyme catalase and hydrogen peroxide was tested. Repeating the reaction with various materials allowed for the concentration of catalase to be inferred. Using liver as our control, we changed the pH of the solution in each trial to test the reaction rate as the acidity increased. We concluded that as the pH of a solution increased the reaction rate increased inversely until a pH of about 5. The error within our experiment only altered the specific pH but the overall result of the last test remains true.


        Costimulation of soil glycosidase activity and soil respiration by nitrogen addition

        Synthesis Research Center of Chinese Ecosystem Research Network, Key Laboratory of Ecosystem Network Observation and Modeling, Institute of Geographic Sciences and Natural Resources Research, Chinese Academy of Sciences, Beijing, 100101 China

        Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, 73019 USA

        Department of Agriculture and Environmental Sciences, Tennessee State University, Nashville, TN, 37209 USA

        Tiantong National Field Observation Station for Forest Ecosystem, School of Ecological and Environmental Sciences, East China Normal University, Shanghai, 200062 China

        Center for Ecological and Environmental Sciences, Northwestern Polytechnical University, Xi'an, 710072 China

        State Key Laboratory of Loess and Quaternary Geology (SKLLQG), Key Laboratory of Aerosol Chemistry and Physics, Institute of Earth Environment, Chinese Academy of Sciences, Xi'an, 710061 China

        University of Chinese Academy of Sciences, Beijing, 100049 China

        Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, 73019 USA

        Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, 73019 USA

        Center for Earth System Science, Tsinghua University, Beijing, 100084 China

        Department of Agriculture and Environmental Sciences, Tennessee State University, Nashville, TN, 37209 USA

        Correspondence: Junji Cao, tel. +86 29 62336233, fax +86 29 62336234, e-mail: [email protected]

        Jianwei Li, tel. +1 615 963 1523, fax +1 615 963 1523, e-mail: [email protected]

        Rui-Wu Wang, tel. +86 29 88460816, fax +86 29 88460816, e-mail: [email protected]

        Tiantong National Field Observation Station for Forest Ecosystem, School of Ecological and Environmental Sciences, East China Normal University, Shanghai, 200062 China

        Center for Global Change and Ecological Forecasting, East China Normal University, Shanghai, 200062 China

        State Key Laboratory of Loess and Quaternary Geology (SKLLQG), Key Laboratory of Aerosol Chemistry and Physics, Institute of Earth Environment, Chinese Academy of Sciences, Xi'an, 710061 China

        Institute of Global Environmental Change, Xi'an Jiaotong University, Xi'an, 710049 China

        Correspondence: Junji Cao, tel. +86 29 62336233, fax +86 29 62336234, e-mail: [email protected]

        Jianwei Li, tel. +1 615 963 1523, fax +1 615 963 1523, e-mail: [email protected]

        Rui-Wu Wang, tel. +86 29 88460816, fax +86 29 88460816, e-mail: [email protected]

        Center for Ecological and Environmental Sciences, Northwestern Polytechnical University, Xi'an, 710072 China

        Correspondence: Junji Cao, tel. +86 29 62336233, fax +86 29 62336234, e-mail: [email protected]

        Jianwei Li, tel. +1 615 963 1523, fax +1 615 963 1523, e-mail: [email protected]

        Rui-Wu Wang, tel. +86 29 88460816, fax +86 29 88460816, e-mail: [email protected]

        State Key Laboratory of Loess and Quaternary Geology (SKLLQG), Key Laboratory of Aerosol Chemistry and Physics, Institute of Earth Environment, Chinese Academy of Sciences, Xi'an, 710061 China

        Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, 73019 USA

        State Key Laboratory of Loess and Quaternary Geology (SKLLQG), Key Laboratory of Aerosol Chemistry and Physics, Institute of Earth Environment, Chinese Academy of Sciences, Xi'an, 710061 China

        Department of Biological Sciences, University of Arkansas, Fayetteville, AR, 72701 USA

        State Key Laboratory of Urban and Regional Ecology, Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences, Beijing, 100085 China

        Synthesis Research Center of Chinese Ecosystem Research Network, Key Laboratory of Ecosystem Network Observation and Modeling, Institute of Geographic Sciences and Natural Resources Research, Chinese Academy of Sciences, Beijing, 100101 China

        Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, 73019 USA

        Department of Agriculture and Environmental Sciences, Tennessee State University, Nashville, TN, 37209 USA

        Tiantong National Field Observation Station for Forest Ecosystem, School of Ecological and Environmental Sciences, East China Normal University, Shanghai, 200062 China

        Abstract

        Unprecedented levels of nitrogen (N) have been deposited in ecosystems over the past century, which is expected to have cascading effects on microbially mediated soil respiration (SR). Extracellular enzymes play critical roles on the degradation of soil organic matter, and measurements of their activities are potentially useful indicators of SR. The links between soil extracellular enzymatic activities (EEAs) and SR under N addition, however, have not been established. We therefore conducted a meta-analysis from 62 publications to synthesize the responses of soil EEAs and SR to elevated N. Nitrogen addition significantly increased glycosidase activity (GA) by 13.0%, α-1,4-glucosidase (AG) by 19.6%, β-1,4-glucosidase (BG) by 11.1%, β-1,4-xylosidase (BX) by 21.9% and β-D-cellobiosidase (CBH) by 12.6%. Increases in GA were more evident for long duration, high rate, organic and mixed N addition (combination of organic and inorganic N addition), as well as for studies from farmland. The response ratios (RRs) of GA were positively correlated with the SR-RRs, even when evaluated individually for AG, BG, BX and CBH. This positive correlation between GA-RR and SR-RR was maintained for most types of vegetation and soil as well as for different methods of N addition. Our results provide the first evidence that GA is linked to SR under N addition over a range of ecosystems and highlight the need for further studies on the response of other soil EEAs to various global change factors and their implications for ecosystem functions.

        Appendix S1 Supplementary notes.

        Table S1 Results for publication bias.

        Table S2 Description of the 12 kinds of enzymes included in our preliminary analysis.

        Table S3 Distribution of the methods of nitrogen addition for the various types of vegetation and soil.

        Figure S1 Global distribution of the nitrogen-addition experiments selected in this meta-analysis. The map was created with ArcGIS.

        Figure S2 Frequency distributions of the response ratios (RR) of (a) α-1,4-glucosidase (AG), (b) β-1,4-glucosidase (BG), (c) β-D-cellobiosidase (CBH) and (d) β-1,4-xylosidase (BX).

        Figure S3 Relationships between the response ratio (RR) of soil respiration (SR) and the RRs of (a) α-1,4-glucosidase (AG), (b) β-1,4-glucosidase (BG), (c) β-D-cellobiosidase (CBH), (d) β-1,4-xylosidase (BX), (e) phenol oxidase (PO), (f) polyphenol oxidase (PHO), (g) invertase, (h) urease, (i) peroxidase (PER), (j) β-1,4-N-acetylglucosaminidase (NAG), (k) acid (alkaline) phosphatase (AP) and (l) leucine amino peptidase (LAP).

        Figure S4 Relationships between the response ratio (RR) of soil glycosidase activity and (a) N-addition rate, (b) N-addition duration, (c) N-addition frequency and (d) sample size.

        Figure S5 Relationships between the response ratio (RR) of glycosidase activity and the RR of soil respiration (SR) for the different methods of SR measurement.

        Figure S6 The effects of N addition on soil respiration from previous meta-analyses. Error bars represent bootstrap 95% confidence intervals (CIs). The effect of N addition was considered significant if the CI of the effect size did not overlap zero. The sample size for each variable is shown next to the CI. This figure was redrawn from previous meta-analyses published by (a, b and c) Zhou et al. 2014 , (d) Liu et al. 2010 , (e) Lu et al. 2011 and (f) Janssens et al. 2010 . Ra, autotrophic respiration Rh, heterotrophic respiration SR, soil respiration.

        Figure S7 Relationships between the possible changes in microbial communities and physiology and the response ratios (RR) of glycosidase activity of (a) microbial abundance, (b) bacterial abundance, (c) fungal abundance, (d) fungi/bacteria, (e) microbial biomass carbon (MBC), (f) microbial biomass nitrogen (MBN) and (g) MBC/MBN. The relationships between the changes in microbial communities and physiology induced by N addition and their links with the corresponding changes in soil respiration were synthesized by Treseder et al. ( 2008 ).

        Figure S8 (a) The effects of N addition on the activities of soil oxidative C-acquiring enzymes. Frequency distributions of the response ratios (RR) of (b) oxidative enzymes, (c) phenol oxidase (PO), (d) peroxidase (PER) and (e) polyphenol oxidase (PHO). Error bars represent bootstrap 95% confidence intervals (CIs). The effect of N addition was considered significant if the CI of the effect size did not overlap zero. The sample size for each variable is shown next to the CI. QB and Qw are defined in the Materials and methods section.

        Figure S9 Relationships between the response ratio (RR) of glycosidase activity and the (a) RR of soil total nitrogen (STN), (b) RR of dissolved organic nitrogen (DON), (c) RR of the substrate C:N ratio and (d) substrate C:N ratio.

        Figure S10 Relationships between the response ratio (RR) of glycosidase activity and (a) the substrate pH and (b) the RR of the substrate pH.

        Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.


        Summary

        Initially, an increase in substrate concentration leads to an increase in the rate of an enzyme-catalyzed reaction. As the enzyme molecules become saturated with substrate, this increase in reaction rate levels off. The rate of an enzyme-catalyzed reaction increases with an increase in the concentration of an enzyme. At low temperatures, an increase in temperature increases the rate of an enzyme-catalyzed reaction. At higher temperatures, the protein is denatured, and the rate of the reaction dramatically decreases. An enzyme has an optimum pH range in which it exhibits maximum activity.


        Immobilization of Enzymes and Cells: Methods, Effects and Applications

        Traditionally, enzymes in free solutions (i.e. in soluble or free form) react with substrates to result in products. Such use of enzymes is wasteful, particularly for industrial purposes, since enzymes are not stable, and they cannot be recovered for reuse.

        Immobilization of enzymes (or cells) refers to the technique of confining/anchoring the enzymes (or cells) in or on an inert support for their stability and functional reuse. By employing this technique, enzymes are made more efficient and cost-effective for their industrial use. Some workers regard immobilization as a goose with a golden egg in enzyme technology. Immobilized enzymes retain their structural conformation necessary for catalysis.

        There are several advantages of immobilized enzymes:

        a. Stable and more efficient in function.

        b. Can be reused again and again.

        c. Products are enzyme-free.

        d. Ideal for multi-enzyme reaction systems.

        e. Control of enzyme function is easy.

        f. Suitable for industrial and medical use.

        g. Minimize effluent disposal problems.

        There are however, certain disadvantages also associated with immobilization.

        a. The possibility of loss of biological activity of an enzyme during immobilization or while it is in use.

        b. Immobilization is an expensive affair often requiring sophisticated equipment.

        Immobilized enzymes are generally preferred over immobilized cells due to specificity to yield the products in pure form. However, there are several advantages of using immobilized multi-enzyme systems such as organelles and whole cells over immobilized enzymes. The immobilized cells possess the natural environment with cofactor availability (and also its regeneration capability) and are particularly suitable for multiple enzymatic reactions.

        Methods of Immobilization:

        The commonly employed techniques for immobilization of enzymes are—adsorption, entrapment, covalent binding and cross-linking.

        Adsorption:

        Adsorption involves the physical binding of enzymes (or cells) on the surface of an inert support. The support materials may be inorganic (e.g. alumina, silica gel, calcium phosphate gel, glass) or organic (starch, carboxymethyl cellulose, DEAE-cellulose, DEAE-sephadex).

        Adsorption of enzyme molecules (on the inert support) involves weak forces such as van der Waals forces and hydrogen bonds (Fig. 21.3). Therefore, the adsorbed enzymes can be easily removed by minor changes in pH, ionic strength or temperature. This is a disadvantage for industrial use of enzymes.

        Entrapment:

        Enzymes can be immobilized by physical entrapment inside a polymer or a gel matrix. The size of the matrix pores is such that the enzyme is retained while the substrate and product molecules pass through. In this technique, commonly referred to as lattice entrapment, the enzyme (or cell) is not subjected to strong binding forces and structural distortions.

        Some deactivation may however, occur during immobilization process due to changes in pH or temperature or addition of solvents. The matrices used for entrapping of enzymes include polyacrylamide gel, collagen, gelatin, starch, cellulose, silicone and rubber. Enzymes can be entrapped by several ways.

        1. Enzyme inclusion in gels:

        This is an entrapment of enzymes inside the gels (Fig. 21.4A).

        2. Enzyme inclusion in fibres:

        The enzymes are trapped in a fibre format of the matrix (Fig. 21.4B).

        3. Enzyme inclusion in microcapsules:

        In this case, the enzymes are trapped inside a microcapsule matrix (Fig. 21.4C). The hydrophobic and hydrophilic forms of the matrix polymerise to form a microcapsule containing enzyme molecules inside. The major limitation for entrapment of enzymes is their leakage from the matrix. Most workers prefer to use the technique of entrapment for immobilization of whole cells. Entrapped cells are in use for industrial production of amino acids (L-isoleucine, L-aspartic acid), L-malic acid and hydroquinone.

        Microencapsulation:

        Microencapsulation is a type of entrapment. It refers to the process of spherical particle formation wherein a liquid or suspension is enclosed in a semipermeable membrane. The membrane may be polymeric, lipoidal, lipoprotein-based or non-ionic in nature. There are three distinct ways of microencapsulation.

        1. Building of special membrane reactors.

        3. Stabilization of emulsions to form microcapsules.

        Microencapsulation is recently being used for immobilization of enzymes and mammalian cells. For instance, pancreatic cells grown in cultures can be immobilized by microencapsulation. Hybridoma cells have also been immobilized successfully by this technique.

        Covalent Binding:

        Immobilization of the enzymes can be achieved by creation of covalent bonds between the chemical groups of enzymes and the chemical groups of the support (Fig. 21.5). This technique is widely used. However, covalent binding is often associated with loss of some enzyme activity. The inert support usually requires pretreatment (to form pre-activated support) before it binds to enzyme. The following are the common methods of covalent binding.

        1. Cyanogen bromide activation:

        The inert support materials (cellulose, sepharose, sephadex) containing glycol groups are activated by CNBr, which then bind to enzymes and immobilize them (Fig. 21.6A).

        Some of the support materials (amino benzyl cellulose, amino derivatives of polystyrene, aminosilanized porous glass) are subjected to diazotation on treatment with NaNO2 and HCI. They, in turn, bind covalently to tyrosyl or histidyl groups of enzymes (Fig. 21.6B).

        3. Peptide bond formation:

        Enzyme immobi­lization can also be achieved by the formation of peptide bonds between the amino (or carboxyl) groups of the support and the carboxyl (or amino) groups of enzymes (Fig. 21.6C). The support material is first chemically treated to form active functional groups.

        4. Activation by bi- or poly-functional reagents:

        Some of the reagents such as glutaraldehyde can be used to create bonds between amino groups of enzymes and amino groups of support (e.g. aminoethylcellulose, albumin, amino alkylated porous glass). This is depicted in Fig. 21.6D.

        Cross-Linking:

        The absence of a solid support is a characteristic feature of immobilization of enzymes by cross- linking. The enzyme molecules are immobilized by creating cross-links between them, through the involvement of poly-functional reagents. These reagents in fact react with the enzyme molecules and create bridges which form the backbone to hold enzyme molecules (Fig. 21.7). There are several reagents in use for cross-linking. These include glutaraldehyde, diazobenzidine, hexamethylene diisocyanate and toluene di- isothiocyanate.

        Glutaraldehyde is the most extensively used cross-linking reagent. It reacts with lysyl residues of the enzymes and forms a Schiff’s base. The cross links formed between the enzyme and glutaraldehyde are irreversible and can withstand extreme pH and temperature. Glutaraldehyde cross- linking has been successfully used to immobilize several industrial enzymes e.g. glucose isomerase, penicillin amidase. The technique of cross-linking is quite simple and cost-effective. But the disadvantage is that it involves the risk of denaturation of the enzyme by the poly-functional reagent.

        Choice of Immobilization Technique:

        The selection of a particular method for immobilization of enzymes is based on a trial and error approach to choose the ideal one. Among the factors that decide a technique, the enzyme catalytic activity, stability, regenerability and cost factor are important.

        Immobilization of L-amino acid acylase:

        L-Amino acid acylase was the first enzyme to be immobilized by a group of Japanese workers (Chibata and Tosa, 1969). More than 40 different immobilization methods were attempted by this group. Only three of them were found be useful. They were covalent binding to iodoacetyl cellulose, ionic binding to DEAE-Sephadex and entrapment within polyacrylamide.

        Stabilization of Soluble Enzymes:

        Some of the enzymes cannot be immobilized and they have to be used in soluble form e.g. enzymes used in liquid detergents, some diagnostic reagents and food additives. Such enzymes can be stabilized by using certain additives or by chemical modifications. The stabilized enzymes have longer half-lives, although they cannot be recycled. Some important methods of enzyme stabilization are briefly described.

        Solvent Stabilization:

        Certain solvents at low concentrations stabilize the enzymes, while at high concentrations the enzymes get denatured e.g. acetone (5%) and ethanol (5%) can stabilize benzyl alcohol dehydro­genase.

        Substrate Stabilization:

        The active site of an enzyme can be stabilized by adding substrates e.g. starch stabilizes a-amylase glucose stabilizes glucose isomerase.

        Stabilization by Polymers:

        Enzymes can be stabilized, particularly against increased temperature, by addition of polymers such as gelatin, albumin and polyethylene glycol.

        Stabilization by Salts:

        Stability of metalloenzymes can be achieved by adding salts such as Ca, Fe, Mn, Cu and Zn e.g. proteases can be stabilized by adding calcium.

        Stabilization by Chemical Modifications:

        Enzymes can be stabilized by suitable chemical modifications without loss of biological activity. There are several types of chemical modifications.

        a. Addition of poly-amino side chains e.g. polytyrosine, polyglycine.

        b. Acylation of enzymes by adding groups such as acetyl, propionyl and succinyl.

        Stabilization by Rebuilding:

        Theoretically, the stability of the enzymes is due to hydrophobic interactions in the core of the enzyme. It is therefore, proposed that enzymes can be stabilized by enhancing hydrophobic interactions. For this purpose, the enzyme is first unfold and then rebuilt in one of the following ways (Fig. 21.8).

        1. The enzyme can be chemically treated (e.g. urea and a disulfide) and then refolded.

        2. The refolding can be done in the presence of low molecular weight ligands.

        3. For certain enzymes, refolding at higher temperatures (around 50°C) stabilize them.

        Stabilization by Site-Directed Mutagenesis:

        Site-directed mutagenesis has been successfully used to produce more stable and functionally more efficient enzymes e.g. subtilisin E.

        Immobilization of Cells:

        Immobilized individual enzymes can be successfully used for single-step reactions. They are, however, not suitable for multi-enzyme reactions and for the reactions requiring cofactors. The whole cells or cellular organelles can be immobilized to serve as multi-enzyme systems. In addition, immobilized cells rather than enzymes are sometimes preferred even for single reactions, due to cost factor in isolating enzymes. For the enzymes which depend on the special arrangement of the membrane, cell immobilization is preferred.

        Immobilized cells have been traditionally used for the treatment of sewage. The techniques employed for immobilization of cells are almost the same as that used for immobilization of enzymes with appropriate modifications. Entrapment and surface attachment techniques are commonly used. Gels, and to some extent membranes, are also employed.

        Immobilized Viable Cells:

        The viability of the cells can be preserved by mild immobilization. Such immobilized cells are particularly useful for fermentations. Sometimes mammalian cell cultures are made to function as immobilized viable cells.

        Immobilized Non-viable Cells:

        In many instances, immobilized non-viable cells are preferred over the enzymes or even the viable cells. This is mainly because of the costly isolation and purification processes. The best example is the immobilization of cells containing glucose isomerase for the industrial production of high fructose syrup. Other important examples of microbial biocatalysts and their applications are given in Table 21.5.

        Limitations of Immobilizing Eukaryotic Cells:

        Prokaryotic cells (particularly bacterial) are mainly used for immobilization. It is also possible to immobilize eukaryotic plant and animal cells. Due to the presence of cellular organelles, the metabolism of eukaryotic cells is slow. Thus, for the industrial production of biochemical, prokaryotic cells are preferred. However, for the production of complex proteins (e.g. immunoglobulin’s) and for the proteins that undergo post- translational modifications, eukaryotic cells may be used.

        Effect of Immobilization on Enzyme Properties:

        Enzyme immobilization is frequently associated with alterations in enzyme properties, particularly the kinetic properties of enzymes.

        Some of them are listed below:

        1. There is a substantial decrease in the enzyme specificity. This may be due to conformational changes that occur when the enzyme gets immobilized.

        2. The kinetic constants Km and Vmax of an immobilized enzyme differ from that of the native enzyme. This is because the conformational change of the enzyme will affect the affinity between enzyme and substrate.

        Immobilized Enzyme Reactors:

        The immobilized enzymes cells are utilized in the industrial processes in the form of enzyme reactors. They are broadly of two types — batch reactors and continuous reactors. The frequently used enzyme reactors are shown in Fig. 21.9.

        Batch Reactors:

        In batch reactors, the immobilized enzymes and substrates are placed, and the reaction is allowed to take place under constant stirring. As the reaction is completed, the product is separated from the enzyme (usually by denaturation).

        Soluble enzymes are commonly used in batch reactors. It is rather difficult to separate the soluble enzymes from the products hence there is a limitation of their reuse. However, special techniques have been developed for recovery of soluble enzymes, although this may result in loss of enzyme activity.

        Stirred tank reactors:

        The simplest form of batch reactor is the stirred tank reactor (Fig. 21.9A). It is composed of a reactor fitted with a stirrer that allows good mixing, and appropriate temperature and pH control. However, there may occur loss of some enzyme activity. A modification of stirred tank reactor is basket reactor. In this system, the enzyme is retained over the impeller blades. Both stirred tank reactor and basket reactor have a well-mixed flow pattern.

        Plug flow type reactors:

        These reactors are alternatives to flow pattern type of reactors. The flow rate of fluids controlled by a plug system. The plug flow type reactors may be in the form of packed bed or fluidized bed (Fig. 21.9B and 21.9C). These reactors are particularly useful when there occurs inadequate product formation in flow type reactors. Further, plug flow reactors are also useful for obtaining kinetic data on the reaction systems.

        Continuous Reactors:

        In continuous enzyme reactors, the substrate is added continuously while the product is removed simultaneously. Immobilized enzymes can also be used for continuous operation. Continuous reactors have certain advantages over batch reactors. These include control over the product formation, convenient operation of the system and easy automation of the entire process. There are mainly two types of continuous reactors-continuous stirred tank reactor (CSTR) and plug reactor (PR). A diagrammatic representation of CSTR is depicted in Fig. 21.9D. CSTR is ideal for good product formation.

        Membrane Reactors:

        Several membranes with a variety of chemical compositions can be used. The commonly used membrane materials include polysulfone, polyamide and cellulose acetate. The biocatalysts (enzymes or cells) are normally retained on the membranes of the reactor. The substrate is introduced into reactor while the product passes out. Good mixing in the reactor can be achieved by using stirrer (Fig. 21.10A). In a continuous membrane reactor, the biocatalysts are held over membrane layers on to which substrate molecules are passed (Fig. 21.10B).

        In a recycle model membrane reactor, the contents (i.e. the solution containing enzymes, cofactors, and substrates along with freshly released product are recycled by using a pump (Fig. 21.10C). The product passes out which can be recovered.

        Applications of Immobilized Enzymes and Cells:

        Immobilized enzymes and cells are very widely used for industrial, analytical and therapeutic purpose, besides their involvement in food production and exploring the knowledge of biochemistry, microbiology and other allied specialties. A brief account of the industrial applications of immobilized cells is given in Table 21.5.

        Manufacture of Commercial Products:

        A selected list of important immobilized enzymes and their industrial applications is given in Table 21.6. Some details on the manufacture of L-amino acids and high fructose syrup are given hereunder.

        Production of L-Amino Acids:

        L-Amino acids (and not D-amino acids) are very important for use in food and feed supplements and medical purposes. The chemical methods employed for their production result in a racemic mixture of D- and L-amino acids. They can be acylated to form D, L-acyl amino acids. The immobilized enzyme aminoacylase (frequently immobilized on DEAE sephadex) can selectively hydrolyse D, L-acyl amino acids to produce L-amino acids.

        The free L-amino acids can separated from the un-hydrolysed D-acyl amino acids. The latter can be recemized to D, L-acyl amino acids and recycled through the enzyme reactor containing immobilized aminoacylase. Huge quantities of L-methionine, L-phenylalanine L-tryptophan and L-valine are produced worldwide by this approach.

        Production of High Fructose Syrup:

        Fructose is the sweetest among the monosaccharide’s, and has twice the sweetening strength of sucrose. Glucose is about 75% as sweet as sucrose. Therefore, glucose (the most abundant monosaccharide) cannot be a good substitute for sucrose for sweetening. Thus, there is a great demand for fructose which is very sweet, but has the same calorific value as that of glucose or sucrose.

        High fructose syrup (HFS) contains approximately equivalent amounts of glucose and fructose. HFS is almost similar to sucrose from nutritional point of view. HFS is a good substitute for sugar in the preparation of soft drinks, processed foods and baking.

        High fructose syrup can be produced from glucose by employing an immobilized enzyme glucose isomerase. The starch containing raw materials (wheat, potato, corn) are subjected to hydrolysis to produce glucose. Glucose isomerase then isomerizes glucose to fructose (Fig. 21.11). The product formed is HFS containing about 50% fructose. (Note: Some authors use the term high fructose corn syrup i.e. HFCS in place of HFS).

        This is an intracellular enzyme produced by a number of microorganisms. The species of Arthrobacter, Bacillus and Streptomyces are the preferred sources. Being an intracellular enzyme, the isolation of glucose isomerase without loss of biological activity requires special and costly techniques. Many a times, whole cells or partly broken cells are immobilized and used.

        Immobilized Enzymes and Cells- Analytical Applications:

        In Biochemical Analysis:

        Immobilized enzymes (or cells) can be used for the development of precise and specific analytical techniques for the estimation of several biochemical compounds. The principle of analytical assay primarily involves the action of the immobilized enzyme on the substrate.

        A decrease in the substrate concentration or an increase in the product level or an alteration in the cofactor concentration can be used for the assay. A selected list of examples of immobilized enzymes used in the assay of some substances is given in Table 21.7. Two types of detector systems are commonly employed.

        Thermistors are heat measuring devices which can record the heat generated in an enzyme catalysed reaction. Electrode devices are used for measuring potential differences in the reaction system. In the Fig. 21.12, an enzyme thermistor and an enzyme electrode, along with a specific urease electrode are depicted.


        Enzyme Reactions: Discussion and Results

        Table 1. Solution concentrations, volumes and observations for Experiment 1: Observing the enzyme reaction.

        Test TubedH2Potato ExtractCatecholObservations
        15 ml + 500μl—–500μLSolution turned milky-white to clear. (High substrate)
        25 ml500μl500μlSolution turned yellowish-brown. (High substrate)
        35 ml + 500μl500μl—–Solution is clear, but cloudy-white at the bottom. (Zero substrate)

        *The chemical reaction was observed with the introduction of catechol to the potato extract (tube 2).

        Table 2. Solution concentrations, volumes and observations for Experiment 2: The effect of substrate concentration on enzyme activity.

        Test TubedH2Potato ExtractCatecholObservations
        15 ml + 500μl500μl500μLSolution turned light, yellowish-brown. (High substrate)
        25 ml + 900μl500μl100μlSolution turned translucent peach. (Low substrate, diluted)

        *The reaction time for tube 1 was the fastest due to the high substrate concentration and lower dH20 concentration.

        Table 3. Solution concentrations, volumes and observations for Experiment 3: The effect of enzyme concentration on enzyme activity.

        Test TubedH2Potato ExtractCatecholObservations
        15 ml + 500μl500μl500μLThe solution turned from bluish-green to light, yellowish-brown. (High substrate)
        25 ml + 900μl100μl500μlSlightly cloudier than the original clear solution no real change in color. (High substrate, diluted, low enzyme)

        *Lower Dh20 and higher potato extract concentrations allowed for a faster reaction time

        Table 4. Solution concentrations, Buffer pH, volumes, and observations for Experiment 4: The effect of pH on enzyme activity.

        Buffer pHBuffer VolumedH2Potato ExtractCatecholObservations
        42ml3ml500μl500μlCloudy white
        62ml3ml500μl500μlDark yellow
        82ml3ml500μl500μLOrangish-brown
        102ml3ml100μl500μlTranslucent peach

        *The reaction rate increased as the pH increased, with a pH of 6 being the best buffer for catechol oxidase activity. Increasing the pH past 6 showed a decrease in the reaction rate.

        Table 5. Solution concentrations, temperatures, volumes and observations for Experiment 5: The effect of temperature on enzyme activity.

        Test TubedH2Potato ExtractCatecholObservations
        1 (0°C)5ml500μl500μlReally light yellowish-brown
        2 (15°C)5ml500μl500μlYellowish-peach
        3 (37°C)5ml500μl500μLOrange-peach
        4 (100°C)5ml500μl500μlReally light peach

        *The fastest reaction rate was observed at 37°C. The colder the temperature (0°C – 15°C), the slower the reaction rate. Enzyme denaturation was observed at 100°C.

        Table 6. Solution concentrations, volumes and observations for Experiment 6: Inhibitor Effects – Inhibiting the Action of Catechol Oxidase

        Test TubedH2Potato ExtractPTUCatecholObservations
        15ml + 1ml500μl—–500μlYellowish-peach (Control)
        25ml + 500μL500μl500μl500μlReally light peach
        35ml500μl500μl500μLCloudy, clear-white

        *The fastest, and most pronounced reaction was observed in tube 1 (the solution without phenylthiourea)

        Enzyme Lab Discussion

        For the first experiment, Observing the Enzyme Reaction, it was hypothesized that the enzyme reaction would only occur in the second test tube due to the fact that it was the only tube to contain both the enzyme and substrate. As expected, the solution in tube 2 was the only solution to show the characteristic yellow-brown pigment of benzoquinone production, which was caused by the potato extract converting its catechol into the new product.

        In experiment 2, The Effect of Substrate Concentration on Enzyme activity, the hypothesis was that the tube with the higher substrate concentration would show a faster and more pronounced chemical reaction than the tube with less catechol.

        The hypothesis was supported by the fact that the higher catechol concentration in tube 1 allowed for a similar result to tube 2 from experiment 1, the only difference being that the extra 5mL of dH20 diluted some of the yellowish-brown color observed in the first reaction.

        While there was a chemical reaction observed in tube 2 (experiment 2), it was much slower (with a translucent peach pigment) due to lower a catechol concentration and a higher dH20 concentration. The higher the concentration of catechol, the more benzoquinone that can be produced.

        It was hypothesized in experiment 3, The Effect of Enzyme Concentration on Enzyme Activity, that the higher the concentration of enzyme in the solution, the faster and more pronounced the chemical reaction would be.

        This hypothesis was able to be accepted based on the rate at which the tube with the higher potato extract concentration reacted. Tube 1 had 400μL more potato extract and 400μL less dH20 than tube 2. Because enzymes are biological catalysts that speed up chemical reaction time, the solution in tube 1 quickly changed from a bluish-green pigment, to the yellowish-brown color associated with benzoquinone.

        The lower concentration of potato extract and a higher concentration of dH20 in tube 2 showed no change in color, other than the cloudiness of the potato extract itself.

        In experiment 4, The Effect of pH on Enzyme Activity, the initial hypothesis was that the lower the pH level of the buffer added to the solution, the quicker the reaction rate would be. This hypothesis was not supported by the data observed because higher acidity levels actually slowed the production of benzoquinone – which was the opposite of what was predicted.

        The solution with a pH buffer of 4 remained cloudy white, while the solution with a 6 pH buffer turned yellowish-brown. As the pH increased, the benzoquinone production rate increased. While lower pH buffers proved to be too acidic, more neutral buffers allowed for the best environment for catechol oxidase activity.

        Buffer pH levels higher than 6 showed a slower and less pronounced chemical reaction as well – illustrating the enzyme reaction’s need for neutrality.

        The hypothesis for experiment 5, The Effect on Temperature on Enzyme Activity, was that extremely low temperature would slow the rate of benzoquinone production, while extremely high temperatures would cause the enzymes to denature. This hypothesis was supported by the rate at which the solutions at 0°C – 15°C slowly reacted, and the rate at which the solution at 37°C quickly produced benzoquinone.

        After five minutes at each solution’s designated temperature, the colder solutions barely started to change color, while the warmer temperatures quickly reacted – so much so that at 100°C, the enzymes denatured and the solution began to pale in pigment. Colder temperatures slowed the movement of molecules in the solutions, while warmer temperatures (not including 100°C) allowed for a better environment for catechol oxidase activity.

        For experiment 6, Inhibitor Effects – Inhibiting the Action of Catechol Oxidase, it was hypothesized that the addition of phenylthiourea (PTU) would keep the enzyme reaction from occurring. The hypothesis was able to be accepted due to the fact that the tubes which contained the PTU showed very little change in pigment.

        Tube 1 served as the control, which showed the production of benzoquinone (yellowish-brown color) and allowed for comparison between the three solutions. Considering PTU is a non-competitive inhibitor, tubes 2 and 3 contained solutions that prevented the enzyme from catalyzing the reaction, regardless of whether or not the substrate was bound to the active site.

        The only real issue with any of the 6 experiments was the unsupported hypothesis for the Effect of pH on the Enzyme Activity experiment. I must have tied the preservative nature of benzoquinone with how acidic lemon juice keeps apples from turning brown, so I assumed a low pH would increase the reaction rate. In reality, acidity slows the reaction rate – which is why the apples don’t change color.

        In conclusion, these experiments have shown that benzoquinone production can only occur with the presence of both an enzyme and substrate. Factors such as substrate and enzyme concentration, pH, temperature, and the presence of noncompetitive inhibitors can affect enzyme reaction. High substrate concentration will allow for greater benzoquinone production, while high enzyme concentration will speed up the reaction rate – and vise versa.

        In order for enzyme reaction to rapidly occur, it must be done in an environment where the pH is as close to neutral as possible, with the reaction rate slowing in both highly acidic or basic solutions. The same goes for temperature – extremely high or extremely cold temperatures can decrease enzyme reaction rates, or cause the enzymes to denature altogether.

        The introduction of a noncompetitive inhibitor (such as phenylthiourea) allows it to bind to the allosteric site on the enzyme, which keeps the reaction from occurring (regardless of the enzyme or substrate concentration).



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