amylase enzyme experiment ph

Practical Biology

A collection of experiments that demonstrate biological concepts and processes.

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Practical Work for Learning

Published experiments

Investigating the effect of ph on amylase activity, class practical.

Measure the time taken for amylase to completely break down starch , by withdrawing samples at 10 second intervals and noting the time at which the solution no longer gives a blue-black colour with iodine solution (but the iodine solution remains orange). Use buffers to provide solutions at different pHs . Calculate the rate of this enzyme controlled reaction by calculating 1÷ time.

Lesson organisation

This procedure is simple enough for individuals to carry out if you have enough dimple tiles. If you choose to investigate five pHs, then groups of five students could complete the investigation by working together and pooling results.

Apparatus and Chemicals

For each group of students:.

Syringes, 5 cm 3 , 2 (1 for starch, 1 for amylase)

Iodine solution in a dropper bottle ( Note 4 )

Test tube rack

Test tube, 1 for each pH to be tested

Dimple tile or white tiles

Teat pipette

For the class – set up by technician/ teacher:

Amylase 1% (or 0.5%) ( Note 1 )

Starch 1% (or 0.5%) ( Note 2 )

Buffer solutions covering a range of pH, each with a labelled syringe/ plastic pipette ( Note 3 )

Health & Safety and Technical notes

Amylase solution and iodine solution are low hazard once made up. Wear eye protection when handling iodine solution. Hazards of buffers may vary. See CLEAPSS Recipe card or supplier’s information and see Note 3 .

Read our standard health & safety guidance

1 Amylase (See CLEAPSS Hazcard and Recipe card) The powdered enzyme is HARMFUL, but solutions less than 1% are LOW HAZARD. It is wise to test, well in advance, the activity of the stored enzyme at its usual working concentration to check that substrates are broken down at an appropriate rate. Enzymes may degrade in storage and this allows time to adjust concentrations or to obtain fresh stocks. Amylase will slowly lose activity, so it is best to make up a fresh batch for each lesson; batches may vary in activity and results collected on different days will not be comparable. The optimum temperature for your enzyme will be listed on the supplier’s label.

Using saliva: the CLEAPSS Laboratory Handbook provides guidance on precautions to take (including hygiene precautions) in order to use saliva safely as a source of amylase. This has the advantage of being cheaper, not requiring technicians to make up fresh solutions each lesson, it is directly interesting to students, and salivary amylase is reliable. It also provides an opportunity to teach good hygiene precautions – including ensuring that students use only their own saliva samples (provide small beakers to spit into); that students are responsible for rinsing their own equipment; and that all contaminated glassware is placed in a bowl or bucket of sodium chlorate(I) before technicians wash up.

2 Starch suspension – make fresh. Make a cream of 5 g soluble starch in cold water. Pour into 500 cm 3 of boiling water and stir well. Boil until you have a clear solution. Do not use modified starch.

3 Buffers: (See CLEAPSS Recipe card) If you make universal buffer it will contain sodium hydroxide at approximately 0.25 M, and should be labelled IRRITANT. Refer to other relevant Hazcards if you choose to make other buffers, or to supplier’s information if you purchase buffer solutions/ tablets. ( Note 1 )

4 Iodine solution (See CLEAPSS Hazcard and Recipe card). A 0.01 M solution is suitable for starch testing. Make this by 10-fold dilution of 0.1 M solution. Once made, the solution is a low hazard but may stain skin or clothing if spilled.

Ethical issues

There are no ethical issues associated with this procedure.

SAFETY: All solutions once made up are low hazard. Wear eye protection, as iodine may irritate eyes.

Preparation

a Check the speed of the reaction with the suggested volumes of reactants to be used – 2 cm 3 of starch: 2 cm 3 of amylase: 1 cm 3 of buffer at pH 6. Ideally the reaction should take about 60 seconds at this pH: this is the usual optimum for amylase (see note 1). If the reaction is too fast, either reduce the enzyme volume or increase the starch volume. If the reaction is too slow, increase the enzyme volume or concentration or reduce the starch volume or concentration.

Investigation

b Place single drops of iodine solution in rows on the tile.

c Label a test tube with the pH to be tested.

d Use the syringe to place 2 cm 3 of amylase into the test tube.

e Add 1 cm 3 of buffer solution to the test tube using a syringe.

f Use another syringe to add 2 cm 3 of starch to the amylase/ buffer solution, start the stop clock and leave it on throughout the test. Mix using a plastic pipette.

g After 10 seconds, use the plastic pipette to place one drop of the mixture on the first drop of iodine. The iodine solution should turn blue-black. If the iodine solution remains orange the reaction is going too fast and the starch has already been broken down. Squirt the rest of the solution in the pipette back into the test tube.

h Wait another 10 seconds. Then remove a second drop of the mixture to add to the next drop of iodine.

i Repeat step h until the iodine solution and the amylase/ buffer/ starch mixture remain orange.

j You could prepare a control drop for comparison with the test drops. What should this contain?

k Count how many iodine drops you have used, each one equalling 10 seconds of reaction time.

l Repeat the whole procedure with another of the pH buffers to be used, or pool the class results.

m Consider collecting repeat data if there is time.

n Plot a graph of time taken to break down starch against pH, or calculate the rate of reaction and plot rate against pH.

Teaching notes

This is a straightforward practical giving reliable, unambiguous results. The main errors will be in the order of mixing the enzyme/ substrate/ buffer, or a delay in sampling so that the reaction time is under-estimated or rate is over-estimated. Temperature variation affects enzyme activity, so results collected on different days are not comparable.

Health and safety checked, September 2008

http://rsc.org/Education/Teachers/Resources/cfb/enzymes.htm Royal Society of Chemistry: Chemistry for Biologists: Enzymes

A clear and thorough presentation of information about enzymes as chemical catalysts and the factors affecting their activity.

(Website accessed October 2011)

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Last Update: November 12, 2023 .

• Introduction

Amylase is a digestive enzyme predominantly secreted by the pancreas and salivary glands and is present in other tissues at minimal levels. [1] Amylase was initially described in the early 1800s and is one of the pioneering enzymes to undergo scientific investigation. Although this enzyme was originally termed diastaste , it was later renamed "amylase" in the early 20th century. [2]

The primary role of amylases is to break down the glycosidic bonds within starch molecules, transforming complex carbohydrates into simpler sugars. Amylase enzymes are categorized into 3 main classes—alpha-, beta-, and gamma amylases—each targeting distinct segments of the carbohydrate molecule. [2] Alpha amylase is present in humans, animals, plants, and microbes, whereas beta amylase is primarily found in microbes and plants. Gamma amylase, on the other hand, can be located in both animals and plants. [3]

In 1908, a study by Wohlgemuth identified the presence of amylase in urine, paving the way for its application as a diagnostic laboratory test. Amylase is a frequently ordered standard diagnostic test, often combined with lipase, particularly when acute pancreatitis is suspected in patients. [2]

• Etiology and Epidemiology

Elevated amylase levels may manifest in various conditions, including pancreatic, salivary, and intestinal diseases, as well as decreased metabolic clearance and macroamylasemia. [4] Approximately 11% to 13% of patients experiencing non-pancreatic abdominal pain exhibit elevated pancreatic enzyme levels. [5]  On at least one occasion, 60% of asymptomatic HIV-positive patients displayed abnormal amylase or lipase measurements. [6] Upon admission, 26 out of 208 patients (12.5%) with acute abdominal pain unrelated to pancreatic issues presented with elevated serum amylase levels. [5]

Abnormally elevated amylase levels are observed in 35% of patients with liver disease. [7] Furthermore, elevated amylase levels are found in around 16% to 25% of diabetic ketoacidosis cases. [8] [9]  In a group of 74 patients with surgically resectable lung cancer, 13 showed hyperamylasemia. [10]

• Pathophysiology

Amylase is a heterogeneous calcium-dependent metalloenzyme with a diverse molecular weight range, typically between 54 and 62 kDa. The compact size of amylase facilitates efficient filtration through the glomeruli. [1] Amylase is eliminated through both the renal system and the reticuloendothelial system. [3]  This enzyme exists as 2 isoenzymes—pancreatic (P-type) and nonpancreatic (S-type)—which are products of 2 closely linked loci on chromosome 1. Additional amylase heterogeneity results from allelic variation, with 12 alleles for the S-type and 6 alleles for the P-type. [11] Both types of amylase also undergo post-translational modifications involving deamidation, glycosylation, and deglycosylation, giving rise to various isoforms. Amylase exhibits a broad tissue distribution, with the most significant levels of P- and S-type activities being found in the exocrine pancreas and salivary glands, respectively. [12]

P-type amylase is synthesized by pancreatic acinar cells and released into the intestinal tract through the pancreatic duct system. The enzymatic activity of P-type amylase is optimized under the slightly alkaline conditions of the duodenum. [13]  Meanwhile, the salivary glands host the highest S-type amylase activity, initiating starch hydrolysis during mastication in the mouth and the passage through the esophagus. However, this action is terminated upon exposure to stomach acid.

S-type amylase is also detectable in extracts from testes, ovaries, fallopian tubes, Mullerian ducts, striated muscle, lungs, and adipose tissue, as well as in bodily fluids such as semen, colostrum, tears, and milk. Approximately 25% of plasma amylase is excreted by the kidneys, with the majority being reabsorbed within the proximal tubules. [14]  The liver is believed to be the primary organ responsible for amylase elimination, leading to a half-life of approximately 10 hours. Serum amylase is intricately controlled within the body, [15] with a delicate balance between its production and clearance rates. Elevated amylase levels can result from heightened production, whether originating in the pancreas or outside it, or from a diminished clearance rate.

Genetic regulation is likely to play a crucial role in the preliminary determination of salivary amylase. [1]  In newborns, the predominant amylase isozymes detected in urine originate from saliva, and as development progresses, both salivary and pancreatic amylase isozymes become more prominent. The functional integrity of amylase is entirely dependent on the presence of calcium. [16]  However, complete functionality is achieved only in the presence of specific anions, including chloride, bromide, nitrate, or monohydrogen phosphate. Chloride and bromide are the most effective activators. The pH optimum for amylase activity falls within the range of 6.9 to 7.0. [17]

The analyte amylase is an endoglycosidase enzyme belonging to the hydrolase class, and it catalyzes the hydrolysis of 1,4-α-glucosidic linkages between adjacent glucose units in complex carbohydrates. [18]  Notably, straight-chain (linear) polyglucans, such as amylose, and branched-chain polyglucans, such as amylopectin and glycogen, are hydrolyzed at different rates. [19]  In the case of amylose, the enzyme cleaves the chains at alternate α-1,4-hemiacetal (-C-O-C-) links, thereby forming maltose and some residual glucose. In the case of branched polyglucans, the enzyme generates maltose, glucose, and a residue of limit dextrins. Notably, the enzyme does not target the α-1,6-linkages at the branch points. [2]

• Specimen Requirements and Procedure

Either serum or heparinized plasma can serve as suitable samples. Nevertheless, a particular study indicated that when using dry-film technology for measurement, heparinized plasma samples yielded significantly higher results than serum samples. [20]  As amylase has an absolute requirement for calcium ions, it is noteworthy that chelating anticoagulants, such as citrate, oxalate, and EDTA, cannot be used when collecting plasma for amylase measurements. However, urine specimens without preservatives, obtained through random or timed collections, are considered valid samples.

Amylase is occasionally measured in ascitic, peritoneal, or pleural fluid, and its presence in these fluids can suggest pancreatitis or the presence of a tumor. In serum, amylase activity remains stable for up to 4 days at room temperature, 2 weeks at 5 °C, 1 year at −28 °C, or 5 years at −75 °C. Urine specimens should be analyzed within 12 hours at room temperature or within 5 days when stored at 5 °C. Notably, urine should not be subjected to freezing. [21]

• Diagnostic Tests

Over the years, amylase has played a central role in diagnosing acute pancreatitis. Amylase can be assessed through either a blood test or a urine test. In the case of a urine test, it can be conducted through a clean catch or a 24-hour urine collection. [3]  The normal range for serum amylase can vary between different laboratories. Notably, it is clinically significant to differentiate pancreatic amylase from other amylase isoforms. If amylase levels are elevated while lipase remains within the normal range, it might indicate an issue originating outside the pancreas. [15]

The lipase-to-amylase ratio may help distinguish between gallstone-induced pancreatitis and alcoholic pancreatitis. Gallstones tend to cause more significant elevations in amylase levels, whereas alcohol typically results in higher elevations in lipase levels. A lipase-to-amylase ratio exceeding 2 exhibits 91% sensitivity and 76% specificity for diagnosing alcoholic pancreatitis, whereas a ratio exceeding 5 demonstrates 31% sensitivity and nearly 100% specificity for identifying alcoholic pancreatitis. [22] Elevated alanine transaminase levels reaching 3 times the normal range are highly specific for diagnosing gallstone pancreatitis. Although the combined measurement of serum amylase and lipase enhances specificity compared to either test in isolation, it does not notably improve sensitivity. [23]

• Testing Procedures

The amylase test is conducted using semiautomatic or fully automated analyzers that operate on the principles of photometry. Photometry entails the measurement of light absorption across the ultraviolet (UV), visible (VIS), and infrared (IR) spectrum. This measurement is used to quantify the concentration of an analyte in a solution or liquid. Photometers utilize a specific light source and detectors, such as photodiodes, photoresistors, or photomultipliers, to transform the light transmitted through a sample solution into a corresponding electrical signal. Photometry applies Beer–Lambert law to calculate coefficients obtained from the transmittance measurement. [24] A test-specific calibration process is utilized to establish a correlation between the absorbance and analyte concentration to achieve highly accurate measurements. [25]

P-type amylase can be differentiated from S-type amylase by selective inhibition of S-type using a wheat germ inhibitor, temperature inhibition, immunoprecipitation, or immunoinhibition with a monoclonal antibody. However, only the methods based on selective inhibition by monoclonal antibodies have demonstrated adequate precision, reliability, practicality, and analytical speed, enabling the reliable measurement of P-type amylase. [26]  The amylase isoforms can be separated by isoelectric focusing, ion-exchange chromatography, or gel/capillary electrophoresis by electrophoretic endosmosis. [27]

• Interfering Factors

Amylase assays typically resist hemoglobin, bilirubin, or triglyceride interference. However, collecting specimens in tubes containing oxalate, citrate, or EDTA may result in falsely decreased values due to the chelation of necessary amylase cofactors. [3] Certain medications, such as aspirin, morphine, antiretrovirals, and estrogen-containing drugs, can potentially affect serum amylase levels. [28]

Elevated amylase activity in serum can be attributed to a condition known as macroamylasaemia, which is characterized by the formation of macromolecular complexes. These complexes typically involve immunoglobulins— predominantly IgA or IgG—although self-polymerization or association with other proteins can also occur. [29] Typically, these complexes maintain their enzymatic activity but cannot be effectively filtered by the renal glomeruli. As a result, this condition leads to delayed clearance and increased serum amylase activity. [30]  This benign condition has been reported in up to 1.5% of hospitalized patients, accounting for as much as 28% of chronic and otherwise unexplained hyperamylasemia cases. [31]  

Macroamylasaemia is a condition associated with autoimmunity, malignancy, cardiovascular disease, diabetes mellitus, and malabsorptive disorders. Macroamylasemia should be considered when evaluating asymptomatic patients with elevated serum amylase levels. [32]  Notably, no specific treatment is mandated for this condition, as it is typically benign. [33]  

The circulating pancreatic amylase levels are higher in female subjects with O blood type than those with A blood type, where pancreatic amylase levels tend to be lower. [34]  Psychosocial stress contributes to elevated salivary amylase levels even in healthy individuals, potentially resulting in higher total serum amylase. However, it remains unconfirmed through clinical studies whether psychosocial stress has a long-term effect on serum amylase levels. [35]

In pancreatitis accompanied by hypertriglyceridemia cases, serum amylase levels may mistakenly appear within the normal range. This discrepancy is linked to an inhibitor associated with elevated triglycerides, which interferes with the assay for the enzyme. By diluting the serum, the inhibitory effect can be lessened, allowing for a recalculation of serum amylase levels to reveal the actual concentration. [36]

• Results, Reporting, and Critical Findings

The reference intervals for amylase can vary among different assay methods due to variations in the substrates used and reagent preparations. [37]  A patient's blood test values should be interpreted based on the reference values established by the specific laboratory conducting the test. Furthermore, it is recommended for each laboratory to establish its unique reference interval based on its specific testing methodology. [38]  A significant proportion of subjects of African and Asian origin exhibit S-type amylase activity levels exceeding the reference interval established for white populations. This can lead to an elevated total amylase measurement without indicating any pathological condition. [39]

The blood amylase activity in newborns is approximately 18% of that in adults. Serum amylase activity does not show significant differences between males and females. In healthy adults, P-AMY typically accounts for about 40% to 50% of the total amylase activity in serum. In most children less than 6 months of age, serum P-AMY activity is not detectable. However, its activity gradually increases after this period and reaches adult concentrations by the age of 5, which corresponds to the postnatal development of exocrine pancreatic function. [40] Therefore, using this enzyme for diagnosing acute pancreatitis in young children should be avoided. [41]

Currently, there is no internationally standardized reference range for amylase levels, with the reference range varying from as low as 20 U/L to as high as 300 U/L. However, elevated amylase levels exceeding 3 times the upper limit of normal (URL) strongly indicate the likelihood of acute pancreatitis. [42] Amylase levels below this threshold are associated with other medical conditions. Abnormally low amylase levels, while less common, can be observed in conditions such as cystic fibrosis, chronic pancreatitis, diabetes mellitus, obesity, and among individuals who smoke. [43]  Clinicians should be aware of such potential causes when interpreting low amylase activity in patients. [44]

A finding of persistently elevated total amylase and normal lipase should raise the possibility of macroamylasemia. Screening tests such as the amylase creatinine clearance ratio (ACCR) or polyethylene glycol precipitation can be valuable in identifying macroamylase. [45]  ACCR can be easily calculated from paired random urine and serum amylase and creatinine measurements. Although an ACCR lower than 1% indicates the possibility of macroamylasemia, each laboratory needs to assess the suitability of these expected values for its unique patient population and, if required, establish its own reference ranges. [46]  An ACCR value greater than 5% suggests acute pancreatitis. [47]  Notably, the ACCR is also known to be increased in diabetic ketoacidosis, renal disease, and after surgical procedures. [48]

• Clinical Significance

Amylase is primarily used for diagnosing pancreatic diseases and is a frequently measured enzyme owing to the accessibility of cost-effective and readily automated methods. Although amylase is a sensitive marker for acute pancreatitis, its lack of specificity is evident, as it can be elevated in numerous conditions unrelated to the pancreas. [49] Pancreatitis can be defined by the presence of at least 2 out of the 3 criteria—abdominal pain, serum amylase, and/or lipase levels—exceeding 3 times the URL and characteristic findings of pancreatitis on abdominal imaging. Therefore, the clinical significance of amylase has been a subject of inquiry. [50]  In cases of elevated amylase levels and insufficient evidence of pancreatitis, healthcare professionals should contemplate alternative causes of hyperamylasemia. [51]  

Amylase does not help predict the severity of an acute pancreatic episode or for monitoring the condition. The magnitude of the increase in serum enzyme activity is not correlated with the severity of pancreatic involvement. However, a more substantial rise in amylase levels does increase the probability of acute pancreatitis. The limited specificity of total amylase measurement has spurred interest in directly measuring P-type amylase, rather than total enzyme activity, for the differential diagnosis of patients experiencing acute abdominal pain. [19] Utilizing the best decision limit, which corresponds to an activity 3 times the URL, the specificity of P-type amylase in diagnosing acute pancreatitis exceeds 90%. Furthermore, P-AMY significantly enhances sensitivity in the late detection of this condition. One week after onset, P-type amylase values remain elevated in 80% of patients with uncomplicated pancreatitis, while only 30% still exhibit increased total amylase activity. [52]  This long-standing increase in P-type amylase activity in serum renders the traditional measurement of total amylase in urine redundant. This test is performed to achieve better diagnostic sensitivity in the late phase of pancreatitis. [53]

Amylase inhibitors such as acarbose have been used in treating type 2 diabetes and have demonstrated the ability to lower hemoglobin A1C levels and reduce peak postprandial glucose. Acarbose has also proven effective in enhancing the remission of dumping syndrome in patients who have undergone bariatric procedures. In addition, this drug has demonstrated the capacity to reduce the risk of cardiovascular disease by slowing the progression of carotid artery thickening. [54] Elevated amylase levels can be observed in a wide array of conditions. Therefore, clinicians need to follow a well-defined, systematic approach when hyperamylasemia is detected. This approach can prevent unnecessary hospitalization and ensure timely and appropriate treatment. [55] Biliary tract diseases, including cholecystitis, can lead to up to a 4-fold increase in serum P-type amylase activity due to primary or secondary pancreatic involvement. [56]

Various intra-abdominal events can result in a substantial rise in serum P-type amylase activities, often reaching up to 4-fold or even greater. These elevations may be attributed to the leakage of P-type amylase from the intestine into the peritoneal cavity and subsequently into the circulation. [57] In cases of renal insufficiency, serum amylase activity increases proportionally with the degree of renal impairment, typically not exceeding 5 times the URL. [58]

Cases of amylase-producing multiple myeloma have been described. Increased amylase activity in most patients results from salivary-type hyperamylasemia—sialyl salivary type. [59]  A common feature of the myeloma cell lines associated with hyperamylasemia is the presence of a chromosome 1 translocation, which harbors the gene for amylase. The link does not seem to be specific to any particular immunoglobulin class. The onset of hyperamylasemia is reported to be associated with rapid disease progression, extensive bone destruction, and increased mortality. Consequently, serum amylase activity may serve as a valuable prognostic "tumor marker" in patients with multiple myeloma, with activity decreasing in response to treatment and increasing during periods of relapse. [60] [61] The amylase isoenzyme in cases of ruptured ectopic pregnancy is not well characterized. In severe cases that are diagnosed late, the elevated isoenzyme may be P-AMY due to pancreatic involvement associated with peritonitis, despite the presence of S-AMY in the fallopian tube. [62]

In some cases, patients with pheochromocytoma or paraganglioma exhibit hyperamylasemia, usually the salivary isotype. In these instances, hyperamylasemia might be associated with a hypertensive crisis and vasoconstriction, resulting in tissue hypoxia rather than being a consequence of tumor secretion. This elevation in amylase activity is often transient. [63]  Salivary-type hyperamylasemia has been documented in various conditions unrelated to salivary gland disorders. These include diabetic ketoacidosis, pneumonia, and postoperative states following a diverse array of surgical procedures, including extra abdominal procedures such as postcoronary bypass. [64]

Benign pancreatic hyperenzymemia or Gullo syndrome was first described by Lucio Gullo and is characterized by elevated serum levels of amylase, pancreatic isoamylase, lipase, and trypsin activities in asymptomatic individuals with no evidence of pancreatic disease on imaging. The syndrome occurs sporadically or in a familial form, and amylase activity may exhibit notable fluctuations, occasionally normalizing transiently in some cases. [65] The etiology of benign pancreatic hyperenzymemia does not appear to involve CFTR , SPINK1 , or PRSS1 gene mutations. In addition, this condition cannot be attributed to mutations in genes known to be associated with pancreatitis or other PRSS1 / SPINK1 genes. [66] Researchers discovered that approximately one-third of patients with chronic non-pathological pancreatic hyperenzymemia had notably elevated fecal calprotectin concentrations. They recommended investigating this discovery to explore the potential connection between intestinal ecology and alterations in pancreatic enzymes. [67]

Damage of salivary glands, leading to salivary hyperamylasemia, has been observed following trauma or surgical procedures on the salivary gland. Furthermore, radiation to the neck area involving the parotid gland can lead to duct obstruction or calculi formation in the salivary glands. [14] Chronic alcoholism and anorexia nervosa can also lead to subclinical damage to the salivary glands. In individuals with alcoholism, 10% of patients exhibit salivary amylase activity, which is 3 times higher than normal and may be associated with chronic liver disease. [68]

Hyperamylasemia in anorexia nervosa is associated with vomiting, and the detection of elevated salivary amylase levels may hint at concealed vomiting. [69]  However, pancreatitis can develop in these patients, especially during the refeeding process. Therefore, it may be warranted to measure plasma lipase and/or amylase isoenzymes to distinguish between pancreatitis and salivary hyperamylasemia. [70]

Hyperamylasemia may be associated with various tumors, either due to ectopic production of the enzyme by the tumors or possibly as an inflammatory response by the tumor cells, thereby leading to the significant release of the enzyme normally produced in these tissues into the bloodstream. [71]  The raised isoenzyme is almost exclusively salivary type in ovarian, lung cancer, multiple myeloma, and pheochromocytoma. [72]  Amylase-producing tumors of the lung are rare, accounting for only 1% to 3% of all lung carcinomas, and in such instances, the salivary amylase isotype is typically present. These amylase-producing lung carcinomas are primarily adenocarcinomas, although hyperamylasemia has also been documented in cases of small-cell carcinoma. [73]  Amylase activity has been suggested as a valuable tumor marker for monitoring the patient's treatment in amylase-producing lung carcinoma cases. [74] In one study, it was noted that 39% of patients with ovarian carcinoma exhibited hyperamylasemia, primarily of the salivary type, suggesting that salivary amylase could be a useful indicator for assessing the effectiveness of radiotherapy in this context. [75]

Gut diseases, including mucosal inflammatory disease of the small intestine, mesenteric infarction, intestinal obstruction, appendicitis, and peritonitis, generally lead to elevated P-type isoamylase levels due to increased absorption of amylase from the intestinal lumen. [76]  Gut perforation is associated with the leakage of intestinal contents into the peritoneum, leading to inflammation and the absorption of amylase across the inflamed peritoneum, which can result in hyperamylasemia. Acidosis can lead to hyperamylasemia and may arise from 2 sources—ketoacidosis, which results in increased S-type and P-type isoamylases, or nonketotic acidosis, which leads to increased S-type isoamylase. [77] Postoperative amylase increases can lead to elevated S-type and P-type isoamylase levels, with a more common increase observed in salivary amylase. [78]  This elevation may occur after procedures involving extracorporeal circulation or nonabdominal surgery. For instance, approximately 30% of patients undergoing cardiac surgery exhibit elevated S-type isoamylase. [79]

Rare cases of hyperamylasemia have been reported in association with systemic lupus erythematosus, as well as with ciprofloxacin treatment. [80]  Additional causes contributing to hyperamylasemia include pneumonia (increased salivary amylase), cerebral trauma, burns, abdominal aortic aneurysms (increased pancreatic amylase), drugs (increased salivary and/or pancreatic amylase), anorexia nervosa and bulimia (increased salivary amylase), non-pathological factors (increased salivary and/or pancreatic amylase), and organophosphate poisoning. Post-procedure balloon-assisted enteroscopy has also been associated with elevated amylase levels. Therefore, it is recommended to measure pancreatic amylase levels rather than total amylase levels following these procedures. [3] [81]  Elevated pancreatic enzymes can be found in critically injured trauma patients, even in the absence of true pancreatitis. [82]

• Quality Control and Lab Safety

The purpose of a clinical laboratory test is to assess the pathophysiological state of a specific patient, aiding in the process of diagnosis, offering guidance or supervision of treatment, and evaluating the likelihood or advancement of the disease. [83]  Implementing a quality management system (QMS) is paramount in upholding the precision and reliability of laboratory tests. [84] Internal quality control (IQC) is a cornerstone within the QMS of a clinical laboratory, systematically monitoring and verifying the accuracy and precision of laboratory test results. [85]

All aspects of laboratory operation, including IQC, must adhere to written standard operating procedures (SOPs). [86]  The SOP for quality control (QC) should include all aspects of the program, including the selection of QC materials, determination of statistical parameters to describe method performance, criteria for accepting QC results, measurement frequency of QC materials, corrective actions when problems arise, and the documentation and review processes. The SOP should specify authorized personnel responsible for setting acceptable control limits and interpretive rules for result release, reviewing performance parameters, including statistical QC results, and granting authorization for exceptions to or modifications of established QC policies and procedures. [87]

The QC samples are periodically measured using the same method as clinical samples, and the results are analyzed to ensure that the measurement procedure meets performance requirements suitable for patient care. [85]  If the QC result falls within the acceptable limits of the known value, it verifies that the measurement procedure is stable and performing as expected. This indicates that results for patient samples can be reported with a high probability that they are suitable for clinical use. If a QC result falls outside the acceptable limits, it indicates that the measurement procedure is not performing correctly. As a result, patient sample results cannot be reported, and corrective action is necessary. After taking corrective measures, patient sample measurements are repeated, along with QC samples, to ensure accurate and reliable results. [88] Adhering to good laboratory practice involves verifying that a method is performing correctly at the moment when patient results are being measured. [89]

For non-waived tests, laboratory regulations mandate the analysis of a minimum of 2 control material levels at least once every 24 hours. In cases where it is necessary, laboratories may choose to assay QC samples more frequently to ensure the accuracy of results. In addition, it is essential to analyze QC samples after the calibration or maintenance of an analyzer to verify the correct method performance. [85] To reduce the frequency of QC testing for assays that have manufacturer-recommended intervals less frequent than those required by regulatory agencies, such as once per month, laboratories have the option to implement an individualized QC plan (IQCP). This plan includes a risk assessment, which evaluates potential sources of errors across all testing phases, and the implementation of a tailored QC strategy to minimize the risk of errors. [90]  Westgard multi-rules are used to assess the QC runs. In case of a rule violation, appropriate corrective and preventive actions must be executed before patient testing. [91]

Proficiency testing (PT) is a program designed to assess method performance by comparing results with those from other laboratories analyzing the same set of samples. [92]  PT providers distribute a set of samples among a group of laboratories for this purpose. Each laboratory analyzes the PT samples in the same manner as they would with patient samples and then reports the results to the PT provider for evaluation. The PT provider assigns a target value to the samples and evaluates whether the individual laboratory's results align closely with the target value, indicating acceptable method performance. [93] The acceptable performance criteria for amylase assay, as defined by the Clinical Laboratory Improvement Amendments (CLIA) and College of American Pathologists (CAP) proficiency program, are within ±30% of the mean value of laboratory peer groups. [94]

If an unacceptable PT result is identified, the method must be investigated for possible causes, and any necessary corrective action must be taken. [95] Furthermore, even when a PT result is within acceptability criteria, it is good laboratory practice to investigate PT results that deviate by more than approximately 2.5 standard deviation index (SDI) from the peer group mean. When the SDI is 2.5, there is only a 0.6% probability that the result will fall within the expected distribution for the peer group. Therefore, it is reasonable to suspect that a method problem may need to be corrected. [96]  Investigative steps, reviewed data, and conclusions derived from the review must be documented in a written report of the unacceptable PT result, which the Director of the laboratory should review. [93]

Ensuring lab safety is paramount in clinical laboratories, where precise and reliable results are essential for patient diagnoses and treatment plans. [97] Maintaining a secure environment requires strict adherence to safety protocols. This includes using personal protective equipment (PPE), including lab coats, gloves, goggles, and masks, to protect against potential hazards. [98]  Chemical safety measures include appropriate labeling, storage, and handling of chemicals, ensuring that hazardous substances are stored in designated areas. Similarly, biological safety is of utmost importance, requiring the use of biosafety cabinets and stringent protocols for managing potentially infectious materials. Regular maintenance and calibration of equipment, as well as staff training on safe operation, are crucial in preventing accidents and ensuring accurate results. In case of emergencies, it is essential to have well-defined procedures for handling accidents, spills, or exposures, along with a clear understanding of the locations of safety equipment and evacuation routes. [99]  

Fire safety precautions, including the availability of fire extinguishers and proper storage of flammable materials, are of utmost importance. In addition, electrical safety measures, such as equipment maintenance and ensuring proper grounding, help prevent electrical hazards. [100]  Safe handling and disposal of sharps and appropriate management of chemical and biohazardous waste are essential to protect lab personnel. Rigorous training on safety protocols, in addition to comprehensive documentation of incidents, procedures, and training, fosters a culture of safety and ensures compliance with regulatory standards. [101] In conclusion, a comprehensive approach to lab safety is indispensable for preserving both the accuracy of test results and the well-being of laboratory personnel in clinical settings. 

• Enhancing Healthcare Team Outcomes

Effective communication among interprofessional healthcare team members is crucial when laboratory results indicate a potential non-pancreatic cause for amylase level abnormalities. Moreover, it is equally important for the healthcare team to understand how various conditions can affect amylase levels. [102] Lipase is generally favored over amylase because of its greater specificity. Lipase levels usually stay elevated for up to 2 weeks, whereas amylase concentrations tend to return to normal within 5 days. Therefore, in cases where there is a gap between symptom onset and the patient seeking medical attention, amylase is less clinically valuable compared to lipase. [18]

The 2013 American College of Gastroenterology guidelines highlight that simultaneously ordering both lipase and amylase is neither cost-effective nor therapeutically advantageous. Furthermore, the healthcare providers emphasize that ordering amylase alone is an unreliable approach and does not enhance diagnostic efficiency compared to lipase. [12]  Hence, when lipase testing is available, including amylase testing not only increases the cost for the patient but also provides minimal value in aiding the diagnosis of pancreatitis. [103]

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Disclosure: Ololade Akinfemiwa declares no relevant financial relationships with ineligible companies.

Disclosure: Muhammad Zubair declares no relevant financial relationships with ineligible companies.

Disclosure: Thiruvengadam Muniraj declares no relevant financial relationships with ineligible companies.

This book is distributed under the terms of the Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International (CC BY-NC-ND 4.0) ( http://creativecommons.org/licenses/by-nc-nd/4.0/ ), which permits others to distribute the work, provided that the article is not altered or used commercially. You are not required to obtain permission to distribute this article, provided that you credit the author and journal.

• Cite this Page Akinfemiwa O, Zubair M, Muniraj T. Amylase. [Updated 2023 Nov 12]. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2024 Jan-.

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Properties of an amylase from thermophilic Bacillu s SP

Raquel vieira de carvalho, thamy lívia ribeiro côrrea, júlia caroline matos da silva, luciana ribeiro coutinho de oliveira mansur, meire lelis leal martins.

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Corresponding Author. Mailing address: Centro de Ciências e Tecnologias Agropecuárias, Universidade Estadual do Norte Fluminense. Av. Alberto Lamego, 2000. CEP. 28015-620. Campos dos Goytacazes, RJ. Brasil. Tel.: (24) 7263880; Fax: (24) 7263875. E-mail: [email protected]

Received 2007 Jan 19; Revised 2007 Jul 16; Accepted 2008 Jan 19; Issue date 2008 Jan-Mar.

All the content of the journal, except where otherwise noted, is licensed under a Creative Commons License

α-Amylase production by thermophilic Bacillus sp strain SMIA-2 cultivated in liquid cultures containing soluble starch as a carbon source and supplemented with 0.05% whey protein and 0.2% peptone reached a maximum activity at 32 h, with levels of 37 U/mL. Studies on the amylase characterization revealed that the optimum temperature of this enzyme was 90°C. The enzyme was stable for 1 h at temperatures ranging from 40-50°C while at 90°C, 66% of its maximum activity was lost. However, in the presence of 5 mM CaCl2, the enzyme was stable at 90°C for 30 min and retained about 58% residual activity after 1 h. The optimum pH of the enzyme was found to be 8.5. After incubation of enzyme for 2 h at pH 9.5 and 11.0 was observed a decrease of about 6.3% and 16.5% of its original activity. At pH 6.0 the enzyme lost about 36% of its original activity. The enzyme was strongly inhibited by Co 2+ , Cu 2+ and Ba 2+ , but less affected by Mg 2+ , Na + and K + . In the presence of 2.0 M NaCl, 63% of amylase activity was retained after 2 h incubation at 45°C. The amylase exhibited more than 70% activity when incubated for 1 h at 50°C with sodium dodecyl sulphate. However, very little residual activity was obtained with sodium hypochlorite and with hydrogen peroxide the enzyme was completely inhibited. The compatibility of Bacillus sp SMIA-2 amylase with certain commercial detergents was shown to be good as the enzyme retained 86%, 85% and 75% of its activity after 20 min incubation at 50°C in the presence of the detergent brands Omo ® , Campeiro ® and Tide ® , respectively.

Keywords: α-Amylase, Thermophilic bacterium, Bacillus sp

INTRODUCTION

The majority of industrial enzymes used currently belong to the hydrolase group, which are active on many natural substrates. For years microorganisms have been the principal source of many different enzymes, which were identified after extensive research and currently find their main uses in industrial applications ( 5 ).

α-Amylase (EC 3.2.1.1) is an important enzyme used in the industry and accounts for approximately 25% of the enzyme market ( 24 ). Thermostable α-amylases have extensive commercial applications in starch processing, brewing and sugar production, desizing in textile industries and in detergent manufacturing processes ( 10 , 15 ).

Amylases can be obtained from several sources ( 1 , 26 ). They are usually produced by bacteria belonging to the genus Bacillus for industrial applications such as B . amyloliquefaciens , B. stearothermophilus , B. subtilis and Bacillus licheniformis ( 22 ), the latter now being of greater industrial importance ( 7 ). The properties of each α-amylase such as thermostability, pH profile, pH stability, and Ca-independency must be matched to its application. For example, α-amylases used in starch industry must be active and stable at low pH but in detergent industry at high pH values ( 19 ).

In this article, the properties of an thermostable α-amylase produced by the thermophilic Bacillus sp strain SMIA-2, isolated from soil in Campos dos Goytacazes City, Rio de Janeiro, Brazil, are reported.

MATERIALS AND METHODS

Microorganism and culture conditions.

The bacterial strain used in this study was Bacillus sp SMIA-2, a thermophilic strain isolated from a local soil sample ( 20 ). Production of α-amylase was carried out in a medium containing (g/L of distilled water): KCl-0.3, MgSO 4 -0.5, K 2 HPO 4 -0.87, CaCl 2 -2.2x10 -3 , ZnO-2.5x10 -3 , FeCl 3 .6H 2 O-2.7x10 -2 , MnCl 2 .4H 2 O-1.0x10 -2 , CuCl 2 .2H 2 O-8.5x10 -4 , CoCl 2 .6H 2 O-2.4x10 -3 , NiCl 3 .6H 2 O-2.5x10 -4 , H 3 BO 3 -3.0x10 -4 , whey protein-0.5, peptone-2.0 and soluble starch-5.0. The pH was adjusted to 7.5 with NaOH and this basal medium was sterilized by autoclaving at 121°C for 15 minutes. The medium (25 mL in 250 mL Erlenmeyer flasks) was inoculated with 1 mL of an overnight culture and incubated at 50°C in a orbital shaker (Thermo Forma, Ohio, USA) operated at 180 min -1 for 48 h. Triplicate aliquots withdraw at regular intervals and analysed for growth (OD 600 nm) and pH. The cells and residues were removed from the culture broth by centrifugation and concentrated by precipitating the enzyme with ammonium sulfate (60% saturation). The precipitate was dissolved and dialyzed overnight against 10 mM phosphate buffer, pH 7.0. The dialyzed enzyme was then concentrated by lyophilization and used for subsequent studies.

Amylase assay

The activity of α-amylase was assayed by incubating 0.5 mL of the diluted enzyme (0.55 mg.mL -1 ) with 0.5 mL soluble starch (0.5%, w/v) prepared in 0.05 M Tris-HCl buffer, pH 8.5. After incubation at 90°C for 10 minutes the reaction was stopped and the reducing sugars released were assayed colorimetrically by the addition of 1 mL of 3-5-dinitrosalicylic acid reagent ( 16 ). One enzyme activity unit (U) was defined as the amount of enzyme releasing 1 μmol of glucose from the substrate in 1 minute at 90°C.

Effect of the pH

The effect of pH on the activity of α-amylase was measured by incubating 0.5 mL of the diluted enzyme (0.55 mg.mL -1 ) and 0.5 mL of buffers presenting pH from 6.0 to 12.0, containing 0.5% soluble starch for 10 minutes at 90°C. The buffers used were: phosphate buffer, pH 6.0-8.0; Tris-HCl buffer, pH 8.5-9.0 and glycine-NaOH, pH 10.0-12.0. The stability of the enzyme at different pH values was also studied by incubating the enzyme at various pH values ranging from 6.0 to12.0 for 2 h and then estimating the residual activity.

Effect of temperature

The effect of temperature on the enzyme activity was determined by performing the previously described standard assay procedure for 10 minutes at pH 8.5 within a temperature range of 40-100°C. Thermostability was determined by incubation of the lyophilized enzyme at temperatures ranging from 40-100°C for 1 h in a constant-temperature water bath. After treatment the residual enzyme activity was assayed.

Effect of metal ions

The effect of metal ions on α-amylase activity was measured incubating the enzyme at 90°C for 2 min with various metal ions at a concentration of 5 mM and 10 mM. The enzyme assay was carried out in the presence of Ca 2+ , Zn 2+ , Mn 2+ , Ni 2+ and Ba 2+ chlorides, and Mg 2+ , Fe 2+ , Co 2+ and Cu 2+ sulphates.

Salt tolerance test

The enzyme was incubated in 50 mM Tris-HCl buffer (pH 8.5) containing NaCl at concentrations of 0 to 5 M for 120 minutes at 45°C. The activity of the enzyme was measured in the same way as mentioned earlier.

Effect of inhibitors and oxidizing agents

The enzyme was incubated with 5 mM sodium dodecylsulphate (SDS), sodium hypochlorite (NaClO), hydrogen peroxide (H 2 O 2 ) and EDTA (Ethylene diaminetetraacetic acid) at 50°C for 1 h. After incubation, the residual activity was determined by the standard enzyme assay. The results were recorded as the percentage of residual activity calculated with reference to activity controls incubated in the absence of these compounds.

Evaluation of enzyme for use in detergent formulation

The detergent brands used were Ariel ® , Biz ® , Cheer ® , Tide ® , Campeiro ® and Omo ® . They were diluted in double distilled water to a final concentration of 7 mg.mL -1 to simulate washing conditions and heated at 100°C for 15 minutes to inactivate the enzymes that could be part of their formulation. The detergents were added to the reaction mixture and the reaction was carried out under standard assay conditions. To determine the stability of Bacillus sp SMIA-2 amylase in the presence of the different detergents, an amylase concentration of 1 mg.mL -1 was added in detergent solution and incubated at 50°C for 60 minutes. Aliquots (0.5 mL) were taken at different time intervals and the residual activity determined at 90°C and compared with the control sample incubated at 50°C without any detergent ( 2 , 21 ).

RESULTS AND DISCUSSION

Enzymatic production.

Fig. 1 shows the time-course profile of amylase production by Bacillus sp strain SMIA-2 in basal medium containing soluble starch (0.5%) as a carbon source and supplemented with whey protein (0.05%) and peptone (0.2%). α-amylase production by Bacillus sp strain SMIA-2 began in the exponential growth phase, reaching a maximum at 32 h, with levels of 37 U/mL. Subsequently amylase levels remained more or less the same up to 36 h and then dropped to 30 U/mL at 48 h. The pH of the medium dropped as cells started to grow, but as soon as enzyme production was initiated, the pH started to rise. This may indicate that some organic nitrogen was being consumed. The end of enzyme production was signalled by a slight decrease of pH. Thus the pH profile provides a useful means of monitoring the production process.

Time course of α-amylase production by Bacillus sp strain SMIA-2 grown at 50°C on soluble starch (0.5%) in shake flasks. Results represent the means of three separate experiments, and bars indicated ± 1 standard deviation. Absence of bars indicates that errors were smaller than symbols.

The effect of pH on α-amylase activity as a function of pH is shown in Fig. 2 . Optimum pH was found to be 8.5. The enzyme activity at pH 7.0 and 11.0 were 72% and 81.4% of that at pH 8.5, respectively. After incubation of the enzyme solution for 2h at pH 6.0-12.0, the original activity at pH 9.5 decreased by 63% and at pH 11.0 the decrease was 16.5%. However, at pH 6.0, the original activity decreased 36%. These results suggest that the activity of the enzyme is higher in alkaline pH, making this enzyme attractive for the detergent industry. α-amylases produced by other Bacillus sp have shown optimum activities at pH values as low as 3.5 or as high as 12 ( 3 , 13 ).

Optimum pH (□) and stability (■) of α-amylase produced by Bacillus sp strain SMIA-2 grown at 50°C for 48 h. Relative activity is expressed as a percentage of the maximum (100% of enzyme activity = 36.1U/mL).

Fig. 3 shows the activity of the lyophilized enzyme preparation assayed at temperatures ranging from 40°C-100°C at pH 8.5 and a substrate concentration of 0.5%. Enzyme activity increased with temperature within the range of 40°C to 90°C. A reduction in enzyme activity was observed at values above 90°C. The optimum temperature of this α-amylase was 90°C, which was higher or similar to that described for other Bacillus α-amylases ( 4 , 6 , 12 , 13 , 24 ).

Optimum temperature (■) and stability temperature (□) of α-amylase produced by Bacillus sp strain SMIA-2 grown at 50°C for 48 h. Relative activity is expressed as a percentage of the maximum. 100% of enzyme activity = 36.4 U/mL.

The residual activities of crude amylase incubated at different temperatures for a period of 1 h were estimated at optimum temperature. The enzyme was stable for 1 h at temperatures ranging from 40-50°C while at 80°C, 18% of its maximum activity was lost. At 90°C, 66% of its maximum activity was lost, however, the addition of 5 mM CaCl2 improved the thermostability of the amylase, as indicated in Fig. 4 . The enzyme was stable at 90°C for 30 min and retained about 50.6% residual activity after 1 h. The requirement of Ca 2+ by α-amylases for their stability at higher temperature has also been reported for Bacillus sp. I-3 ( 9 ), Bacillus sp. ANT-6 ( 6 ), B. subtilis ( 18 ), Bacillus clausii BT-21 ( 8 ), and Bacillus licheniformis ( 14 ). The stabilizing effect of Ca 2+ on thermostability of the enzyme can be explained by the salting out of the hydrophobic residues by Ca 2+ in the protein, thus causing the adoption of a compact structure ( 27 ).

Termostability of amylase at 90°C in the presence (■) or absence of Ca 2+ (□). 100% of enzyme activity = 50.5 U/mL.

The amylase did not require any specific ion for catalytic activity ( Fig. 5 ). Najafi et al. (2005) observed that the α-amylase from Bacillus subtilis AX20 did not have an obligate requirement for divalent metal ions to be active and its activity was not stimulated in the presence of metal ions. A stronger inhibitory effect was observed in the presence of Cu 2+ , Co 2+ , Ba 2+ and Mn 2+ . On the other hand, no significant inhibition of activity was observed in the presence of 5 mM of Mg 2+ , Na + , Zn 2+ and K + . Some amylases are metalloenzymes, containing a metal ion with a role in catalytic activity. The inhibition of Bacillus sp strain SMIA-2 amylase by Co 2+ , Cu 2+ and Ba 2+ ions could be due to competition between the exogenous cations and the protein-associated cation, resulting in decreased metalloenzyme activity ( 15 ).

Effect of metal ions on amylase activity. The activity is expressed as a percentage of the activity level in the absence of metal ion. 100% of enzyme activity = 40.62 U/mL.

In the presence of 2.0 M NaCl, 63.4% of amylase activity was retained after 2 h incubation at 45°C. The α-amylase produced by Bacillus sp. MD 124 was stable in NaCl solution and retained 75% of its original activity in 5 M after 24 h of incubation ( 11 ).

Effect of inhibitors and some oxidizing agents on enzyme activity and compatibility with various commercial detergents

Besides pH and temperature stability, a good detergent amylase should also be stable to various detergent ingredients, such as surfactants, chelators and oxidants ( 23 ). The amylase from Bacillus sp SMIA-2 retained more than 70% activity after incubated for 1 h at 50°C with sodium dodecyl sulphate, an anionic detergent. However, very little residual activity was obtained with sodium hypochlorite. The enzyme was completely inhibited by hydrogen peroxide. The amylase from Bacillus sp. PN5 retained more than 80% activity when incubated with sodium perborate and sodium dodecyl sulphate and more than 70% activity when incubated with hydrogen peroxide for an hour. However, very little residual activity was obtained with sodium hypochlorite ( 23 ). Regarding to the effect of EDTA, the enzyme retained 53% of its activity when incubated for 20 min at 50°C. The α-amylase from Bacillus sp PS-7 retained almost 100% activity when 1mM EDTA was added to the reaction mixture ( 25 ).

Effect of NaCl concentration on α-amylase produced by Bacillus sp strain SMIA-2 grown at 50°C for 24 h. Relative activity is expressed as a percentage of the maximum. 100% of enzyme activity = 34.8 U/mL.

Effect of inhibitors and oxidizing agents on α-amylase activity. The activity is expressed as a percentage of the activity level in the absence of inhibitors and oxidizing agents. 100% of enzyme activity = 31.9 U/mL.

Studies on the effect of detergents on amylase activity ( Fig. 8a ) show that the enzyme activity increased in the presence of Omo ® and was almost the same as that of the control in the presence of Campeiro ® . On the other hand, the enzyme was severely inhibited in the presence of Biz ® and Cheer ® . The compatibility of Bacillus sp. SMIA-2 amylase with certain commercial detergents was shown to be good as the enzyme retained 86%, 85% and 75% of its activity after 20 minutes incubation at 50°C in the presence of the detergent brands Omo ® , Campeiro ® and Tide ® respectively ( Fig. 8b ). After 40 minutes, the enzyme retained 49%, 47% and 40% activity at 50°C in the presence of the detergents Tide ® , Omo ® and Campeiro ® respectively.

Compatibility of α-amylase activity from Bacillus sp. SMIA-2 with commercial detergents (♦ Ariel ® , ■ Biz ® , Δ Cheer ® , x Tide ® , ▅ Campeiro®, ● Omo®). The activity is expressed as a percentage of the activity level in the absence of detergents (100% of enzyme activity = 47 U/mL).

ACKNOWLEDGMENTS

The authors are highly thankful to the Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro for financial support.

Propriedades de uma amilase de um termofílico Bacillus sp

A produção de α-amilase por um termofilico, Bacillus sp SMIA-2, cultivado em meio líquido contendo amido solúvel como fonte de carbono, alcançou uma atividade máxima de 37 U/mL em 32 horas. Estudos sobre a caracterização da amilase revelaram que a temperatura ótima desta enzima foi 90°C. A enzima foi estável por 1 hora a temperaturas de 40 e 50°C enquanto a 90°C, 66% da atividade máxima foi perdida. Entretanto, na presença de 5 mM de CaCl-2, a enzima foi estável a 90°C por 30 minutos e manteve cerca de 58% de sua atividade residual por 1 hora. O pH ótimo da enzima encontrado foi de 8.5. Após a incubação da enzima por 2 horas a pH 9.5 e 11.0 foi observado um decréscimo de aproximadamente 6.3% e 16.5% da atividade original. Em pH 6.0 a enzima perdeu cerca de 36% de sua atividade original. A enzima foi fortemente inibida por Co 2+ , Cu 2+ , e Ba 2+ , porém pouco afetada por Mg 2+ , Na + e K + . Na presença de 2.0 M de NaCl, 63% da atividade da amilase foi mantida após 2 horas de incubação a temperatura de 45°C. A amilase exibiu atividade acima de 70% quando incubada por 1 hora a 50°C em presença de sódio dodecil sufato (SDS). Entretanto, uma baixa atividade residual foi obtida quando na presença do hipoclorito de sódio e uma completa inibição quando a enzima foi incubada em peróxido de hidrogênio. A compatibilidade da amilase produzida pelo Bacillus sp SMIA-2, em relação a alguns detergentes comerciais mostrou que a enzima manteve 86%, 85%, e 75% da atividade após 20 minutos de incubação a 50°C na presença dos detergentes Omo ® , Campeiro ® e Tide ® , respectivamente.

Palavras-chave : α-Amilase, Bactéria termofílica, Bacillus sp.

• 1. Aquino A.C.M.M., Jorge J.A., Terenzi H.F., Polizeli M.L.T.M. Studies on a thermostable α-amylase from the thermophilic fungus Scytalidium thermophilum. Appl. Microbiol. Biotechnol. 2003;61:323–328. doi: 10.1007/s00253-003-1290-y. [ DOI ] [ PubMed ] [ Google Scholar ]
• 2. Banerjee U.C., Sani R.K., Azmi W., Soni R. Thermoestable alkaline protease from Bacillus brevis and its characterization as a laundry detergent additive. Process Biochem. 1999;35:213–219. [ Google Scholar ]
• 3. Bernhardsdotter E.C.M.J., Ng J.D., Garriott O.K., Pusey M.L. Enzymic properties of an alkaline chelator-resistant α-amylase from an alkaliphilic Bacillus sp. isolate L1711. Process Biochem. 2005;40:2401–2408. [ Google Scholar ]
• 4. Bertoldo C., Antranikian G. Starch hydrolyzing enzymes from thermophilic archae and bacteria. Curr. Opin. Chem. Biol. 2002;6:151–160. doi: 10.1016/s1367-5931(02)00311-3. [ DOI ] [ PubMed ] [ Google Scholar ]
• 5. Bon E.P.S. A tecnologia enzimática no Brasil. ENZITEC. 1995;95:9–14. [ Google Scholar ]
• 6. Burhan A., Nisa U., Gokhan C., Omer C., Ashabil A., Osman G. Enzymatic properties of a novel thermostable, thermophilic, alkaline and chelator resistant amylase from an alkaliphilic Bacillus sp. isolate ANT-6. Process Biochem. 2003;38:1397–403. [ Google Scholar ]
• 7. Declerck N., Machius M., Wiegand G., Huber R., Gaillardin C. Probing structural determinants specifying high thermostability in Bacillus licheniformis α-amylase. J. Mol. Biol. 2000;31:1041–1057. doi: 10.1006/jmbi.2000.4025. [ DOI ] [ PubMed ] [ Google Scholar ]
• 8. Duedahl-Olesen L., Kragh K.M., Zimmermann W. Purification and characterisation of a malto-oligosaccharide-forming Amylase from amylase active at high pH from Bacillus clausii BT-21. Carbohydr. Res. 2000;329:97–107. doi: 10.1016/s0008-6215(00)00153-1. [ DOI ] [ PubMed ] [ Google Scholar ]
• 9. Goyal N., Gupta J.K., Soni S.K. A novel raw starch digesting thermostable α-amylase from Bacillus sp.I-3 and its use in the direct hydrolysis of raw potato starch. Enzyme and Microb. Technol. 2005;37:723–734. [ Google Scholar ]
• 10. Hendriksen H.V., Pedersen S., Bisgard-Frantzen H. A process for textile warp sizing using enzymatically modified starches. Patent Application. WO 99/35325. [ Google Scholar ]
• 11. Jana M., Pati B. Thermostable, salt-tolerant α-amylase from Bacillus sp. MD-124. J. Bras. Microbiol. 1997;37:323–326. [ Google Scholar ]
• 12. Jin F., Li Y., Zhang C., Yu H. Thermostable α-amylase and a-galactosidase production from the thermophilic and aerobic Bacillus sp. JF strain. Process Biochem. 2001;36:559–564. [ Google Scholar ]
• 13. Konsula Z., Liakopoulou-Kyriakides M. Hydrolysis of starches by the action of an α-amylase from Bacillus subtilis. Process Biochem. 2004;39:1745–1749. [ Google Scholar ]
• 14. Krishnan T., Chandra A.K. Purification and characterization of α-amylase from B. licheniformis CUMC 305. Appl. Environ. Microbiol. 1983;46:430–437. doi: 10.1128/aem.46.2.430-437.1983. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ]
• 15. Lévêque E., Janecek S., Haye B., Belarbi A. Thermophilic archaeal amylolytic enzymes. Enzyme Microbiol. Technol. 2000;26:3–14. [ Google Scholar ]
• 16. Miller G.L. Use of dinitrosalicylic acid reagent for determination of reducing sugars. Anal. Chem. 1959;3:426–428. [ Google Scholar ]
• 17. Najafi M.F., Deobagkar D., Deobagkar D. Purification and characterization of an extracellular α-amylase from Bacillus subtilis AX20. Protein Expr. and Purif. 2005;41(2):349–354. doi: 10.1016/j.pep.2005.02.015. [ DOI ] [ PubMed ] [ Google Scholar ]
• 18. Nielsen A.D., Pusey M.L., Fuglsang C.C., Westh P. A proposed mechanism for the thermal denaturation of a recombinant Bacillus halmapalus α-amylase the effect of calcium ions. Biochim. Biophys. Acta. 2003;1652:52–63. doi: 10.1016/j.bbapap.2003.08.002. [ DOI ] [ PubMed ] [ Google Scholar ]
• 19. Nielsen J.E., Borchert T.V. Protein engineering of bacterial α-amylases. Biochim Biophys. Acta. 2000;1543:253–274. doi: 10.1016/s0167-4838(00)00240-5. [ DOI ] [ PubMed ] [ Google Scholar ]
• 20. Nunes A.S., Martins M.L.L. Isolation, properties and kinetics of growth of a thermophilic Bacillus. Braz. J. Microbiol. 2001;32:271–275. [ Google Scholar ]
• 21. Phadatare S.U., Deshpande V.V., Srinivasan M.C. High activity alkaline protease from Conidiobolus coronatus (NCL 86.8.20): Enzyme production and compatibility with commercial detergents. Enzyme Microbiol. Technol. 1993;15:72–76. [ Google Scholar ]
• 22. Sajedi R.H., Naderi-Manesh H., Khajeh K., Ahmadvand R., Ranjbar B., Asoodeh A., Moradian F. A Ca-independent α-amylase that is active and stable at low pH from the Bacillus sp. KR-8104. Enzyme and Microb. Technol. 2005;36:666–671. [ Google Scholar ]
• 23. Saxena R.K., Dutt K., Agarwal L., Nayyar P. A highly thermostable and alkaline amylase from a Bacillus sp. PN5. Bioresour. Technol. 2007;98(2):260–265. doi: 10.1016/j.biortech.2006.01.016. [ DOI ] [ PubMed ] [ Google Scholar ]
• 24. Sidhu G.S., Sharma P., Chakrabarti T., Gupta J.K. Strain improvement for the production of a thermostable α-amylase. Enzyme and Microb. Technol. 1997;21:525–530. [ Google Scholar ]
• 25. Sodhi H.K., Sharma K., Gupta J.K., Soni S.K. Production of a thermostable α-amylase from Bacillus sp. PS-7 by solid state fermentation and its synergistic use in the hydrolysis of malt starch for alcohol production. Process Biochem. 2005;40:525–534. [ Google Scholar ]
• 26. Van der Maarel M.J., Van der Veen B., Uitdehaag J.C., Leemhuis H., Dijkhuizen L. Properties and applications of starch-converting enzymes of the α-amylase family. J. Biotechnol. 2002;94:137–155. doi: 10.1016/s0168-1656(01)00407-2. [ DOI ] [ PubMed ] [ Google Scholar ]
• 27. Volkin D.B., Klibanov A.M. Thermal destruction processes in proteins involving cysteine. J. Biol. Chem. 1989;262:2945–2950. [ PubMed ] [ Google Scholar ]
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The determinants of α-amylase pH–activity profiles

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Jens Erik Nielsen, Torben V. Borchert, Gerrit Vriend, The determinants of α-amylase pH–activity profiles, Protein Engineering, Design and Selection , Volume 14, Issue 7, July 2001, Pages 505–512, https://doi.org/10.1093/protein/14.7.505

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The glycosyl hydrolases present a large family of enzymes that are of great significance for industry. Consequently, there is considerable interest in engineering the enzymes in this family for optimal performance under a range of very diverse conditions. Until recently, tailoring glycosyl hydrolases for specific industrial processes mainly involved stability engineering, but lately there has also been considerable interest in engineering their pH–activity profiles. We mutated four neutral residues (N190, F290, N326 and Q360) in the chimeric Bacillus Ba2 α-amylase to both charged and neutral amino acids. The results show that the pH–activity profile of the Ba2 α-amylase can be changed by inserting charged residues close to the active site. The changes in the pH–activity profile for these neutral → charged mutations do not, however, correlate with the predictions from calculations of the p K a values of the active site residues. More surprisingly, the neutral → neutral mutations change the pH–activity profile as much as the neutral → charged mutations. From these results, it is concluded that factors other than electrostatics, presumably the dynamic aspects of the active site, are important for the shape of the pH–activity profiles of the α-amylases.

α-Amylases are used in several industrial processes such as starch liquefaction, laundering, dye removal and feed pre-processing (Guzman-Maldonado and Paredes-Lopez, 1995). Several of these processes take place at pH values which are very different from those where the α-amylases perform optimally ( Nielsen and Borchert, 2000 ) and consequently there is great interest in changing the pH–performance profile of the α-amylases ( Nielsen et al. , 1999a ; Shaw et al. , 1999 ) and related enzymes ( Fang and Ford, 1998 ; Wind et al. , 1998 ). The engineering of pH–activity profiles for these enzymes has proven particularly difficult and in the present paper we present experiments and calculations that suggest several reasons for the difficulties encountered when engineering the pH–activity profiles of α-amylases and related enzymes.

The α-amylases consist of three domains called A, B and C. Domain A is a TIM-barrel [(α/β) 8 -barrel], which is interrupted by an irregular β-domain (domain B) inserted between the third β-strand and the third α-helix of the TIM-barrel. Domain C is a Greek key motif which is located approximately on the opposite side of the TIM-barrel with respect to domain B. The active site is situated in a cleft at the interface between domains A and B.

The active site consists of a large number of charged groups, among which are three acids essential to catalytic activity. Two of these, Asp231 and Glu261 (numbering according to the BLA sequence), are believed to be the two catalytic groups. Asp231 is the catalytic nucleophile, while evidence has been presented for Glu261 being the catalytic hydrogen donor ( McCarter and Withers, 1996 ; Uitdehaag et al. , 1999 ). The third essential acid (Asp328) is believed to assist catalysis by hydrogen bonding to the substrate and by elevating the p K a of Glu261 ( Klein et al. , 1992 ; Knegtel et al. , 1995 ; Strokopytov et al. , 1995 ; Uitdehaag et al. , 1999 ).

The catalytic reaction is believed to consist of three steps ( Sinnot, 1990 ; McCarter and Withers, 1994 ; Davies and Henrissat, 1995 ; McCarter and Withers, 1996 ) (see Figure 1 ). Step one is the protonation of the glycosidic oxygen by the proton donor (Glu261). This is followed by a nucleophilic attack on the C1 of the sugar residue in position –1 by Asp231 [nomenclature as described by Davies et al. ( Davies et al. , 1997 )]. After the aglycone part of the substrate has left, a water molecule is activated, presumably by the now deprotonated Glu261. This water molecule hydrolyses the covalent bond between the nucleophile oxygen (of Asp231) and the C1 of the sugar residue in position –1, thus completing the catalytic cycle.

α-Amylase catalysis is thought to be limited by the protonation of the nucleophile at low pH and by the deprotonation of the hydrogen donor at high pH ( Qian et al. , 1994 ; Strokopytov et al. , 1995 ; Fang and Ford, 1998 ). The rate-limiting step at intermediate pH values is not known. This step has to be largely independent of pH since many α-amylases have a pH–activity profile with a `flat top'. Binding of the substrate and release of the product are expected to be independent of pH and it has therefore been speculated that the rate-limiting step is either substrate binding or product release at intermediate pH values.

Changing the pH–activity profile

If we accept the assumption that α-amylase catalysis is limited at low pH by protonation of the nucleophile (Asp231) and at high pH by deprotonation of the hydrogen donor (Glu261), then the pH–activity profile is determined by the p K a values of the these two active site groups ( Kyte, 1995 ). We consider two types of pH–activity profiles: k cat profiles and the k cat / K m profiles. The k cat profile of an enzyme is determined by the p K a values of the active site groups in the substrate bound form of the enzyme ( Kyte, 1995 ).

Since the substrate is present in high concentrations in the majority of industrial processes where α-amylases are used, it is obviously the k cat profile and not the k cat / K m profile that limits the activity of the enzyme. In order to change the k cat profile, we therefore have to identify the factors that determine the p K a values of the active site residues when the substrate is bound. The protein environment certainly is important and although not much is known about the effect of the substrate, it is undoubtedly also important, since shifts in α-amylase pH–activity profiles have been measured when changing the substrate ( Keating et al. , 1998 ). Also, elegant experiments with Bacillus circulans xylanase have shown that the p K a value of the catalytic hydrogen donor `cycles' during the catalytic reaction in order to fulfil its dual role as hydrogen donor and hydrogen acceptor ( McIntosh et al. , 1996 ).

Changing the substrate is unfortunately not an option when trying to engineer the pH–performance profile of an α-amylase for a specific industrial process and we are therefore left with the option of changing the enzyme (in our case by site-directed mutagenesis) and in that way changing the p K a values of the active site groups and thereby the k cat profile.

Changing pK a values

The p K a value of a residue depends on the free energy difference between the neutral and the charged states of the residue in the protein. The free energy difference between these two states is influenced both by desolvation effects and by the charges and dipoles in the protein and the substrate. Desolvation effects and the interaction with dipoles are short-ranged and therefore result mainly from interactions with residues in the immediate environment. Mutations that aim at changing the p K a value of a titratable group by changing these energies should therefore be placed in the vicinity of the titratable group.

Mutations that introduce or remove unit charges can be placed further away from the titratable group, because the interaction energy between a titratable group and a unit charge (which may itself be a part of another titratable group) is less dependent on distance.

The direction of the expected p K a shift when perturbing the environment of an uncoupled titratable group is summarized in Figure 2 . Using this figure, we can therefore easily predict the direction of an α-amylase pH–activity profile shift, if we assume that the active site residues of the α-amylases are relatively uncoupled or if the charged residue is placed a short distance from the active site. Thus, placing a negative charge near the hydrogen donor will elevate the p K a of the hydrogen donor and give a basic shift in the high-pH limb of the pH–activity profile. Placing a positive charge close to the hydrogen donor will shift the basic limb of the pH–activity profile to more acidic values.

Identifying the determinants of α-amylase pH–activity profiles

Previously ( Nielsen et al. , 1999a ), we constructed 15 mutants in and around the active site of Bacillus licheniformis α-amylase (BLA), in an attempt to change its pH–activity profile. The mutations in the active site were conservative in nature and were an attempt to change the pH–activity profile of the enzyme by changing the hydrogen-bond interactions and the solvent accessibility of the active site residues. The mutations outside the active site aimed at changing the pH–activity profile by introducing or removing a charge and in this way perturbing the p K a values of the active site acids.

The mutations in the active site were found to be highly deleterious to the activity of the enzyme, whereas most of the mutations further away from the active site did not change the activity of the enzyme significantly, but nevertheless they produced some changes in the pH–activity profile. Unfortunately, the changes in the pH–activity profiles did not correlate with the predictions from electrostatic potential calculations, prompting us to suggest that effects other than electrostatics were important for determining the pH–activity profile.

In this paper, we describe how these `other effects' are indeed important for determining the pH–activity profile. We designed mutations at four positions around the active site of a chimeric Bacillus α -amylase (Ba2). This enzyme was chosen as a model system because high-resolution structures are available for both the apo and holo forms of this enzyme ( Brzozowski et al. , 2000 ). We mutated the original neutral residues at the four positions to both neutral and charged residues and examined the effects on the pH–activity profile.

The results indicate that point mutations that are likely to change the dynamics of the active site can change the pH–activity profile. The effects on the pH–activity profile of a neutral → neutral mutation are so large that effects caused by neutral → charged mutations in this study are also likely to be dominated by the associated differences in active site dynamics.

Site-directed mutagenesis

The `MegaPrimer' method for site-directed mutagenesis ( Sarkar and Sommer, 1990 ) was used to construct a DNA fragment carrying the mutation. Mutagenic primers were designed so as to introduce or remove a unique site in the gene encoding the hybrid α-amylase (Ba2). The mutant DNA fragments were inserted into a Bacillus expression plasmid in the context of the Bacillus licheniformis α-amylase promoter, signal sequence and transcriptional terminator. The resulting mutant plasmids were transformed into Bacillus subtilis. Mutant colonies were identified by endonuclease digests of colony polymerase chain reaction (PCR) fragments and confirmed by DNA sequencing throughout the region covered by the mutagenic primer.

Fermentation

Fermentation was carried out at 37°C for 5 days in shake flasks containing a complex medium mainly consisting of potato flour, barley flour and sucrose soy meal ( Bang et al. , 1999 ; Beier et al. , 2000 ).

Protein purification

The supernatant of the fermentation mixture was isolated by flocculation and centrifugation. During ultrafiltration the buffer was changed to 20 mM sodium acetate, pH 5.5. The protein was subsequently applied to an SP-Sepharose Hi-Load column (Pharmacia). A dialysis step was used to change the buffer to 20 mM H 3 BO 3 + 5 mM KCl, pH 9.6. The final step of the purification procedure consisted of anion-exchange chromatography on a Mono-Q Hi-Load column (Pharmacia). All protein preparations were at least 95% pure as shown by SDS–PAGE.

Activity assays

The activity as a function of pH was measured over the range pH 4.0–10.5, using the Phadebas α-amylase test kit (Pharmacia). Measurements were carried out in duplicate at 30°C in 50 mM Britton–Robinson buffer (50 mM H 3 PO 4 + 50 mM CH 3 COOH + 50 mM H 3 BO 3 ) containing 0.1 mM CaCl 2 . The Phadebas α-amylase test kit is based on the release of blue colour from the substrate (blue-coloured starch) upon cleavage. Hydrolysis is stopped by adding 1/6 volume of 1 M NaOH to the reaction mixture. After removal of the unhydrolysed blue starch by filtration, the amount of hydrolysed substrate is proportional to the absorbance at 620 nm.

Since the substrate is insoluble (and added in large quantities), the activity measurements obtained by this method can be regarded as k cat for insoluble starch.

Stability assays

Stability was measured as the residual activity after incubation at 30°C for 15 min. Measurements were carried out at pH 4.5, 7.0 and 9.0 and were performed in duplicate. None of the mutants showed any detectable differences from the wild-type stability.

pK a calculations

p K a calculations were performed with the WHAT IF p K a calculation routines (Nielsen and Vriend, 2001). The routines apply the hydrogen-bond optimization procedure of Hooft et al. ( Hooft et al. , 1996 ) in order to model each of the protein protonation states accurately. DelPhi II ( Nicholls and Honig, 1991 ) was used to calculate electrostatic energies. The parameters for DelPhi II were set as described previously (Nielsen and Vriend, 2001), namely protein dielectric constant, 16 (for residues that participate in crystal contacts, have an alternative accessible rotamer or an average B -factor of >20) or 8 (all other residues); ionic strength, 0.160 M; solvent probe radius, 1.4 Å; and solvent dielectric constant, 80. The calculations were performed without water molecules, because the inclusion of water molecules was found to decrease the accuracy of the p K a calculations for a test set of nine proteins (Nielsen and Vriend, 2001).

A model of Ba2 in complex with malto-nonaose was constructed from the Ba2 α-amylases–acarbose X-ray structure ( Brzozowski et al. , 2000 ). The few changes that were necessary in order to convert the inhibitor to a substrate were made using Quanta. The CHARMm 22 parameter set was used as the source of charges and radii for the substrate in the p K a calculations.

Preparation of mutant structures

Mutant structures for use in the p K a calculations were designed using the WHAT IF position-specific rotamer libraries ( Chinea et al. , 1995 ). Mutant structures were inspected visually.

We constructed 12 point mutations to examine the determinants of the pH–activity profile for Ba2. The activity of all mutant amylases was within one order of magnitude of the activity of the wild-type at pH 7.0, with three variants having higher activity than the wild-type. The stability assays showed that the pH-dependent stability of the mutants was indistinguishable from that of the wild-type. The results are summarized in Table I .

Mutations were clustered at four positions, namely N190, F290, N326 and Q360 (Figure 3 ). The sites for the mutations were chosen reasonably close to the active site in order for the mutations to have a significant effect on the p K a values of the active site acids (Table II ). Also, we required that the wild-type residue at the position should be neutral. This was necessary in order to be able to construct neutral → neutral mutations. Visual inspection and WHAT IF's ( Vriend, 1990 ) structure analysis module furthermore indicated that mutations could be made at these positions without perturbing the local structure significantly. The apparent k cat profiles for the wild-type and the mutants were measured with the Phadebas assay. These k cat profiles will be referred to as `pH–activity profiles' in the following.

N190 is positioned in domain B in a turn that points into the edge of the substrate binding cleft (Figure 3 ). The residue does not hydrogen-bond to any other residue, except that its NH 2 group points to Tyr193 and most likely interacts with the π-electrons of the aromatic ring. The rotamer distributions for Asp and Lys at this position reveal that the Asp and Lys side chains will most likely point in the same direction as the Asn side chain, thereby causing almost no changes in the local structure. The unfavourable interaction of the Asp side chain with the π-electrons of Tyr193 is, however, likely to cause some rearrangements.

The pH–activity profiles of N190D, N190K and N190G show no significant changes compared with the wild-type pH–activity profile (Figure 4a ).

F290 is situated in the α-helix that lies between β-strand 6 and α-helix 6 of the TIM-barrel. This `in-between' α-helix is not a part of the classical TIM-barrel. Phe290 sits next to His289, which hydrogen bonds to Ser337. Ser337 is positioned in the loop that covers the active site and its mutation to glycine results in a protein with 2% of wild-type activity at pH 7.0, thus hinting at an important role played by the hydrogen bond between Ser337 and His289.

The pH–activity profiles for F290K, F290E and F290A are shown in Figure 4b . Both F290E and F290A are more active than the wild-type and both enzymes are much more active at basic pH than they would have been if they had had a pH–activity profile shaped as the wild-type protein. F290K is slightly less active than the wild-type, but its pH–activity profile has also been shifted slightly to more basic pH values.

Asn326 sits in the loop that covers the active site and is fairly close to Asp328 (Table I ). Based on the position-specific rotamer distributions and a bump analysis, it was judged that neither a lysine nor an arginine could be accommodated at this position. Furthermore, the rotamer distribution for leucine gave only two hits at this position, although visual inspection suggested that only minor adjustments in the local structure are needed to accommodate this mutation.

The pH–activity profile of N326A is almost identical with that of the wild-type (Figure 4c ), although the mutant has only ~60% of the wild-type activity at pH 7.0. N326D has relatively higher activity than the wild-type at basic pH, which is in agreement with the results reported previously by Takase (1993).

The pH–activity profile of N326L has lost the characteristic peak around pH 5.5 and has become almost completely flat. Furthermore, the basic limb of the pH–activity profile is shifted slightly towards more basic pH values.

The mutants Q360A and Q360K (Figure 4d ) have ~50% of the wild-type activity and their pH–activity profiles are very similar to that of the wild-type. Q360E, on the other hand, is more than twice as active as the wild-type at pH 6.0 and has a pH–activity profile which is much more bell-shaped than the wild-type profile.

p K a calculations were carried out as described in the Materials and methods section. The titration curves for the active site residues were mostly irregular and did not follow the classical Henderson–Hasselbalch shape (Figure 5 ). It was therefore not possible to determine a p K a value for any of the active site residues. Instead, we have given a qualitative description of the differences in the titration curves. The detailed titration curves for the active site residues in all mutant structures can be found at http://www.cmbi.kun.nl/gv/nielsen/amylase/pKa/ .

Wild-type pK a calculations

p K a calculations were performed for both the apo and holo forms of the enzyme. In the apo form, Glu261 and Asp328 form a tightly coupled system that titrates roughly as one group (hence the large oscillations in the titration curves for these two residues) with a p K a value of ~10, whereas Asp231 titrates with an apparent p K a of ~2. This corresponds fairly well with the established view of the α-amylase catalytic mechanism with Glu261 being the hydrogen donor and Asp231 being the nucleophile. In the calculations with the substrate, however, the titration curve for Asp231 becomes biphasic with apparent p K a values of ~4 and 7, whereas Glu261 remains negatively charged over the entire pH range. Asp328 is predicted to have a p K a value of ~8. The calculations for the holo form thus suggest that Asp328 is the hydrogen donor. It is, however, not clear how reliable the calculations for the holo form are as the coordinates for the holo form complex are derived from an enzyme–inhibitor complex rather than from an enzyme–substrate complex.

pK a calculations for the mutants

The p K a calculations for the mutants were carried out with the holo structure of the enzyme to give the p K a shifts for the active site acids when the substrate is bound. The perturbations in the titration curves for the active site residues were fairly small for all mutations of Asn190 and Phe290. This is in perfect agreement with the experimental results for the mutations at position 190, but does not correlate with the results for the mutations at position 290. Changes were observed for the mutations of Asn326 and Gln360. These changes are described in more detail below.

The calculations for N326A show an upward shift in the p K a value of His327 and a downward shift in the p K a value of Asp328. The calculations for N326L show a slight upward shift in the p K a value of Asp328 and an even smaller upward shift in the p K a value of Glu261. The N326D p K a calculations predict that the p K a values of Asp231, His327, Asp328 and Glu261 increase.

The p K a calculations for the mutants in position 360 predict that the p K a value of Asp231 becomes elevated upon all three mutations. In the case of Q360A and Q360K, the shift is very large, whereas a smaller shift is seen for Q360E. The p K a values of His327 and Asp328 are predicted to become slightly lower for Q360A. The calculations for Q360K predict that the p K a values for His327 and Asp328 increase.

Correlation of the calculated pK a shifts with the experimentally observed pH–activity profile shifts

The magnitude and direction of the pH–activity profile shifts for the Ba2 mutants are not reproduced by the p K a calculations. This is most clearly seen when comparing the calculations with the experimental results for F290A and N326A. F290A gives the largest shift in the pH–activity profile, but the p K a calculations produce insignificant changes in the active site p K a values for this mutation. In the case of N326A, the p K a calculations show a significant perturbation of the p K a values of Asp328 and His327, but the pH–activity profile for this mutant is almost identical with that of the wild-type.

The calculations for the mutations that introduce charges also do not correlate well with the experimental pH–activity profile shifts and this strongly suggests that long-range electrostatics are less important for the pH–activity profile of the α-amylases than previously thought.

The most pronounced effect on the pH–activity profile for a neutral → neutral mutation is seen for F290A. Phenylalanine and alanine have different sizes and the large effect is seen for this mutation is therefore not surprising. The imidazole ring of His289 packs against the aromatic ring of Phe290 in the wild-type structure and removing the aromatic ring of Phe 290 (by the F290A mutation) makes His289 more solvent accessible. The higher solvent accessibility changes the p K a of His289 slightly (by 0.1 unit in the p K a calculations) and in this way the F290A mutation changes the electrostatic field in the protein. This change in the electric field cannot, however, be responsible for the observed change in the pH–activity profile, since both the F290E and F290K mutations should change the electrostatic field even more than F290A. They should thus produce pH–activity profiles that are even more different from the wild-type profile than the pH–activity profile of F290A. This is, however, clearly not the case and the changes in the pH–activity profile caused by F290A therefore have to be ascribed to effects other than electrostatics as outlined below.

The His289–Ser337 hydrogen bond

His289 hydrogen bonds to Ser337 and the change in the p K a , and possibly in the dynamics of His289, is likely to affect the strength of this hydrogen-bond. Ser337 is situated in a loop that covers the active site and it is therefore likely that a change in the Ser337–His289 hydrogen bond could affect the dynamics of the active site. The hydrogen bond between His289 and Ser337 is known to be important, since the mutation of Ser337 to glycine produces an enzyme with a 50-fold reduction activity (J.E.Nielsen, G.Vriend and T.V.Borchert, unpublished results).

The reason for the discrepancy between the p K a calculations and the experiments for the mutants F290K and F290E now becomes clear, since these mutations are also likely to change the solvent accessibility and the dynamics of His289. The pH–activity profile shifts resulting from these two mutations are therefore the combined effects of the charge and the change in the mobility of His289 that are induced by the mutations. The p K a calculations do not attempt to model such effects and are therefore unable to reproduce the pH–activity profile shifts.

Takase constructed the N326D mutation ( Takase, 1993 ) in the Bacillus stearothermophilus α-amylase and reported the same downward shift in the pH–activity profile as we found. This shift is not readily explainable and is possibly due to changes in the mobility. The neutral → neutral mutations in this position (N326A and N326L) are both likely to influence the active site dynamics, but only N326L shows a significant change in the pH–activity profile.

The mutations at position 360 are remarkable in that Q360E has a large impact on the pH–activity profile, whereas the mutations Q360K and Q360A, which are expected to change the dynamics of the enzyme more than Q360E, have wild-type pH–activity profiles. This suggests that the effect of Q360E is purely electrostatic, although it is difficult to understand why Q360K does not have an equally large effect on the pH–activity profile.

Effect of introducing a point mutation on a tightly coupled system of titratable groups

In constructing the point mutations in BLA and BA2 we have silently assumed that the effect of inserting a titratable group near the active site could be predicted using the well-established rules that are summarized in Figure 2 . The p K a calculations for Q360K show that this is not always the case, as the p K a of Asp328 is calculated to increase when Gln360 is mutated to a lysine. This is clearly not what would be expected and shows that counterintuitive effects can indeed be achieved when a system of tightly coupled titratable groups is perturbed. The phenomenon is illustrated in Figure 6 , where the rules of Figure 2 are shown to break down for an Asp–Glu system much like the two catalytic acids in the α-amylases. It is therefore essential in many cases to use p K a calculations to predict the effect of charged point mutations on the p K a values in the active site.

We have shown that significant changes in the activity of pH–activity profiles of a Bacillus α -amylase can indeed be achieved using site-directed mutagenesis. The shifts in the pH–activity profiles for the mutants did not agree with the calculated changes in the active site electrostatics.

We speculate that changes in the dynamics of the active site residues are at least as important for the pH–activity profile as the changes in the active site electrostatics caused by the introduction of a charged residue. This is corroborated by the mutations F290A and N326L, which change the pH–activity profile without changing the net charge on the molecule. This implies that the p K a values of the active residues can be changed significantly by altering the dynamics of the active site and thus suggests an alternative approach to the engineering of pH–activity profiles.

A detailed explanation of the effects of the neutral → neutral mutations is difficult, as we do not know the motions that are important for the catalytic activity of the α-amylases. It is likely that very accurate and long molecular dynamics (MD) simulations could provide insights into this, but in view of the present day MD force fields and computer speeds, this is not a feasible solution. The p K a calculation method that we used does not model heavy atom mobility and the p K a shifts induced by mobility changes can therefore not be reproduced in the calculations. Furthermore, we may well have underestimated the value of the dielectric constant in the active site in our calculations. Employing a higher dielectric constant in the p K a calculations would reduce the magnitude of the calculated shifts in the titration curves, but it is unlikely that it would give a better qualitative agreement with the experimental data.

The p K a calculations also show that the α-amylase active site is a strongly connected system of titratable groups. The example with a strongly coupled Asp–Glu system given in Figure 6 shows that the effect of inserting a titratable group cannot always be predicted by using the simple scheme of Figure 2 . If the circumstances are right, one can observe the exact opposite of what was predicted. We speculate that the shifts in the `wrong' direction seen for several of the mutants in this and in the previous study ( Nielsen et al. , 1999a ) could be partly due to such effects. It seems more likely, though, that the change in the pH–activity profiles result primarily from the change in the active site dynamics.

The findings of this study stress the point that dynamics are an essential part of every enzyme and that rational engineering of enzyme activity is more likely to succeed if a detailed description of the enzyme mobility is available when designing the point mutations.

Specific activity of the Ba2 point mutations at pH 7.0 as measured by the Phadebas assay

Distance from mutated residues to the active site acids

Catalytic mechanism of the retaining glycosyl hydrolases. ( I ) Protonation of the glycosidic oxygen by the hydrogen donor (Glu261) and attack on the glucose C1 by the nucleophile (Asp231). Departure of the reducing end of the substrate. ( II ) Activation of a water molecule, cleavage of C1–Asp231 covalent bond. ( III ) Regeneration of the initial protonation states.

Environmental effects on the p K a of titratable residues. The effect of placing a titratable group in a negative, positive and hydrophobic environment is summarized.

The position of the point mutations in Ba2. Domain A, green; domain B, cyan; domain C, light grey. The calcium ions are shown as red spheres and the sodium ion is shown as an orange sphere. The nonaose substrate used in the p K a calculations is shown in yellow. The active site acids Asp231, Glu261 and Asp328 are shown in red.

pH–activity profiles for the Ba2 wt and the mutants. ( a ) wt, N190K, N190G and N190D; ( b ) wt, F290E, F290K and F290A; ( c ) wt, N326L, N326A and N326D; ( d ) wt, Q360K, Q360A and Q360E.

Calculated titration curves of the three active site acids Asp231, Glu261 and Asp328 in the apo-form and in the holo-form of the enzyme. Asp231, red; Glu261, magenta; Asp328, green.

An example of the effect of a charged point mutation on a hypothetical Glu–Asp system. It is seen that the simple predictive rules in Figure 2 break down when the system of titratable groups is strongly coupled. The electrostatic interaction energies between the titratable groups corresponds approximately to the calculated interaction energies between Glu261, Asp328 and Lys 234 in Ba2. ( a ) Schematic representation of the Glu–Asp system. The p K a of each residue in the absence of the other is 3.5 (intrinsic p K a in the Figure). The interaction energy between the two groups is 3.0 ln(10) kT (298.15 K). ( b ) Schematic representation of the Glu–Asp–Lys system. The interaction energy between the Asp and the Glu is the same as in (a). The interaction energy with the Lys is twice as large for the Asp as for the Glu. ( c ) The titration curves for the Asp–Glu system shows that the Asp and the Glu have identical titration curves. ( d ) The titration curves for the Asp–Glu–Lys system shows that the titration curve of the Asp is shifted to more acidic pH values (= lower p K a ), whereas the titration curve of the Glu is shifted to more basic pH values (= higher p K a ), thus defying the rules specified in Figure 2 .

To whom correspondence should be addressed. Present address: 4222 Urey Hall, Department of Chemistry and Biochemistry, University of California, San Diego, Mail Code 0365, La Jolla, CA 92093–0365, USA.

Email: [email protected]

The authors thank Vibeke Holbo for help with the construction and purification of mutant proteins, Jytte Piil for providing purified wild-type protein and Rebecca Wade, Barry Honig and Lawrence McIntosh for stimulating discussions.

Bang,M.L., Villadsen,I. and Sandal,T. ( 1999 ) Cloning and characterization of an endo-beta-1,3(4)glucanase and an aspartic protease from Phaffia rhodozyma CBS 6938. Appl. Microbiol Biotechnol. , 51 , 215 –222.

Beier,L., Svendsen,A., Andersen,C., Frandsen,T.P., Borchert,T.V. and Cherry,J.R. ( 2000 ) Conversion of the maltogenic alpha-amylase Novamyl into a CGTase. Protein Eng. , 13 , 509 –513.

Brzozowski,A.M., Lawson,D.M., Turkenburg,J.P., Bisgaard-Frantzen,H., Svendsen,A., Borchert,T.V., Dauter,Z., Wilson,K.S. and Davies,G.J. ( 2000 ) Structural analysis of a chimeric bacterial α-amylase. High resolution analysis of native and ligand complexes. Biochemistry , 39 , 9099 –9107.

Chinea,G., Padron,G., Hooft,R.W.W., Sander,C. and Vriend,G. ( 1995 ) The use of position-specific rotamers in model building by homology. Proteins , 23 , 415 –421.

Davies,G.J. and Henrissat,H. ( 1995 ) Structures and mechanisms of glycosyl hydrolases. Structure , 3 , 853 –859.

Davies,G.J., Wilson,K.S. and Henrissat,B. ( 1997 ) Nomenclature for sugar-binding subsites in glycosyl hydrolases. Biochem. J. , 321 , 557 –559.

Fang,T.Y. and Ford,C. ( 1998 ) Protein engineering of Aspergillus awamori glucoamylase to increase its pH optimum. Protein Eng. , 11 , 383 –388.

Guzman-Maldano,H. and Paredes-Lopez,O. ( 1995 ) Amylolytic enzymes and products derived from starch; a review. Crit. Rev. Food. Nutr. , 36 , 373 –403.

Hooft,R.W., Sander,C. and Vriend,G. ( 1996 ). Positioning hydrogen atoms by optimising hydrogen-bond networks in protein structures. Proteins , 26 , 363 –376.

Keating,L., Kelly,C. and Fogarty,W. ( 1998 ). Mechanism of action and the substrate-dependent pH maximum shift of the α-amylase of Bacillus coagulans . Carbohydr. Res. , 309 , 311 –318.

Klein,C., Hollender,J., Bender,H. and Schulz,G.E. ( 1992 ) Catalytic center of cyclodextrin glycosyltransferase derived from X-ray structure analysis combined with site-directed mutagenesis. Biochemistry , 31 , 8740 –8746.

Knegtel,R.M, Strokopytov,B., Penninga,D., Faber,O.G., Rozeboom,H.J., Kalk,K.H., Dijkhuizen,L. and Dijkstra,B.W. ( 1995 ) Crystallographic studies of the interaction of cyclodextrin glycosyltransferase from Bacillus circulans strain 251 with natural substrates and products. J. Biol. Chem. , 270 , 29256 –29264.

Kyte,J. ( 1995 ) Mechanism in Protein Chemistry . Garland Publishing, New York, pp. 258–283.

McCarter,J.D. and Withers,S.G. ( 1994 ) Mechanisms of enzymatic glycoside hydrolysis. Curr. Opin. Struct. Biol. , 4 , 885 –892.

McCarter,J.D. and Withers,S.G. ( 1996 ) Unequivocal identification of Asp-214 as the catalytic nucleophile of Saccharomyces cerevisiae α-glucosidase using 5-fluoro glycosyl fluorides. J. Biol. Chem. , 271 , 6889 –6894.

McIntosh,L.P., Hand,G., Johnson,P.E., Joshi,M.D., Korner,M., Plesniak,L.A., Ziser,L., Wakarchuk,W.W. and Withers,S.G. ( 1996 ) The p K a of the general acid/base carboxyl group of a glycosidase cycles during catalysis: a 13C-NMR study of Bacillus circulans xylanase. Biochemistry , 35 , 9958 –9966.

Nicholls,A. and Honig,B. ( 1991 ) A rapid finite difference algorithm, utilizing successive over-relaxation to solve the Poisson–Boltzmann equation. J. Comput. Chem. , 12 , 435 –445.

Nielsen,J.E. and Borchert,T.V. ( 2000 ) Protein engineering of bacterial α-amylases. Biochim. Biophys. Acta , 1543 , 253 –274.

Nielsen,J.E. and Vriend,G. ( 2001 ) Optimising the hydrogen-bond network in Poisson–Boltzmann equation based p K a calculations. Proteins , 43 , 403 –412.

Nielsen,J.E., Beier,L., Otzen,D., Borchert,T.V., Frantzen,H.B., Andersen,K.V. and Svendsen,A. ( 1999 ) Electrostatics in the active site of an α-amylase. Eur. J. Biochem. , 264 , 816 –824.

Qian,M., Haser,R., Buisson,G., Duee,E. and Payan,F. ( 1994 ) The active centre of a mammalian α-amylase. Structure of the complex of a pancreatic α-amylase with a carbohydrate inhibitor refined to 2.2-Å resolution. Biochemistry , 33 , 6284 –6294.

Sarkar,G. and Sommer,S.S. ( 1990 ). The `megaprimer' method of site-directed mutagenesis. Biotechniques , 8 , 404 –407.

Shaw,A., Bott,R. and Day,A.G. ( 1999 ) Protein engineering of α-amylase for low pH performance. Curr. Opin. Biotechnol. , 10 , 349 –352.

Sinnot,M.L. ( 1990 ) Catalytic mechanisms of enzymic glycosyl transfer. Chem. Rev. , 90 , 1171 –1202.

Strokopytov,B., Penninga,D., Rozeboom,H.J., Kalk,K.H., Dijkhuizen,L. and Dijkstra,B.W. ( 1995 ) X-ray structure of cyclodextrin glycosyltransferase complexed with acarbose. Implications for the catalytic mechanism of glycosidases. Biochemistry , 34 , 2234 –2240.

Takase,K. ( 1993 ) Effect of mutation of an amino acid residue near the catalytic site on the activity of Bacillus stearothermophilus alpha-amylase. Eur. J. Biochem. , 211 , 899 –902.

Uitdehaag,J.C., Mosi,R., Kalk,K.H., van der Veen,B.A., Dijkhuizen,L., Withers,S.G. and Dijkstra,B.W. ( 1999 ) X-ray structures along the reaction pathway of cyclodextrin glycosyltransferase elucidate catalysis in the α-amylase family. Nature Struct. Biol. , 6 , 432 –436.

Vriend,G. ( 1990 ) WHAT IF: a molecular modelling and drug design program. J. Mol. Graphics , 8 , 52 –56.

Wind,R.D., Uitdehaag,J.C., Buitelaar,R.M., Dijkstra,B.W. and Dijkhuizen,L. ( 1998 ) Engineering of cyclodextrin product specificity and pH optima of the thermostable cyclodextrin glycosyltransferase from Thermoanaerobacterium thermosulfurigenes EM1. J. Biol. Chem. , 273 , 5771 –5779.

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