concentration of amylase on starch experiment

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Amylase on Starch Lab

INTRODUCTION:

In this experiment you will observe the action of the enzyme amylase on starch. Amylase changes starch into a simpler form: the sugar maltose , which is soluble in water. Amylase is present in our saliva, and begins to act on the starch in our food while still in the mouth. Exposure to heat or extreme pH (acid or base) will denature proteins. Enzymes, including amylase, are proteins. If denatured, an enzyme can no longer act as a catalyst for the reaction. Benedict’s solution is a test reagent that reacts positively with simple reducing sugars like maltose, but will not react with starch. A positive test is observed as the formation of a brownish-red cuprous oxide precipitate. A weaker positive test will be yellow to orange.

Add 1g of cornstarch to a beaker containing 100ml of cold distilled water. While stirring frequently, heat the mixture just until it begins to boil. Allow to cool.

1. Fill the 250-mL beaker about 3/4 full of water and place on the hot plate for a boiling water bath. Keep the water JUST AT BOILING .

2. Mark 3 test tubes A , B and C . “Spit” between 1 and 2 mL of saliva into each test tube.

3. Into tube A , add 2 mL of vinegar. Into tubes B and C , add 2 mL of distilled water. Thump the tubes to mix.

4. Place tube B into the boiling water bath for 5 minutes . After the five minutes, remove from the bath, and place back into the test tube rack.

5. Add 5 mL of the starch solution to each tube and thump to mix. Allow the tubes to sit for 10 minutes , occasionally thumping the tubes to mix.

6. Add 5 mL of Benedict’s solution to each tube and thump to mix. Place the tubes in the hot water bath. The reaction takes several minutes to begin.

OBSERVATIONS:

Tube A : Starch + saliva treated with vinegar (acid)

• Was the test positive or negative? _______________________

What does this indicate?__________________________________________________

____________________________________________________________________

Tube B : Starch + saliva and water, treated in a boiling water bath

Tube C : Starch + saliva

1. What is the function of an enzyme?

2. Where does a substrate attach to an enzyme?

3. If an enzyme is present in a reaction, less ________________ _________________ will be needed to get the reaction started.

4. What is a common suffix found at the end of most biological enzymes?

5. Most enzymes are macromolecules called ________________.

6. Define denaturation of proteins.

7. Name 3 things that can denature or unfold an enzyme.

8. In this lab, what weak acid denatured the protein?

9. What was the purpose of placing one test tube in a hot water bath?

10. What happens to enzymes in your body whenever you run fever?

EXPERIMENT NO. 5

Starch hydrolysis by amylase, table of contents, introduction, list of reagents and instruments, a. equipment, b. reagents, discussions, tabular forms.

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 amylase on a starchy foodstuff, class practical or demonstration.

Place rice in a Visking tubing bag to model food in the gut . Investigate amylase action by adding water, amylase, or boiled amylase to the rice. Leave for 10-15 minutes in a water bath at around 37 °C then test for the presence of a reducing sugar in the water surrounding the Visking tubing bag.

Lesson organisation

This experiment could be done as a demonstration or in groups. Each group needs to set up three Visking tubing bags, so a group of 3 students is ideal.

Apparatus and Chemicals

For each group of students:.

3 x 15 cm lengths of Visking tubing

Syringe barrels, sawn off, 3

Boiling tube, 3

Test tubes, 6

Test tube racks to accommodate 6 test tubes and 3 boiling tubes per group

Teat pipettes, 6

White dimple (spotting) tile

Beaker, 250 cm 3

Kettle for boiling water for Benedict’s test

Eye protection for each student

For the class – set up by technician/ teacher:

Length of Visking tubing, knotted at one end, 15 cm, 3 per group ( Note 1 )

Syringe barrel, sawn off, 3 per group ( Note 2 )

Elastic bands, 3 per group

Electric water baths set at 35-40 °C, with thermometer to show temperature accurately

Cooked rice

Iodine solution ( Note 3 )

Benedict’s reagent ( Note 4 )

Amylase solution, 5 cm 3 per group ( Notes 5 and 6 )

Boiled amylase, 5 cm 3 per group

Clinistix (as an alternative to Benedict’s reagent) ( Note 7 )

Health & Safety and Technical notes

Students should wear eye protection when handling chemicals. Electrical apparatus should be maintained and checked according to your employer’s instructions. Ensure students know how to deal with breakages of glass or thermometers

Read our standard health & safety guidance

1 Soak the Visking tubing in warm water beforehand so it is ready to use.

2 The end of an old syringe makes a convenient support for the Visking tubing, and makes it easier to take samples of the contents with a teat pipette.

3 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.

4 Benedict’s (qualitative) reagent. (See CLEAPSS Recipe card) No hazard warning is required on the bottle, as the concentrations of each of the constituents are low. Take care making up the reagent; sodium carbonate is an irritant to the eyes and copper(II) sulfate(VI) is harmful if swallowed. See CLEAPSS Hazcards.

5 Amylase solution: Check your amylase supply as many contain starch or reducing sugars, which would interfere with the results of this test. Alpha amylase is bacterial amylase with high activity, and does not give a positive reducing sugar test or starch test. You can use lower concentrations of this enzyme. Some bacterial amylases may survive boiling!

Using saliva: the CLEAPSS Laboratory Handbook provides guidance on precautions, including hygiene precautions, for safe use of saliva as a source of amylase. This has the advantage of being cheaper and technicians do not need 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. Students must be responsible for rinsing their own equipment. All contaminated glassware is placed in a bowl or bucket of sodium chlorate(I) for technicians wash up.

6 Working with enzymes: It is wise to test, well in advance, the activity of stored enzymes at their usual working concentrations 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.

7 Clinistix are quick and easy to use. Each stick can be cut into two or three pieces.

Ethical issues

There are no ethical issues associated with this protocol.

Preparation

a Prepare boiled rice, enzyme solution, boiled enzyme solution, iodine solution, and Benedict’s reagent.

b Set up a water bath at 37 °C.

c Soak Visking tubing, cut 15 cm lengths (3 per group) and set up model guts with syringe barrels, or leave for students to assemble.

Investigation

d Label 3 boiling tubes 1, 2, 3.

e Label 3 test tubes 1, 2, 3.

f Set up 3 model guts: take a wet piece of Visking tubing, tie a knot in one end, place the sawn off syringe barrel in the other end and secure with an elastic band. These may have been set up for you (see diagram).

g Use the spatula to add rice to each of the model guts until they are half full.

h Rinse the outside of each piece of Visking tubing under a running tap.

i Place the rice-filled model gut in a labelled boiling tube. Add warm water to boiling tube outside the Visking tubing until it reaches about 2 cm higher than the level of the liquid inside the Visking tubing (see diagram).

j Immediately withdraw one drop of the water you have added and test it with iodine on a dimple tile.

k Add 5 cm 3 of water to model gut 1.

l Add 5 cm 3 of amylase to model gut 2.

m Add 5 cm 3 of boiled amylase to model gut 3.

n Place all the boiling tubes containing the model guts in the water bath at approximately 37 °C.

o Leave for at least 15 minutes.

p While you are waiting:

• Place a grain of rice in a well on the white tile and add a drop of iodine.
• Put some rice in a test tube. Add 2 cm 3 of water and 2 cm 3 of Benedict’s reagent, and place into a large beaker of boiling water. Check the colour after 2-3 minutes.
• Record your results in a suitable table.

q After 15 minutes, use a teat pipette to remove 2-3 cm 3 of the water surrounding the model gut in boiling tube 1.

r Place one drop of this water in a well on the white tile and add a drop of iodine. Record the result.

s Place the rest (around 2 cm 3 ) of the water from boiling tube 1 into test tube 1. Add an equal volume of Benedict’s reagent and place test tube 1 into a large beaker of boiling water. Check the colour after 2-3 minutes. Record the result.

t Repeat steps q , r , s with water from boiling tubes 2 and 3. Record the results.

Teaching notes

The sawn off syringe barrel acts as a model mouth to the gut. It is a good idea to use cooked rice, as this is real food and can be seen in the (model) gut.

Many students will need help to understand this activity. When interpreting the results, students have to think in terms of two types of model: the model gut with Visking tubing representing the selectively permeable membranes lining the gut wall, and a simplified chemical model of large and small molecules. A further complication is that the movement of chemicals is unseen and only inferred from the results of chemical tests. An additional model could be used, with chicken wire or mesh, fruit or satsuma bags to represent the membrane, and poppet beads in chains to represent starch and singly to represent glucose.

Health and safety checked, September 2008

Related experiments

Evaluating Visking tubing as a model for a gut

Investigating the effect of pH on amylase activity

Digestion of starch by microbes

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Amylase Starch Experiments

Enzyme Model Science Projects

Amylase is an enzyme responsible for converting starches into the sugar maltose, which is a disaccharide. This enzyme, present in saliva, is a key component in germinating plants. The starches contained within the seed are converted to sugars, providing energy to the plant before photosynthesis begins. Experiments with amylase demonstrate how the enzyme reacts with starches and variables, which affect the rate of the reaction.

Chewing Bread

Bread is full of carbohydrates. Starches are considered a type of complex carbohydrate, which begins to be broken down into maltose as soon as it's in our mouths. Give each student a slice of bread that has been cut in two. The students chew one half of the bread for three minutes and write down their observations as to the changes in how the bread tastes. The other half of the bread is chewed for 10 seconds, then placed in a safe container for 10 minutes. After 10 minutes are up, the students chew the bread again. In both cases, the bread should begin to get sweeter as the amylase begins to convert the carbohydrates into maltose, which tastes sweet.

Give the students three corn seeds -- one dry, another that has been boiled, and one that has been soaked in water. The students cut the seeds in half and place the seeds on an agar petri dish that has a starch solution. The students then incubate the seeds for 30 minutes. After removal, they add an iodine solution over the plates. Starches remaining on the plate react with the iodine, creating purple areas. Students observe the differences between the seeds to determine which type of seed had more active amounts of amylase present.

As with all enzymes, amylase has a preferred pH level in which it operates. This can be determined by creating different pH levels and amylase reactions that measure the speed of the reaction. Place iodine solution drops in a test tube. In test tubes mix amylase, starch and a buffer solution with different pH levels. After mixing the solution, remove a small amount using a pipette and add it to the iodine. The iodine must turn orange when the reaction is complete. The students test the solution every 10 seconds until they arrive at the correct color. The experiment is repeated at each pH level. The pH level that turned orange the fastest is the preferred pH of amylase.

Temperature

Amylase reactions happen more rapidly at certain temperatures. Place iodine solution in a tray. Mix the amylase, starch and buffer, use the same pH this time, and test how long it takes to turn orange. Raise the temperature of the solution by 10 degrees for the next solution and retest the time it takes for the reaction to test. Students should determine the optimum temperature for the amylase reaction through multiple trials.

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Action of Salivary Amylase on Starch

To study the action of salivary amylase on starch solution.

The interaction between salivary amylase and starch constitutes a fundamental aspect of our digestive process, providing a glimpse into the complex biochemical mechanisms that enable our bodies to extract energy from the food we consume. Salivary amylase, an enzyme secreted by salivary glands, initiates the breakdown of complex starch molecules present in our diet. As food enters the mouth, the enzyme catalyses the hydrolysis of starch into simpler sugars like maltose. This enzymatic action marks the first step in carbohydrate digestion.

Experiment Procedure

To demonstrate the experiment on the action of salivary amylase on starch, we need to follow the given procedure:

• First of all, rinse your mouth with fresh water and collect saliva using a spatula/spoon.
• Then, filter saliva through a cotton swab.
• Now, take 1 mL of filtered saliva in a test tube and add 10 mL of distilled water to the test tube. Label it as “saliva solution.”
• Next, take 2 mL of 1% starch solution in 2 labelled test tubes (A and B).
• Add 1 mL diluted saliva to test tube B and shake well.
• We will not add anything to test tube A and keep it in control.
• After 5 minutes, take 5 drops from test tube A on a tile or a glass slide.
• Add 2 drops of 1% iodine solution in it, mix and observe colour.
• Place 5 drops from test tube B away from A’s mixture.
• Add 2 drops of 1% iodine solution to B’s drops, mix and observe.
• Repeat the iodine test after 5, 10, 15, and 20 minutes.

In conclusion, the experiment about “Action of Salivary Amylase on Starch” shows how enzymes in our bodies help break down food. By collecting saliva, diluting it, and mixing it with starch, it demonstrates how starch changes into simpler sugars. This change is important for getting energy from our food. Iodine solution is used to see this change, which makes the colour of the mixture different. These findings remind us how our bodies work with chemicals to stay alive. Understanding how salivary amylase acts on starch helps us see how our bodies make use of the food we eat.

FAQs on the Action of Salivary Amylase on Starch

Q.1 what is salivary amylase and its role in digestion.

Ans. Salivary amylase is an enzyme produced by the salivary glands, primarily in the mouth. Its main role is to initiate the digestion of complex carbohydrates, specifically starches, into simpler sugars.

Q.2 How does salivary amylase work on starch?

Ans. Salivary amylase breaks down the starch molecules into smaller fragments by catalysing the hydrolysis of the glycosidic bonds that link the glucose units in the starch molecule. This results in the production of maltose and other shorter carbohydrate chains.

Q.3 What factors can affect the activity of salivary amylase on starch?

Ans. The activity of salivary amylase can be affected by factors like pH, temperature, and the presence of inhibitors. An optimal pH level is necessary for its activity, and extreme temperatures or certain inhibitors might denature or inhibit the enzyme’s function.

Q.4 Why do we rinse the mouth with fresh water at the beginning of the experiment?

Ans. Rinsing the mouth with fresh water helps remove any residual food particles or substances that might interfere with the experiment. It ensures that only the collected saliva is being used in the experiment.

Q.5 What happens to the pH during salivary amylase action on starch?

Ans. Salivary amylase works optimally in a slightly acidic to neutral pH range, typically around pH 6.7. It starts the starch digestion process in the mouth, where the slightly acidic environment due to the presence of acids from foods and beverages helps activate the enzyme. 

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The Effects of Temperature, pH and Enzyme Concentration on Amylase

Written by Sarah

Introduction

Enzymes are proteins that are critical to catalyzing reactions (Brooker, Widmaier, Graham & Stiling, 2011). Like most proteins, they are synthesized by the ribosomes in the cell. They react with a specific substrate in order to increase the rate of a chemical reaction within the cell. Without enzymes, reactions would be significantly slower and we would not be able to do the most basic functions, such as breathing or digesting food.

Amylase is an type of enzyme. Amylase has an active site organized in subsites, each of which accommodates a glucose residue (Talamond, Noirot & de Kochko, 2005). It breaks down starch to glucose, giving food that sweet taste. An example of amylase in the natural world is in bananas. When they are green, the amylase has yet to break down the starch, but by the time they’ve turned brown, the reaction has been completed. This is why brown bananas taste sweeter than their green counterpart.

Within this experiment, the objective was to test how temperature, pH level and enzyme concentration changed the effectiveness of amylase.

First, an indicator experiment was performed. In two test tubes, a 1 mL (millimeter) sample of starch solution was pipetted. The same was done with 1 mL of maltose solution into another two tubes. Into one of the test tubes of maltose and one of starch, 5 drops of I 2 KI were added. In the other two test tubes of maltose and starch, 5 drops of Benedict’s reagent were added. The test tubes with Benedict’s reagent were placed in a hot sand bath. All four tubes were then watched for color change, indicating a reaction.

The activity of amylase was then observed through three reaction mixtures. The first mixture was 1 mL of starch solution and 1 µL (microliter) of amylase solution. The second was 1 mL of starch and 50 µL of water. The third and final was 50 µL of amylase and 1 mL of water. Immediately a drop of each mixture was transferred to a separate well on a spot plate and a drop of I 2 KI was added. This was then repeated every minute. The color was observed with the passage of time to conclude whether amylase activity was present or not. After 8 minutes, 5 drops of Benedict’s reagent was added to all three tests tubes which were then placed in the hot sand bath. The tubes were observed for color change, indicating the presence of maltose.

The effects of temperature were observed through three water baths set to 4°C (Celsius), 23°C and 37°C with a solution of pH 7 starch solution resting in all three. Fifty µL of amylase solution was pipetted into a test tube which was placed in the water bath for 1 minute. Then, 1 mL of the temperature equilibrated starch was added. Immediately a drop of the reaction was transferred to the spot plate and 1 drop of I 2 KI was added. This step was repeated every minute to test the presence of starch. Complete this process for all three temperatures. At the end, 5 drops of Benedict’s reagent was added and the tubes were placed in the hot sand bath.

Next, pH levels were tested. Four test tubes were filled with 1 mL of pH 4, 5, 6 and 7 starch solutions respectively. Then 50 µL of amylase were added to each tube and immediately, one drop of each mixture was transferred to the spot plate and a drop of I 2 KI was added. This was repeated at one minute intervals. After 8 minutes, add 5 drops of Benedict’s reagent was added to the test tubes and they were placed in the hot sand bath.

Finally, the effects of enzyme concentration were tested. One mL of pH 7 starch solution was pipetted into three test tubes. To one test tube, 50 µL of 5% enzyme was added, 50 µL of 10% enzyme to another and 50 µL of 20% enzyme to the final tube. One drop of each mixture was immediately transferred to the spot plate and a drop of I 2 KI was added. This was repeated every minute. After 8 minutes, 5 drops of Benedict’s reagent was added to the test tubes which were then placed within the hot sand bath.

Within the indicators experiment, the first test tube changed from clear to dark purple. The second tube changed from blue to yellow. The third tube didn’t change and neither did the fourth.

For the activity of the amylase experiment, tube 1 indicated starch within the first test but didn’t after. Tube 2 indicated starch for all 8 minutes. Tube 3 never indicated starch. Only tube 1 changed any color after exposure to Benedict’s reagent and heat, proving maltose was present.

Temperature seemed to have a positive correlations with the speed of the reaction. The 37°C completed the reaction the quickest. This is shown in Figure 1. Maltose was shown to be present in all three test tubes.

Figure 1. The Effects of Temperature on Enzyme Activity

The correlation of pH doesn’t seem as clear. Test tubes 2 and 3, which contained pH 5 and 6 starch, completed the reaction at the same time. Figure 2 shows the reaction rates of the different pH levels. Maltose was present in all test tubes.

Figure 2. The Effects of pH Level on Enzyme Activity

As the concentration of amylase increased, the reaction time decreased. Figure 3 shows the direct correlation. All three tubes had maltose present.

Figure 3. The Effects of Enzyme Concentration on Activity

In the indicator experiment, it was necessary to test what each indicator marked. This allowed for I 2 KI to be used to indicate starch and Benedict’s reagent to be used to indicate maltose. The controls/amylase activity experiment showed that it is necessary for both starch and amylase to be mixed in order for the reaction to occur. As expected, as the temperature increased, so did the speed of the reaction. This is expected because this reaction often occurs in the human body where the temperature is normally 37°C. The results of the pH experiment did not clearly show what the best level for the enzyme was. The concentration experiment was much more clear. The 20% concentration reacted the fastest because there was more enzymes to react with the substrate and create maltose.

Literature Cited

Brooker, Robert J., Eric P. Widmaier, Linda E. Graham, and Peter D. Stiling. Biology . 2nd ed.

New York: McGraw Hill, 2011. Print.

Talamond, Pascale, Michel Noirot, and Alexandre De Kochko. “The Mechanism of Action of

α-amylase from Lactobacillus Fermentum on Maltooligosaccharides.” Journal of

Chromatography B (2005): 42-47. Science Direct . Web.

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Biochemical Characterization of the Amylase Activity from the New Haloarchaeal Strain Haloarcula sp. HS Isolated in the Odiel Marshlands

Patricia gómez-villegas.

1 Laboratory of Biochemistry, Department of Chemistry, Marine International Campus of Excellence (CEIMAR), University of Huelva, Avda. de las Fuerzas Armadas s/n, 21071 Huelva, Spain; moc.liamg@livmogtap (P.G.-V.); se.uhu@aragiv (J.V.)

Javier Vigara

Luis romero.

2 Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and Universidad de Sevilla, Avenida Américo Vespucio 49, 41092 Seville, Spain; se.cisc.fvbi@oremorl (L.R.); se.cisc.fvbi@rotog (C.G.)

Cecilia Gotor

Sara raposo.

3 CIMA—Centre for Marine and Environmental Research, FCT, Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal; tp.glau@osopars (S.R.); tp.glau@seugirdorgb (B.G.)

Brígida Gonçalves

Rosa léon, associated data.

All DNA and protein sequences of the studied enzymes are included as Supplementary Materials ; other information is available upon request.

Simple Summary

Amylases are a group of enzymes that degrade starch into simple sugars. These proteins are produced by a wide variety of organisms and are supposed to be one of the most valuable industrial enzymes. However, the extreme conditions required for many industrial operations limit the applicability of most amylases found in nature. In this context, halophilic archaea entail an excellent source of novel proteins that tolerate harsh conditions, as they live in environments with high salt concentration and temperature. In this work, a screening of haloarchaea, isolated from Odiel salterns in the southwest of Spain, was carried out to select a new strain with a high amylase activity. This microorganism was identified as Haloarcula sp. HS and showed amylase activities in both, the cellular and the extracellular extracts. Both amylase activities were poly-extremotolerant, as their optimal yields were achieved at 60 °C and 25% NaCl. Additionally, the study of the protein sequences from Haloarcula sp. HS allowed the identification of three different amylases, which conserved the typical structure of the alpha-amylase family. Finally, the applicability of the extracellular amylase to treat bakery wastes under high salinity conditions was demonstrated.

Alpha-amylases are a large family of α,1-4-endo-glycosyl hydrolases distributed in all kingdoms of life. The need for poly-extremotolerant amylases encouraged their search in extreme environments, where archaea become ideal candidates to provide new enzymes that are able to work in the harsh conditions demanded in many industrial applications. In this study, a collection of haloarchaea isolated from Odiel saltern ponds in the southwest of Spain was screened for their amylase activity. The strain that exhibited the highest activity was selected and identified as Haloarcula sp. HS. We demonstrated the existence in both, cellular and extracellular extracts of the new strain, of functional α-amylase activities, which showed to be moderately thermotolerant (optimum around 60 °C), extremely halotolerant (optimum over 25% NaCl), and calcium-dependent. The tryptic digestion followed by HPLC-MS/MS analysis of the partially purified cellular and extracellular extracts allowed to identify the sequence of three alpha-amylases, which despite sharing a low sequence identity, exhibited high three-dimensional structure homology, conserving the typical domains and most of the key consensus residues of α-amylases. Moreover, we proved the potential of the extracellular α-amylase from Haloarcula sp. HS to treat bakery wastes under high salinity conditions.

1. Introduction

Haloarchaea are the main representatives of extreme halophiles, which can thrive in media with salt concentrations ranging from 20 to 30% [ 1 , 2 ]. These singular microorganisms are characterized by the accumulation of large amounts of KCl in the cytoplasm, to maintain the osmotic balance with the medium, in contrast to moderate or facultative halophiles, which usually store compatible solutes for the same purpose [ 3 ]. Therefore, intra- and extracellular proteins from haloarchaea are specially adapted to work properly at high salt concentrations. These proteins possess exceptional features that make them distinguishable from non-halophilic proteins; they have a unique amino acid composition with acidic surfaces and low overall hydrophobicity, to prevent aggregation and, at the same time, retain flexibility in such high salinity [ 4 ]. Most haloarchaeal enzymes are considered poly-extremotolerant, as they work appropriately under more than one extreme condition, usually elevated temperatures, in addition to high salinity. For this reason, enzymes from these halophilic microorganisms can be of interest in many harsh industrial and biotechnological processes [ 5 , 6 ].

Amylases are a diverse group of hydrolase or transferase enzymes that degrade large alpha-linked polysaccharides, such as starch and related oligosaccharides, and are one of the most required enzymes in industrial operations. They stand for about 30% of the world enzyme market, and this value is expected to grow in the following years, due to the global increase in the demand for bakery and sugar-derived products, biofuels, detergents, breweries, animal feeds, pharmaceuticals, paper, and textiles [ 7 , 8 ]. At present, the best market for amylases is in the production of maltose and glucose syrups from corn, which are used as sweeteners for soft drinks [ 9 , 10 ]. They are also required for the enhancement of dough for baking, for clarification of fruit juices and beers, and in the pretreatment of animal feed to improve the digestibility of fiber [ 11 ]. In addition, amylases are widely used in textile and paper industries to remove the starch employed for the desizing process and the coating treatment, respectively. Moreover, several hydrolytic enzymes, including amylases, are usually added to detergents because they permit the use of milder conditions in laundry and automatic dishwashing, making them eco-friendlier. Other interesting fields of application of amylases are biomedicine and pharmacy, to treat digestive disorders or as reporter genes in molecular biology [ 12 ]. Furthermore, amylases are widely used for the conversion of starch-rich agronomic and food wastes into fermentable sugars, which are required as feedstocks for the production of fuels and chemicals with high demand and market value [ 13 ].

Amylases are ubiquitous ancient enzymes found in plants, animals, and microorganisms. Among them, bacteria of the genus Bacillus or fungi belonging to the Aspergillus genus are the most preferred source of amylases for large-scale production [ 14 ]. Hydrolytic amylases can be classified into two broad categories—endoamylases, which hydrolyze the interior of the starch molecule; and exo-amylases, which successively degrade starch from the non-reducing ends [ 9 ]. Most endoamylases belong to the α-amylase family (EC 3.2.1.1) and cleave internal α,1-4 glycosidic bonds between glucose units, producing oligosaccharides with varying lengths and α-limit dextrins. Additionally, α-amylases are typically divided into two groups, according to the hydrolysis products and the degree of starch hydrolysis; saccharifying α-amylases that produce free sugars, and liquefying α-amylases that break down the starch polymer without producing free sugars [ 5 ]. The mechanism of action and the catalytic properties of these amylases are well-known and can be correlated with their structural characteristics, as was detailed in several reviews [ 7 , 15 , 16 ].

The ability of haloarchaea to produce and excrete hydrolytic enzymes, including amylases to degrade extracellular polysaccharides, as many other microorganisms do, was previously described [ 6 ]. However, the application of archaeal amylases, which could be beneficial to many industrial operations that require extreme conditions, remains scarcely studied when compared to those from other microorganisms. Intracellular or cell-associated amylases from haloarchaea are particularly understudied, although they are an important haloarchaeal trait and can represent an interesting source of halotolerant enzymes.

The saltern ponds of the Odiel Marshlands are an interesting saline ecosystem, which harbors a rich diversity of prokaryotic and eukaryotic microorganisms. Our previous studies showed that at very high salinity (33%), the most abundant archaea species belong to the genera Halorubrum and Haloquadratum [ 17 ]. Metagenomic microbial profiling by high-throughput 16S rRNA sequencing revealed the existence of various strains that belonged to the Haloarcula genus. Although the abundance of these representatives was quite low, with less than 0.2% of the total sequenced reads, our data suggest that some of the haloarchaea of this group are able to produce bioactive compounds [ 18 ] and excrete hydrolytic haloenzymes, including proteases, amylases, or lipases [ 17 ].

In this study, a collection of haloarchaea isolated from the saltern ponds of the Odiel Marshlands was screened for their amylolytic activity, and the one that exhibited the highest activity was selected and identified on the basis of its 16S rRNA coding gene. The extracellular and cellular starch-degrading activities of the selected archaea were characterized, revealing different optimal parameters and modes of action. To get a further insight into the identity of these starch-degrading enzymes, the proteome composition of the partially purified cell-free supernatant and the cellular extracts was analyzed by tryptic digestion, followed by nano-liquid chromatography coupled to an electrospray ionization tandem mass spectrometry system. This study allowed the identification of three amylase sequences (two were exclusively cell-associated and one was also found in the extracellular medium) with high homology to amylases of other haloarchaeal species, and the typical alpha-amylase conserved regions. Furthermore, the potential applicability of the amylase enzymes of this new haloarchaea on the treatment of bakery waste was assessed and compared with a commercial amylase.

2. Materials and Methods

2.1. screening and selection of amylase producing haloarchaea.

The screening of amylase-producing haloarchaea was performed by detection of the extracellular amylase activity of the isolates on starch agar plates, with 20% NaCl. The plates were flooded with commercial Lugol’s iodine solution, 0.5% I 2 and 1% KI ( w/v ) (Chem Lab, Zedelgem, Belgium), every three days, to check the formation of degradation halos around the colonies. The isolate that presented the highest ratio of halo zone with respect to colony diameter was chosen for further studies. Screenings were done in triplicates.

2.2. Identification of the Selected Microorganism

Genomic DNA of the isolated amylase-producing strain was purified using the GeneJET Genomic Purification kit (Thermo Fisher Scientific, Waltham, MA, USA), following the manufacturer’s instructions. The quantification and the purity assessment of the genomic DNA obtained was done on a Nanodrop Spectrophotometer ND-1000 (Thermo Fisher Scientific). The full length of the 16S rRNA encoding gene was amplified with the archaeal specific primers 21F (5′-TTCCGGTTGATCCTGCCGGA-3′) and 1492R (5′-GGTTACCTTGTTACGACTT-3′). Polymerase chain reactions (PCR) were performed in a total volume of 25 µL, using an Eppendorf thermo-cycler. The reaction mixture contained—1 µL of genomic DNA, 0.2 U REDTaq ® DNA polymerase from Sigma Aldrich (St. Louis, MO, USA), and 2.5 µL of its specific 10× buffer that contained 10 pM of each primer, 0.2 mM dNTPs, and 2.5 mM MgCl 2 . The thermal profile was set to 0.5 min at 96 °C, 0.5 min at 55 °C, and 1 min at 72 °C for 30 cycles, followed by 10 min of final extension. The PCR products were analyzed by electrophoresis, on a 1% agarose gel to check their quality, and sent to Stabvida (Lisbon, Portugal) for Sanger sequencing. The 1.4 kb 16S rRNA gene sequences obtained were compared to those available at the GenBank and the European Molecular Biology Laboratory (EMBL) databases, using the Basic Local Alignment Search Tool (BLAST) at the National Center for Biotechnology Information (NCBI) [ 19 ].

2.3. Culture Conditions for Enzyme Production

All cultures were incubated at 37 °C with a shaking rate of 100 rpm, with either standard rich medium or minimal medium. The standard rich medium for archaea growth contained per liter—156 g NaCl, 13 g MgCl 2 ·6H 2 O, 20 g MgSO 4 ·7H 2 O, 1 g CaCl 2 ·6H 2 O, 4 g KCl, 0.2 g NaHCO 3 , 0.5 g NaBr, and 5 g yeast extract, with a pH value of 7, measured before autoclaving. For the minimal medium, yeast extract was substituted for 1% ( w/v ) of ammonium acetate. The amylase activity was induced by the addition of starch (3 g L −1 ) to either the rich or the minimal medium.

2.4. Partial Purification of Cell-Associated and Extracellular Amylases

Haloarcula sp. HS cells were first cultured in the rich medium, containing yeast extract and starch. When the culture reached the end of the exponential phase (OD 580 ≈ 3), cells were harvested through centrifugation, washed, and transferred to the minimal medium, where the biomass was cultivated until the extracellular starch was completely exhausted, about 3 days after the transference. Starch content was periodically measured every 24 h, by mixing 1 mL of culture medium with 5 µL of commercial Lugol’s iodine solution and by reading the absorbance at 580 nm. Then, the biomass was harvested by centrifugation for 20 min at 12,000 rpm and 4 °C. The supernatant was 100-fold concentrated by an ultrafiltration process in an Amicon ® system with a 10 kDa cut-off membrane and used as the source of the extracellular amylase. The specific amylase activity in the medium supernatant was 8 U mg −1 and it was increased to 350 U mg −1 in the concentrated supernatant, with a purification factor of 43.75. On its part, the cell pellet was disrupted by sonication in phosphate buffer (50 mM, 20% NaCl, pH 7) and centrifuged again to remove the cell debris and unbroken cells. The obtained cell extract was loaded onto a DEAE Sephacel TM column equilibrated with the same phosphate buffer. The absorbed proteins were eluted using a linear gradient of NaCl from 0 to 500 mM and a final washing with NaCl 1 M, with a flow rate of 15 mL h −1 . All fractions that presented amylase activity were collected and used as the cellular amylase source. In this case, the specific activity was increased from 20 U mg −1 in the crude cell extract to 120 U mg −1 in the partially purified preparation, with a purification factor of 6. Determination of the protein content in all the obtained extracts was performed according to the Bradford method [ 20 ], using bovine serum albumin (BSA) as standard.

2.5. Amylase Activity Assay

Unless otherwise indicated, the amylase activity was measured following the degradation of soluble starch by the standard iodine assay, based on the decrease of the absorbance at 580 nm of the iodine–starch complex produced by starch hydrolysis. The standard reaction mixture contained 50 µL of enzyme solution, 100 µL of 1% ( w/v ) potato starch solution in 20% NaCl, and 100 µL of phosphate buffer (50 mM, pH 7, 20% NaCl). The reaction mixture was incubated at 50 °C for 30 min, previously set as the best time to conserve the linearity of the activity. The reaction was stopped by cooling on ice and 100 µL were employed to reveal the remaining starch, by mixing 5 µL of four times diluted commercial Lugol’s iodine solution with the sample. Thereafter, 1 mL of distilled water was added to the sample before reading the absorbance at 580 nm. A standard curve was prepared with soluble starch. One unit of amylase activity was defined as the amount of enzyme degrading one microgram of starch per minute from soluble starch, under the assay conditions. To study the substrate specificity, potato starch was substituted by the indicated compounds (carboxymethyl cellulose, sucrose, and lactose) at a concentration of 1% ( w/v ), and incubated in the same conditions.

To analyze the starch hydrolysis products, aliquots were withdrawn from the incubation mixture at the initial reaction time and after 2 h of incubation at 50 °C. The reaction mixture contained 300 µL of the corresponding amylase extract and 600 µL of 1% ( w/v ) potato starch solution in 20% NaCl ( w/v ). The hydrolysis products were examined by a high-performance liquid chromatographic (HPLC) system (Merck-Hitachi LaChrom Elite), equipped with a refractive index detector (Merck-Hitachi L-2490) and an Aminex ® HPX-87H Column (Bio-Rad, Hercules, CA, USA), using an isocratic elution method with 5 mM H 2 SO 4 at 50 °C, and a flow rate of 0.6 mL min −1 . Glucose, maltose, and dextrin standards were obtained from Merck, Sigma-Aldrich (St. Louis, MO, USA).

2.6. Native Electrophoresis and Zymogram

The presence of amylase activity in the concentrated supernatant and the cell extract was revealed by in situ staining of a native PAGE containing 0.2% of soluble starch in the separating gel. A volume of 15 µL of the sample was mixed with 5 µL of loading dye and electrophoretically separated into two parallel gels of acrylamide, 10% supplemented with starch 0.2% ( w/v ) and run at 130 V. After electrophoresis, one of the gels was incubated in phosphate buffer (50 mM, pH 7, 20% NaCl) at 50 °C and 50 rpm for 1 h. Subsequently, the gel was stained with commercial Lugol´s reagent and the appearance of clear bands revealed the amylase activity. Meanwhile, the other gel was stained with 0.1% ( w/v ) Coomasie Brillant Blue R-250 in 45% ( v/v ) ethanol-10% ( v/v ) acetic acid, and faded with 25% ( v/v ) ethanol-10% ( v/v ) acetic acid. Molecular markers (NativeMark TM Unstained Protein Ladder, Thermo Fisher Scientific, Waltham, MA, USA) were used as a reference for the molecular weight of proteins. Molecular mass estimation of the proteins was calculated by plotting the log (MW) as a function of Rf (migration distance of the protein/migration distance of the dye front).

2.7. Effect of NaCl, Temperature, pH, Metals, and Detergents on the Amylase Activities of the New Isolated Strain Haloarcula sp. HS

The effect of salt concentration was evaluated until a maximum of 32% NaCl with intervals of 4% salinity increase. The desired NaCl concentration was obtained by adding the required NaCl to the phosphate buffer (50 mM, pH 7) and to the 1% ( w/v ) starch solution. The influence of temperature on cell-associated and extracellular amylase activities was studied in phosphate buffer (50 mM, pH 7, 20% NaCl), over the range of 30–80 °C, with temperature increments of 10 °C. For pH studies, amylase activity was measured at 50 °C and 20% of salt, in the following buffers—50 mM acetate for pH 2 and 3; 50 mM MES for pH from 4 to 6; and 50 mM Tris-HCl for pH from 7 to 11. All the assays were done at least in triplicates, and the results were presented as a percentage of relative activity.

To test the influence of different metals on cell-associated and extracellular amylase activities MgSO 4 , CaCl 2 , CuCl 2 , FeCl 2 , FeCl 3 , or EDTA (ethylenediaminetetraacetic acid), were added to the reaction mixture, in a final concentration of 10 mM. Similarly, the effect of various surfactants were studied, including Tween20 (Polyoxyethylene (20) sorbitan monolaurate), Tween80 (Polyoxyethylene (80) sorbitan monooleate), Triton-X100 (2-[4-(2, 4, 4-trimethylpentan-2-yl) phenoxy] ethanol), CHAPS (3-[(3-Cholamidopropyl) dimethylammonio]-1-propanesulfonate), SB-12 (N-Dodecyl-N,N-dimethylammonio-3-propane sulfonate), and SDS (sodium dodecyl sulfate), in a final concentration of 0.5% ( w/v ). Amylase residual activity was measured as previously detailed for the standard assay and expressed as a percentage, with respect to a control sample incubated in the absence of additives. All the determinations were conducted in triplicates.

2.8. Proteomic Analysis

For the proteomic analysis, the concentrated supernatant and the partially purified cell extract fractions with starch degrading activity were dialyzed for 48 h, against a solution of 1% NaHCO 3 and 0.01% EDTA in milli-Q water, to eliminate excess salt. The proteins were first precipitated with TCA/acetone and resuspended in ammonium bicarbonate-trifluoroethanol (50%). After that, the proteins were treated with dithiothreitol, 10 mM, and methyl ethanethiosulfonate, 10 mM. Prior to trypsin digestion, the samples were diluted with ammonium bicarbonate 25 mM, until the concentration of trifluoroethanol was under 5%. The digestion with trypsin was done overnight at 37 °C. Subsequently, the samples were analyzed by LC-MS/MS in a triple quadrupole-TOF system (5600 plus, ABSciex, Vaughan, ON, Canada), equipped with a nano-electrospray ion source, coupled to a nano-HPLC (Eksigent, Vaughan, ON, Canada). The Analyst TF 1.7 software was used for equipment controlling and data acquisition. Peptide mass tolerance was set to 25 ppm and 0.05 Da, for fragment masses, and only 1 or 2 missed cleavages were allowed. The peptide and protein identifications were performed using the Protein Pilot software (version 5.0.1, SCIEX, Vaughan, ON, Canada), with the Paragon algorithm. The search was conducted against the Uniprotproteome_Haloarcula_hispanica database 11_24_2020. The false discovery rate (FDR) was set to 0.01 for both peptides and proteins. Protein comparison was performed with the Basic Local Alignment Search Tool for proteins (BLASTp) of the NCBI (National Center for Biotechnology Information). The obtained sequences were analyzed using the CLC Workbench software (version 8, Qiagen, Hilden, Germany).

2.9. Identification of Amylase Coding Genes Based on Protein Sequences

With the aim of completing the full protein sequences of the amylases identified, the sequences of their encoding genes were amplified by PCR, using sets of primers specifically designed on the basis of the sequences of peptides obtained in the proteomic analysis. Concretely, six pairs of primers were employed to cover almost the full length of the DNA sequences of the three amylases found, obtaining two overlapping sequences for each amylase gene ( Table 1 ).

Sequences of the primers employed for the amylase encoding genes amplification.

Primer sets used for PCR amplification of the three amylase coding genes found in Haloarcula sp. HS. For each gene ( AMY_HS1 , AMY_HS2 , and AMY_HS3 ), two pairs of primers were designed on the basis of the sequences of the peptides obtained by proteomics.

Polymerase chain reactions were performed in a total volume of 25 µL, using an Eppendorf thermo-cycler. The reaction mixtures contained—1 µL of genomic DNA, 0.2 U REDTaq ® DNA polymerase from Sigma Aldrich (St. Louis, MO, USA), and 2.5 µL of its specific 10× buffer that contained 10 pM of each primer, 0.2 mM dNTPs, and 2.5 mM MgCl 2 . The thermal profile was set to 0.5 min at 96 °C, 0.5 min at 62 °C, and 1 min at 72 °C for 30 cycles, followed by 10 min of final extension. The PCR products were analyzed by electrophoresis on a 1% agarose gel and sent to Stabvida (Lisbon, Portugal) for Sanger sequencing. The sequences obtained were translated to protein and both, DNA and protein sequences, were compared to those available at National Center for Biotechnology Information (NCBI) databases, using the Basic Local Alignment Search Tool (BLAST). Finally, different alignments were conducted in the CLC Workbench software (version 8, Qiagen), the predicted structural models were built using the Phyre2 [ 21 ] and NetSurfP [ 22 ] online web servers, and three-dimensional (3D) molecular graphics were analyzed in the UCSF Chimera version 1.15 [ 23 ] (University of California, Oakland, CA, USA). Physicochemical characteristics of the proteins were obtained using the ProtParam tool (ExPASy) [ 24 ].

2.10. Starch Hydrolysis from Bakery Waste

Bread from bakery waste was chosen for the present experiment. Bread crumbs were dried on a stove at 70 °C and milled in a porcelain mortar to obtain a fine powder. Starch was recovered by mashing dried crumbs in distilled water, in saltwater at 20% NaCl ( w/v ), and in saturated saline solution (33% NaCl). The ability of the extracellular amylase of Haloarcula sp. HS to degrade the starch from bread was comparatively tested against a commercial α-amylase (Megazyme cat. no. E-BSTAA). Starch and enzyme solutions were mixed in a proportion of 1:1 ( v/v ) in a final volume of 1 mL. The hydrolysis of the starch was performed for 15 min at 60 °C in 50 mM acetate buffer pH 5. The amount of remaining starch was measured by the iodine-starch method. A control, containing starch recovered from bread without the enzyme solution, was incubated in the same conditions. All assays were conducted in triplicates.

3.1. Selection of Amylase-Producing Haloarchaea Isolated from Odiel Salterns Ponds

An in vitro screening was carried out to select the best amylase-producing strain, among a collection of archaea previously isolated from the saltern ponds of the Odiel Marshlands (SW, Spain) with a salinity of 33%. Amylase activity of each isolate was screened for 9 days on starch-agar plates, as detailed in Material and Methods ( Figure 1 A). Eight colonies showed a considerable amylase activity, and that with the largest halo was selected and identified, by amplification and sequencing of its 16S rRNA full-length coding gene ( Supplementary Material, Figure S1 ), followed by the comparison of the obtained sequence with the NCBI database using the BLASTn tool. The results showed that the selected strain was closely related to the Haloarcula genus, showing 98% homology with different species of this taxonomic group. Therefore, the new strain isolated was named Haloarcula sp. HS.

( A ) In vitro selective screening for amylase-releasing haloarchaea. Semi-quantitative estimation of amylase activity from four haloarchaeal strains isolated from the Odiel Marshlands, grown on starch agar plates ( top ) and revealed with Lugol´s iodine solution ( down ), as detailed in the Material and Methods section. ( B ) Molecular phylogenetic analysis by the maximum likelihood method. The tree represents a comparison among the complete 16S rRNA coding gene sequences, including a series of reference haloarchaeal species and the new isolated strain, Haloarcula sp. HS. Multiple alignments were generated by MUSCLE (MUltiple Sequence Comparison by Log-Expectation) and the tree was constructed with MEGA X. The numbers at the nodes indicate the bootstrap values calculated for 1000 replicates. Arrows point to the new strain Haloarcula sp. HS.

Molecular phylogenetic analysis was performed using the Molecular Evolutionary Genetics Analysis (MEGA X) [ 25 ], on a series of reference haloarchaeal species and on the new isolated strain, Haloarcula sp. HS ( Figure 1 B). The hyperthermophilic archaea Methanococcus vulcanus was used as an outgroup and the bootstrap was set at 1000 replicates. The 16S rRNA encoding sequence of the isolate clustered with the corresponding genes of the representatives of the Haloarcula genus, especially close to the species Haloarcula hispanica.

3.2. Optimization of a Two-Stage Culture Strategy to Induce the Production of Amylase

In the studied archaea, significant levels of amylase activity were only found when the biomass was grown under inductive conditions in the presence of starch. The culture conditions that induced the production of amylases are widely studied for bacteria and hyperthermophilic archaea, as reviewed by Mehta and Satyanarayana [ 5 ], but more limited information exists on the production of amylase in haloarchaea [ 26 , 27 ]. Most authors agree that amylase production is growth-associated and is strongly induced by starch.

To establish the best culture conditions for the production and excretion of amylase by Haloarcula sp. HS, the haloarchaea was cultured in a (i) rich medium, which contained yeast extract and a (ii) minimal medium, in which the yeast extract was substituted by ammonium acetate. In both cases, starch (3 g L −1 ) was added to the culture medium, as detailed in Materials and Methods. The optical density of the cultures, protein secretion into the media, and hydrolysis of extracellular starch were followed in both, rich and minimal medium cultures. As shown in Figure 2 , cell growth and protein secretion were higher when the microorganism was grown in the rich media, which contained yeast extract. In this medium, the studied archaea reached the stationary phase of growth in about 4 days and excreted 3 mg L −1 of proteins into the culture medium. The archaea cultured in the minimal medium exhibited very slow growth and excreted much fewer proteins to the culture medium, about 1 mg L −1 , after 30 h of culture. However, starch degradation activity was much higher for the archaea cultured in the minimal medium, which despite a much lower biomass, showed an initial starch degradation rate 5.5 times higher than that of the rich medium. Although starch was completely hydrolyzed in both media, there was a 24 h lag phase before starch degradation started in the medium with yeast extract, probably due to the presence of more easily assimilable carbon sources in this medium.

Time course evolution of Haloarcula sp. HS cultures in rich and minimal media. Optical density ( A ), secretion of proteins ( B ), and starch hydrolysis ( C ) were measured during the time of culture in rich (■) and minimal ( ♦ ) broths. All data are expressed as the mean ± SD of at least triplicate experiments.

For this reason, a two-step culture was set up to get both, a high biomass and amylase productivity. Cells were first grown in a rich medium in order to obtain a large amount of biomass, and when the culture reached the end of the exponential phase of growth, at the third day of culture, the cells were transferred to the fresh minimal medium to induce the production of amylase, and was cultured for another 4 days. Through this two-step approach, high starch consumption activity and a high protein excretion were achieved, reaching an extracellular protein concentration of 10 mg L −1 and undetected levels of extracellular starch, on the 7th day of culture ( Figure 3 ).

Time course evolution of Haloarcula sp. HS in the two-step culture. Optical density, secreted proteins, and starch concentration were measured along the full cultivation time. The red arrow indicates the moment of transference of the cells from the rich to the fresh minimum medium. All data are expressed as the mean ± SD of at least triplicate experiments.

3.3. Extracellular and Cell-Associated Amylase Activities in Haloarcula sp. HS

To identify the enzyme responsible for the amylase activity and characterize its properties, the haloarchaeal strain Haloarcula sp. HS was grown in a two-stage culture with starch (3 g L −1 ), as previously described ( Figure 3 ). Cell-associated proteins and the concentrated extracellular proteins excreted into the cultured medium were electrophoretically separated in a polyacrylamide gel containing starch. After electrophoresis, the gel was split lengthwise with a razor blade. One half was stained with Coomassie blue and the other with Lugol’s iodine solution to detect both proteins and amylase activity, respectively. The zymogram analysis proved the presence of amylase activity in both samples, the culture medium, and the crude extract. A unique band with amylase activity was observed in the culture medium, after a 100-fold concentration step through ultrafiltration with a 10 kDa cut-off membrane, as indicated in Material and Methods. However, in the crude extract, two bands with starch hydrolyzing activity were observed, indicating the presence of several cell-associated enzymes with amylase activity ( Figure 4 and Figure S2 ). The crude extract was partially purified through ion-exchange chromatography in DEAE Sephacel TM , as indicated in the Materials and Methods section. All fractions with amylase activity were pooled and used as the source of cellular amylase. The electrophoretic analysis of the purified extracts showed a unique band with amylase activity ( Figure 4 A). The size of the observed bands was between 20 and 27 kDa, however, the protein mobility was strongly affected by the starch added to the polyacrylamide gel and these apparent sizes observed were not representative.

Native-PAGE and zymogram of cell-associated ( A ) and extracellular ( B ) proteins obtained from Haloarcula sp. HS. Lanes 1, 3, and 5—samples on native-PAGE followed by Coomassie Blue staining. Lanes 2, 4, and 6—samples on native PAGE followed by Lugol´s solution staining. Lane M—molecular mass marker in kDa, lanes 1 and 2—crude extract, lanes 3 and 4—partially purified crude extract, and lanes 5 and 6—concentrated supernatant. The whole gels for each staining are available in Supplementary Material, Figure S2 .

3.4. Characterization of Extracellular and Cell-Associated Amylase Activities

A series of in vitro experiments were carried out with the extracellular and cellular amylase-enriched extracts to characterize the hydrolysis products, the substrate specificity, and the optimal kinetic parameters of the amylase activity of these extracts. Both, extracellular and cellular, enzymatic preparations were incubated with 1 mg of starch in the standard conditions, described in Material and Methods, excepting that the incubation time was fixed at 2 h. The products of starch hydrolysis were identified by HPLC and an IR detector, as detailed in Materials and Methods. A parallel reaction with a commercial α-amylase purchased from Megazyme (cat. no. E-BSTAA) was done in the same conditions for comparison. The results ( Table 2 ) revealed that starch degradation was almost complete in all cases, being especially efficient in the case of the extracellular amylase extract, with a remaining starch of only 1.7%. The main product obtained with the three enzymatic sources was maltose, which represents between 73.8% and 86.1% of the total carbohydrate content in the reaction mixtures. In addition, the enzymatic preparation from the cellular extract was also able to catalyze the liberation of glucose (6.6%). On the other hand, the extracellular extract and the commercial reference amylase catalyzed the liberation of dextrins, which supposed 18.5% and 20.8% of the total carbohydrate content in the reaction mixture, respectively, in addition to maltose. No glucose was found as the end product in these reactions ( Table 2 ).

Percentage of the different products obtained from starch hydrolysis.

Comparison of the products obtained from the hydrolysis of starch by cell-associated or extracellular partially purified extracts of Haloarcula sp. HS and a commercial α-amylase. The percentage (%) of dextrins, maltose, and glucose produced and the remaining starch are indicated as the mean of three replicates with the corresponding standard deviation. ND, not detected.

With respect to the substrate specificity, neither extracellular nor cellular Haloarcula sp. HS extracts were able to hydrolyze other glucose polysaccharides, such as carboxymethyl cellulose, or disaccharides, such as sucrose or lactose, which not contain alpha-1,4-linked glucose.

The most characteristic feature of halophilic enzymes is their ability to operate under very high salinities. As it is shown in Figure 5 , the extracellular enzymatic preparation presented the optimal activity at 28% salt and retained less than half of its activity when the salt content was under 20% ( Figure 5 A), showing that it is more salt-dependent than the cellular one, which instead showed the maximum activity at 16% salinity and retained more than 60% of its activity at all salinities studied ( Figure 5 B).

Effect of salt, pH, and temperature on amylase activities. Relative amylase activities in different salt concentrations (%, w/v ), pH values, and temperatures are shown for the extracellular ( A , C , E ) and cellular ( B , D , F ) extracts. Relative activity was defined as the percentage of maximum activity for each case. The 100% activity corresponded to 70 ± 6.4 U mL −1 (350 U mg −1 ) for the extracellular amylase extract and 60 ± 5.6 U mL −1 (120 U mg −1 ) for the mix of cell-associated amylases. Mean and standard deviations are shown.

The influence of pH in both amylase activities was studied in different buffers, as detailed in Materials and Methods. The optimal activity was found at pH 5 for the extracellular amylase, and at pH 7 for the cellular amylase-enriched extract, as shown in Figure 5 C,D, respectively. It should be noted that the cell-associated activity was stable under a wide range of pH values, retaining more than 50% activity at pH values between 2 and 11, and more than 80% activity at pH values comprising pH 5 to 9. Contrarily, the extracellular amylase activity appeared to be more susceptible to extreme pH, losing more than 50% of its activity both at low and high pH values.

The effect of the temperature on the amylase activities showed, once again, that the extracellular amylase activity had a higher dependence on the physicochemical parameters of the assay than the cell-associated one. The extracellular amylase activity showed an optimal temperature of 60 °C, losing more than 65% of its activity below 50 °C or above 60 °C ( Figure 5 E). The cellular amylase-enriched extract, on the other hand, conserved a high activity over a wide range of temperatures, retaining more than 75% of activity from 30 to 80 °C ( Figure 5 F). This weak temperature dependence was due to the fact that a mix of three different cell-associated enzymes could contribute to the amylase activity, as later shown by the proteomic analysis of the cellular enzymatic preparation.

3.5. Effects of Metals and Surfactants on the Amylase Enzymatic Activities

Many microbial α-amylases are reported to be calcium-dependent metalloenzymes [ 9 ]. Therefore, the effect of EDTA, Ca 2+ , and other metallic ions including Mg 2+ , Cu 2+ , Fe 2+ , and, Fe 3+ was tested. The results revealed that the addition of Ca 2+ , Fe 2+ , or Mg 2+ causes a slight increase in the amylolytic activity of both extracellular and cellular enzymatic extracts ( Figure 6 ). The possible existence of divalent metals in the partially purified enzymatic preparations makes it difficult to obtain accurate conclusions on the effects of these metals on the amylase activities of Haloarcula sp. HS. However, the strong inhibition observed in the presence of the metal chelating agent EDTA confirmed the divalent cation dependence of both, extracellular and cell-associated amylase activities, which decreased to 39% and 33%, respectively, in the presence of EDTA.

Effect of metal ions on the amylase activities. Extracellular and cell-associated relative amylase activities under the presence of different metal ions and EDTA (10 mM) are represented. Relative activity was defined as the percentage of maximal activity with respect to control, with no additives. The control activity was 70 ± 6.4 U mL −1 (350 U mg −1 ) for the extracellular amylase and 60 ± 5.6 U mL −1 (120 U mg −1 ) for the cellular amylase. Mean and standard deviations are shown.

On the other hand, the presence of Cu 2+ and Fe 3+ in the reaction mixture, at a concentration of 10 mM, caused a drastic reduction in both, extracellular and cell-associated amylase activities ( Figure 6 ). The cellular extract only retained 28% of its amylase activity in Fe 3+ ; and similarly, the extracellular amylase conserved 26% of its activity. Under the presence of the cupric ion, the activity of the extracellular amylase dropped to 8%, while the activity of the cell-associated amylase decreased to 33%. This fact suggests the involvement of tiol/carboxyl groups, typically inhibited by Cu 2+ , in the function of the enzymes [ 28 ].

The in vitro effect of anionic (SDS), cationic (SB-12), zwitterionic (CHAPS), and no ionic (Tween 20, Tween 80, and Triton X-100) detergents on the amylase activities were assayed. The obtained results revealed that both activities were very stable in different detergents, retaining more than 80% activity in all, with the exception of SDS, which caused a decrease by almost half in the amylase activities of both, extracellular and cellular amylase enriched extracts.

3.6. Identification of Amylases in Cellular and Extracellular Concentrated Extracts of Haloarcula sp. HS by a Proteomic Approach

A proteomic study of both, extracellular and cellular partially purified extracts with amylase activity, was carried out to identify the sequence of the proteins responsible for these starch-degrading activities in Haloarcula sp. HS. Both samples were submitted to tryptic digestion and analyzed by LC-MS/MS in a triple quadrupole-TOF system, as described in Materials and Methods.

The results revealed that the main extracellular protein secreted to the culture media was α-amylase. The rest of the proteins identified in the extracellular fraction were membrane-ligated proteins, probably from broken cell remains. This extracellular amylase (AMY_HS1) was identified by 42 unique peptides, presenting 73.56% identity and 60% query cover with the α-amylase of Haloarcula hispanica N601 (UniProt: V5TMJ3_HALHI). For its part, in the cellular fraction, three different amylases were found. One of them corresponded to the same α-amylase (UniProt: V5TMJ3_HALHI) detected in the extracellular fraction, which in this case was identified according to 13 unique peptides and showed 81.48% identity and 36% query cover. The other two proteins, denoted as AMY_HS2 and AMY_HS3, showed high homology with two different α-amylases of Haloarcula hispanica N601 (UniProt codes: V5TRA6_HALHI and V5TQD3_HALHI, respectively), both determined according to 15 unique peptides. AMY_HS2 presented 45.25% identity and 65% query cover with V5TRA6_HALHI, while AMY_HS3 showed 75.76% identity and 31% query cover with V5TQD3_HALHI.

Due to the low percentage of protein covering achieved, the sequences of amylase coding genes were amplified by PCR, as detailed in Material and Methods, with primers designed to target the peptide sequences identified by proteomics for each enzyme. Through this approach, practically the full length of each protein sequence was completed.

The alignment of the three obtained protein sequences revealed that the extracellular amylase only presented a 17% identity with the cell-associated amylases, which in turn showed a 38% identity between them. Nonetheless, as it is shown in their predicted three-dimensional ( Figure 7 ) and secondary structures ( Supplementary Material, Figure S3 ), the three amylase sequences from Haloarcula sp. HS conserve the typical structural domains of the GH-13 family, according to the Carbohydrate-Active enZYmes (CAZY) database; including the catalytic (β/α) 8 -barrel (TIM-barrel) located in the domain A, a small domain B located in the loop between the β 3 -strand and the α 3 -helix of the barrel, and the domain C showing an antiparallel β-sandwich structure in the C-terminal end of the protein. In addition to this typical conformation, the two cellular amylases show an N-terminal domain, which was reported in some maltogenic amylases and seems to be involved in increasing the binding of the enzyme to raw starch. This N domain forms a large groove, the N–C groove, which might be responsible for thermostabilization via oligomerization and substrate affinity modifications in some microbial maltogenic amylases [ 29 ]. For its part, the extracellular protein, AMY_HS1, presents the conserved TAT (Twin-Arginine-Translocation) motif and its corresponding processing site ( Supplementary Material, Figure S4 ).

Three-dimensional predicted structures of the three amylase sequences identified in Haloarcula sp. HS; ( A ) mature extracellular amylase, AMY_HS1; ( B ) cellular amylase, AMY_HS2; ( C ) cellular amylase, AMY_HS3. Phyre2 software and the Chimera program were employed for 3D structure visualization. Helixes are represented by blue ribbons and strands by red arrows.

Regarding the physicochemical characteristics, the three alpha-amylases of Haloarcula sp. HS had a low isoelectric point, negative net charge, and low hydrophobicity ( Table 3 ) as other haloarchaeal enzymes.

Physicochemical properties of the amylase enzymes from Haloarcula sp. HS.

Main physicochemical characteristics of the amylase enzymes found in the Haloarcula sp. HS strain. N, number of nucleotides; MW, molecular weight; IP, theoretical isoelectric point; Z, net charge; GRAVY (Grand average of hydropathicity), mean of the hydropathy index of each amino acid residue. The aliphatic index stands for the relative volume occupied by the aliphatic side chains.

Moreover, although the three protein sequences had a low percentage of sequence homology, they conserved the catalytic triad of aspartate, glutamate, and aspartate in the active site, along with other conserved residues that were described to be indispensable for the structure of the enzyme [ 30 ]. These conserved amino acids are shown in Figure 8 . One of the first consensus amino acids found is aspartic acid, which is essential for active site integrity. This aspartic residue is in the position Asp92 for the mature AMY_HS1 protein after TAT processing, and occupies the position 359 and 303 in AMY_HS2 and AMY_HS3, respectively (Asp92/359/303). Following the same notation, the rest of the conserved amino acids are distributed as follows—Asn96/363/307, which coordinates the conserved calcium ion between the A and B domains [ 31 ]; and His93/360/304, which stabilizes the interaction between the C-terminal of β3 and the rest of TIM barrel through hydrogen bonding to Asn61/328/272 and the backbone oxygen of Tyr57/324/268. The first catalytic residue is Asp177/446/379, located in β4, which is preceded by Arg175/444/377, both of which are indispensable amino acids for the catalytic activity. Lys and His are usually present in this region in the position Asp+3 and +4, binding the reducing end of the glucose chain in the substrate-binding cleft [ 32 ], however, these residues were only found in the extracellular amylase. The second catalytic residue is the proton donor Glu205/475/408, which lies in the fifth L-strand of the TIM-barrel. The following conserved residues protect the active site from the solvent and contain the last catalytic amino acid Asp268/539/471, postulated to be involved in substrate binding, substrate distortion, and in elevating the pKa of Glu205/475/408. This residue is usually accompanied in α-amylases by His, Asn, Val, and Phe in the positions −1, −2, −4, and −5, respectively, as it occurs in AMY_HS1, while in AMY_HS2 and AMY_HS3, Phe is substituted by Tyr, and Val is changed by Ala in AMY_HS3. Finally, the other two conserved residues were found in Gly287/564/496, followed by Pro289/566/498 [ 33 , 34 , 35 , 36 ].

Alignment of the three amylase sequences from Haloarcula sp. HS. The mature extracellular protein is named AMY_HS1, while the cell-associated amylases are denoted as AMY_HS2 and AMY_HS3. Purple stars highlight the catalytic triad (Asp-Glu-Asp), blue star denotes the canonical calcium-binding site and the black stars point other essential residues for enzyme structure. Identical residues in the three sequences are shaded in white, residues that coincide in two of the sequences or do not coincide at all are shaded in pink and red, respectively.

Finally, a multispecies study was carried out to compare the degree of conservation among the amylases reported and those from different haloarchaeal members. The protein sequence of the extracellular amylase (AMY_HS1) was aligned and compared to various extracellular amylase sequences available in the NCBI database, selecting different representatives of the order Halobacteriales ( Supplementary Material, Figure S4 ). Among these amylase sequences, different percentages of identity were found with the extracellular amylases from Haloarcula hispanica N601 (90%), Halomicroarcula salina (73%), Halapricum salinum (57%), and Haloterrigena turkmenika (40%). Likewise, the two cell-associated amylases (AMY_HS2 and AMY_HS3) from Haloarcula sp. HS showed the following percentages that identity with the amylase sequences from Haloarcula hispanica N601 (90 and 80%, respectively), Halomicroarcula salina (70 and 65%, respectively), Haloferax mediterranei (59, and 50%, respectively), Halogeometricum rufum (57% with AMY_HS2), and Halogeometricum limi (49% with AMY_HS3) ( Supplementary Material, Figures S5 and S6 ). In all alignments, it could be appreciated that the catalytic regions were conserved among the different haloarchaeal species ( Supplementary Material, Figures S4–S6 ).

3.7. Hydrolysis of Bakery Waste

Bakery residues, including dough, flour dust, burnt or rejected bread, and biscuits, can be exploited for the production of fermentable sugars. Among them, bread was chosen as the substrate for this assay, as it is one of the most abundant food waste products worldwide. In addition, many of the discarded by-products during the bread manufacturing process are fundamentally constituted by starch, such as substandard bread and the bread crust removed to make special types of sandwich bread [ 37 ].

The extracellular amylase activity was selected for this assay, as it works better in high NaCl concentrations than the cell-associated amylase. This was compared to a commercial thermostable α-amylase. Sodium chloride was found to have a complex effect on the gelatinization and rheological properties of starch. Some studies pointed out that the enthalpy for the gelatinization process decreased at high salt concentrations, an effect of great importance for the production and properties of several cereal-based products, as well as for the manufacture of modified starches [ 38 ].

The results proved that a high percentage of the initial starch (75–85%) was hydrolyzed by both enzymes, with maximum activities above 100 U mL −1 . However, the concentration of salt was determined in their maximum activities. The extracellular amylase hydrolyzed around 75% of the starch under 20% salt or even under salt saturation, with maximum activities of 107 and 105 U mL −1 , respectively; while when no salt was added, the degradation rate decreased to 22%, with an activity of 24.7 U mL −1 . Conversely, the commercial α-amylase presented the highest activity (101.8 U mL −1 ) when no salt was added to the mixture, hydrolyzing 85% of the starch; and its activity dropped drastically to 6–7 U mL −1 , under elevated salt concentration, degrading only 5–6% of the starch.

The obtained hydrolysate was very rich in simple sugars that could now be used for the bioproduction of many high-value molecules like glycerol, hydrogen, ethanol, or lactic acid, among others. These molecules are required for different purposes, including renewable energy sources, fuel additives, and food preservers [ 13 ]. To our knowledge, this study supposes the first attempt to use a halophilic amylase to degrade starch from bread into simple sugars. The applicability of the amylase from a haloarchaea was tested using starch from agricultural waste [ 39 ], however, the starch content of this residue was considerably lower, and also the starch extraction method was costlier.

4. Discussion

Alpha-amylases are a large family of endo-glycosyl hydrolases that cleave the internal α,1-4 glycosidic bonds between the glucose units in polysaccharides, such as amylose and amylopectin. They are common in all kingdoms of life and their general properties, three-dimensional structures, and mechanisms of action are extensively reviewed [ 40 , 41 , 42 , 43 ].

Alpha-amylases are particularly spread among microbial species, such as bacteria and yeasts. These amylases are often extracellular enzymes that allow microorganisms to use environmental polysaccharides for their nutrition. Thermostable α-amylases were produced and commercially exploited from yeast and bacterial species, such as Bacillus subtilis , B. licheniformis , or B. amyloliquefacines [ 14 ]. Some archaea, including halophilic archaea, were found to produce halotolerant α-amylases, which in addition were highly or moderately thermotolerant. The extracellular α-amylases of Haloarcula sp. S-1 [ 44 ], N. amylolyticus [ 45 ], H. mediterranei [ 26 ], H. xinjiangense [ 46 ], Haloferax sp. HA10 [ 47 ], H.turkmenica [ 39 ], or Halococcus GUVSC8 [ 48 ] are some examples. Although much less studied, the intracellular α-amylases of some haloarchaea, such as Haloarcula japonica [ 49 ] were also characterized.

The new strain isolated from the Odiel Marshlands and selected by its high ability to degrade starch in iodine–starch agar plate assays was foumd to be closely related to the genus Haloarcula , as shown in the phylogenetic tree ( Figure 1 ). The high homology of its 16S rRNA with that of Haloarcula hispanica (98%) or Haloarcula japonica (97%) confirmed it. We found that the new strain exhibited a high amylolytic activity when cultured in the presence of starch; this activity was higher in a minimal medium with ammonium acetate ( Figure 2 ).

In agreement with our observations, several reports indicate that amylase production in haloarchaea is induced by starch, as described for Halorubrum [ 46 ], Haloferax [ 47 ], Haloarcula [ 50 ], and some halophilic bacteria [ 51 ]. Other culture conditions and nutrients that were reported to influence the induction of alpha-amylases are nitrogen, metal ions, or phosphate [ 5 ]. Pérez-Pomares et al. [ 26 ], for example, reported low amylase excretion when using a minimal medium containing ammonium acetate as carbon and nitrogen source, plus starch in Haloferax mediterranei.

Our observations indicate that the conditions that induce the amylase activity are not the best for growth ( Figure 2 ). Therefore, a two-step culture was set up, in which a large amount of biomass was obtained in a rich medium, followed by transfer to a minimal medium with starch, to induce the production of amylase activity ( Figure 3 ). This two-stage method allowed the improvement of amylase production, and at the same time facilitated the recovery of the extracellular enzyme. Since the minimal medium had no yeast extract, there are no foreign proteins that could interfere with the amylase secreted into the culture medium.

Partially purified extracellular and cellular-amylase-enriched extracts were obtained from Haloarcula sp. HS, through ultrafiltration and anion exchange chromatography, respectively, and were electrophoretically separated in native conditions. In situ staining of the obtained acrylamide gel allowed the identification of bands with starch degrading ability, one band in the extracellular enzymatic preparation, and two bands in the cellular extract, with apparent relative molecular masses between 21.6 and 30.4 kDa ( Figure 4 ). The zymogram indicates that the new isolated strain presents amylase activities in both, the supernatant and the crude cell extract, suggesting that there is more than one cell-associated amylase that does not coincide with the extracellular one. The proteomic analysis of the extracts and the subsequent amplification of the whole gene sequences that encode for these amylases allowed us to confirm this hypothesis ( Figure 8 ), and also indicated that the real masses of the amylases were much higher than the apparent molecular masses shown in the electrophoresis gel. These differences could be due to the fact that the electrophoresis was carried out in native conditions and in the presence of starch, which could modify their electrophoretic mobilities, besides the fact that halophilic proteins usually show altered electrophoretic properties [ 52 ].

The apparent molecular masses reported for the amylases of other haloarchaea are slightly higher than the molecular weight observed for the amylases of Haloarcula sp. HS. For example, the intracellular amylase of Haloarcula japonica presented a molecular mass of 83 kDa [ 49 ]; the extracellular amylases of Haloterrigena tukmenica and Haloferax sp. HA10 showed a molecular weight of 66 kDa [ 39 , 47 ], in Haloferax mediterranei , the weight of the enzyme was around 50–58 kDa [ 26 ], 60 kDa in Halorubrum xinjiangense [ 46 ], and 74 kDa in Natronococcus sp. Ah-36 [ 45 ], while in the Haloarcula species, the molecular mass varied from 43 to 70 kDa [ 27 , 44 ]. The physicochemical parameters of the new alpha-amylases, with low isoelectric point and negative net charge ( Table 3 ) also meet the usual characteristic of haloarchaeal enzymes, as reported by other authors. Yan and Wu [ 43 ] analyzed the sequences of 88 α-amylases from archaea and observed that amylases from haloarchaea have a highly negatively charged surface, and a higher percentage of acidic residues as a mechanism of adaptation to high salinity. Other authors describe similar features for H. hispanica , which has an extracellular amylase with an isoelectric point of 4.2 and a low level of aromatic and hydrophobic residues [ 27 ]. In haloarchaea, most studies about amylases focused on extracellular enzymes, given that sometimes no activity was found in the crude cell extract, as was observed in Haloferax mediterranei and Halorubrum xinjiangense [ 26 , 46 ] or because, as in the case of Haloterrigena turkmenica , the amylase activity was quite higher in the supernatant than in the cell extract [ 39 ]. With respect to the Haloarcula genus, Hutcheon et al. confirmed the overexpression of an extracellular amylase (AmyH) in the mutant strain Haloarcula hispanica B3, which was secreted in a folded conformation via the TAT (Twin-Arginine-Translocation) pathway, indicating that it was active in the cytoplasm before the secretion to the media [ 27 ]. Additionally, Onodera et al. reported the overexpression of an intracellular amylase (malA), which was not secreted to the media in Haloarcula japonica [ 49 ]. Based on the above mentioned, it seems that there were diverse amylases with probably different modes of action, which to our knowledge, are not yet deeply elucidated.

Maltose is the main end-product released from the starch hydrolysis by the extracellular and cellular partially purified amylase extracts of Haloarcula sp. HS ( Table 2 ). This is the main product of maltogenic α-amylases, like most α-amylases from haloarchaea, e.g., intracellular α-amylase from Haloarcula japonica [ 49 ] or the extracellular α-amylase from Haloterrigena tukmenica [ 39 ]. A small proportion of glucose was found in the assay catalyzed by the cell extract. However, it is difficult to predict if it is directly due to the action of the cellular amylases in Haloarcula sp. HS or due to the contribution of additional cell-associated enzymes.

With regards to the optimal enzymatic parameters, both cell-associated and extracellular starch-degrading activities exhibited their maximum activities around 60 °C. The low-temperature dependence of the cell-associated amylase, which only loses around 25% of its activity in the temperature range 20–80 °C, is noteworthy. However, it should be considered that this activity might be, as shown in this study, the result of the action of three different amylase enzymes. This fact contributes to broadening the range of optimal temperatures for the cell-associated amylase activity. High retention of enzyme activity over a wide range of temperatures was reported for other partially purified amylases, such as that from Haloferax sp. HA10 [ 47 ], which showed the highest amylase activity at 55 °C.

The optimal pH values were 7 for the cellular amylase activity, and 5 for the extracellular activity. Additionally, extracellular and cell-associated, amylase activities were extremely halophilic, showing their maximum activities at 25% NaCl. Therefore, it is noteworthy that the cell-associated amylase activity seemed to be more tolerant to changes in salinity, pH, and temperature than the extracellular one ( Figure 5 ). This was probably due to the presence of three different amylases in the cell extract, as was later confirmed in the proteomic analysis. In addition, the high salt and temperature tolerance could be of interest for many industrial applications in which these extreme conditions are needed.

A comparison with the optimal parameters reported for amylases of other related haloarchaea is summarized in Table 4 . Most extracellular α-amylases from haloarchaea showed their best activity at temperatures from 45 to 70 °C, pH from 6.5 to 8.7, and in salt concentrations from 1 to 5 M. The extracellular amylase found in Haloarcula sp. HS is one of the most halophilic and acidophilic α-amylase described within the haloarchaea group ( Table 4 ).

Optimal parameters reported for α-amylase activity in haloarchaea.

Additionally, extracellular and cell-associated amylase, activities from Haloarcula sp. HS, exhibit a strong inhibition in the presence of the metal chelating agent EDTA ( Figure 6 ). In addition, the analysis of the amylase sequences obtained allowed the identification of the canonical Ca-binding residue in the three amino acidic sequences, as shown in Figure 8 , indicating that they must be calcium-dependent. Dependence of calcium is a common feature within haloarchaeal amylases, as was reported for Haloferax mediterranei , Haloarcula hispanica , Haloarcula sp. S-1, Halorubrum xijiangense , and [ 26 , 27 , 44 , 46 ]. However, some haloarchaeal amylases showed to be resilient to EDTA, indicating no dependence on Ca 2+ , as revealed by the studies on Haloterrigena turkmenica and Haloarcula japonica [ 39 , 49 ].

Furthermore, both amylase activities showed high stability in most tested surfactants, excepting the anionic detergent SDS. There are few reports on the stability of amylase from haloarchaea on surfactants. In this context, detergent-stable amylases were recently found in Haloterrigena tukmenica , Halorubrum xijiangense , and Haloferax sp. HA10 [ 39 , 46 , 47 ]. Additionally, a surfactant-stable amylase was characterized from the halophilic bacteria Nesterenkonia sp. F [ 53 ]. Therefore, to the best of our knowledge, the two detergent-stable amylase activities described in this work entailed the first report on this specific feature of the amylase activity from the Haloarcula genus.

Analyzing the sequences of the peptides generated by the tryptic digestion of the partially purified extracellular and cell extracts, it was possible to identify three different amylases in Haloarcula sp. HS. One (AMY_HS1) was found both in cells and in the culture medium, while the other two amylases (AMY_HS2 and AMY_HS3) were exclusively found in the cell extract.

Despite the low percentage of sequence identity that the three amylases of Haloarcula sp. HS shared ( Figure 8 ), the three enzymes exhibited a high three-dimensional structure homology, with the three typical domains of alpha-amylases of the glycosyl hydrolase GH-13 family ( Figure 7 ) and many of the key conserved residues ( Figure 8 ). For example, the three amino acids (Asp-Glu-Asp), which constituted the active site of alpha-amylases, were identified in α-amylases of haloarchaeal strains, such as Halogeometricum borinquense [ 30 ] and Haloterrigena turkmenica [ 39 ]. These residues were conserved in all alpha-amylases of the GH-13 family of many different origins, which were compiled in the Carbohydrate-Active enZYmes (CAZy) database [ 54 ].

The calcium-binding domain located between the 3rd beta-strand and the 3rd alpha-helix contained an asparagine amino acid found in the three amylases of Haloarcula sp. HS ( Figure 8 ), as was described for other haloarchaeal amylases [ 30 ] and many other α-amylases, which were calcium-dependent metalloenzymes [ 55 ]. In some cases, more than one Ca-binding domain was described, as in H. orenii [ 56 ]. Additional conserved amino acids reported in alpha-amylases, which help to stabilize the structure or the binding of the substrate [ 5 ], were found in the three amylases of Haloarcula sp. HS ( Figure 8 ).

Most available archaeal amylase sequences were from the thermophilic archaea and could work at very high temperatures, which was of great industrial interest. The amylases identified from the Odiel Marshlands were medium or highly thermotolerant, in addition to being extremely halophilic. The α-amylases from hyperthermophilic archaea were closely related to plant amylases [ 57 ]. However, as new potential α-amylases from the halophilic archaea were identified, evident differences were observed with the sequences of their known hyperthermophilic counterparts. Unfortunately, most of those halophilic amylolytic enzymes were only putative proteins from genome sequencing projects [ 58 ], or their complete sequences were not available [ 26 , 44 , 46 ], making it difficult to establish accurate phylogenetic relationships. In addition, an enormous diversity was observed among the amylases characterized and sequenced from a member of the Halobacteriaceae family, which showed similarities with marine bacteria, fungal, or even animal sources [ 59 ]. Therefore, more insightful biochemical characterization studies are needed to reveal the exact features of these amylolytic enzymes from haloarchaea.

Although several copies of alpha-amylases appear in the sequenced genomes of haloarchaea, most studies are focused on the extracellular ones. The role of the extracellular amylase in haloarchaea is well-established, as it allows the conversion of starch, produced by the marine plankton, into simple sugars that could be incorporated into the cell and used as a carbon source [ 60 ]. Nonetheless, the function of the intracellular amylases is less understood, not only in haloarchaea but also in other heterotrophic microorganisms [ 61 ]. Most intracellular amylases of haloarchaea were assigned by sequence homology without a functional characterization, with few exceptions, like the intracellular α-amylase from H. japonica , whose activity is well-studied [ 49 ]. Further insight is necessary to complete the characterization of haloarchaeal amylases, to understand their role in archaeal metabolism, and to evaluate their biotechnological applications.

5. Conclusions

The detailed biochemical characterization of the cell-associated and the extracellular amylase activities from the new isolated strain Haloarcula sp. HS revealed that both are active at high salinity conditions and at considerably high temperatures. These features, joined to their stability under a wide range of surfactants, make them suitable for industrial applications. The proteomic analysis showed that three different cell-associated enzymes, one of which was also found in the extracellular medium, might be responsible for the amylase activities. The three proteins conserve the consensus domains and residues of the α-amylase family. Further studies aim to decipher the function of a hypothetical ancestral gene and to increase our understanding of the biochemical behavior of these polyextremophilic enzymes. Developing new techniques for high-scale production in the industry is also needed.

Acknowledgments

Technical support of Rocío Rodríguez from the IBVF-CSIC in the proteomic analysis is acknowledged.

Supplementary Materials

The following are available online at https://www.mdpi.com/article/10.3390/biology10040337/s1 . Figure S1: Full length of the 16S rRNA encoding gene from Haloarcula sp. HS. Figure S2: Complete original polyacrylamide gels. Figure S3: Predicted structures of the three amylase sequences identified in Haloarcula sp. HS. Figure S4: Multiple alignments of the amino acid sequence of the extracellular amylase identified in Haloarcula sp. HS (AMY_HS1). Figure S5: Multiple alignments of the amino acid sequence of the cell-associated amylase from Haloarcula sp. HS (AMY_HS2). Figure S6: Multiple alignments of the amino acid sequence of the cell-associated amylase from Haloarcula sp. HS (AMY_HS3).

Author Contributions

Conceptualization, J.V., R.L., and P.G.-V.; Funding acquisition, J.V., R.L. and P.G.-V.; Investigation, P.G.-V. and J.V.; Methodology, P.G.-V., B.G. and S.R.; Supervision, J.V. and R.L.; Software: P.G.-V. and L.R. Data curation, P.G.-V., J.V., L.R. and C.G.; Writing—original draft, P.G.-V. and R.L.; Writing—review & editing, P.G.-V., J.V. and R.L. All authors have read and agreed to the published version of the manuscript.

This research was funded by the Operative FEDER Program-Andalucía 2014–2020, the University of Huelva, the Spanish Agencia Estatal de Investigación (grants PID2019-109785GB-I00 and PID2019-110438RB-C22 -AEI/FEDER) and the SUBV. COOP.ALENTEJO-ALGARVE-ANDALUCIA 2019. P.G.-V. and C.G. acknowledge financial support from the University of Huelva (EPIT 2016-17) and the Junta de Andalucía (grant P18-RT-3154), respectively.

Institutional Review Board Statement

This study did not involve humans or animals.

Informed Consent Statement

Not applicable.

Data Availability Statement

Conflicts of interest.

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

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Enzymes - investigate the affect of amylase concentration on starch breakdown into glucose.

The aim of this investigation is to investigate the affect of amylase concentration on starch breakdown into glucose. To do this a couple of variable should be brought into consideration. They are the amount of amylase and the time taken for the starch to break down into glucose.

I predict that if I increase the concentration of amylase in the starch then the amount of time taken for the starch to break down into glucose will decrease, hence making it’s rate of reaction higher. This is because there will be more active sites for reactions, resulting in more chemical reactions caused by successful collisions between the active site of the amylase and the starch.

The higher the concentration of amylase in the starch then the amount of time taken for the starch to break down into glucose will decrease. This is because there is more amylase in a higher concentration gradient to break down the starch into glucose, thus making the reaction time much less. However there is less amylase in a lower concentration gradient that makes the reaction time longer because there is not enough amylase to break down he starch in one go.

Enzymes are biological catalysts/scissors. They speed up reactions within the body so that they can take place at body temperature. The enzyme is not used up in the reaction and can continue work on other substrate molecules. Enzymes are proteins with a special shape. All enzymes are made up of hundreds of amino acids joined together in a specific way. The protein (enzyme) molecule becomes twisted to form a special 3 dimensional shape, which can combine with a reactant (substrate) molecule, like a fitting in a particular key.

Enzymes combine with reactants, called the substrate. The enzyme allows the products to form the substrate by making and breaking chemical bonds easily.

An enzyme molecule can bring about hundreds of changes like this each second.

The enzyme is not used up at the end of the reaction and can continue to combine with other substrate molecules, forming more products. The enzyme is linked to a specific key, which fits onto a specific substrate (the lock). This is sometimes called the lock and key theory of enzyme action. These diagrams show two ways in which enzymes can bring about a change in the substrate: -

The special shape of a particular enzyme is held together by many different kinds of chemical bonds, it is these, which are affected by temperature (above 40°C) and incorrect pH.

Enzymes only one reaction in which the same end product is formed each time. They act at a certain optimum temperature, usually 36.7°C. Most enzymes are de-activated above 40°C. With most chemical reactions, raising the temperature will only increase the rate of reaction. This is true of enzyme-catalysed reactions, but only up to a point.

As you can see, initially as the temperature is increased so the rate of reaction goes up. You would expect this as the number of molecules possessing the amount of activation energy needed to react is being increased as more heat energy is supplied. The temperature coefficient, or Q 10 , is used to show the relationship between the reaction rate and a 10°C rise in temperature.

Q 10  = Rate of reaction at t°C + 10°C

       Rate of reaction at t°C

Calculate the value of Q10 for the reaction above if the temperature is raised from 25°C to 35°C.

For most enzyme reactions, Q 10  is about 2, i.e. for every 10°C rise in temperature, the reaction rate doubles. If the reaction above was kept at 35°C for a long period of time, and then Q 10  was calculated, it would appear to be a lot lower than 2. This is because some the enzymes have been de-natured.

As the reaction proceeds, the level of substrate would fall as it is converted into product by the enzyme. As the rate of reaction is also dependent on the concentration of substrate, this too would 'fall and has nothing to do with the effect of temperature. Hence the need for controls, i.e. keep all experimental factors constant except for the one being investigated.

This is a preview of the whole essay

After a certain point, the rate of reaction begins to fall drastically – at about 60°C, when you might think things should really be moving, there is no reaction at all. Why not? The answer lies in the fact that all enzymes are proteins. As with other proteins an enzyme has a very specific shape, and is held in this shape by strong disulphide bridges and much weaker hydrogen bonds, ionic interactions, etc. Heating the enzymes above a certain level will provide enough energy to break these weaker bonds and unfold the amino acid chain. When this happens the enzyme is said to be ‘denatured’. The enzyme is effectively destroyed, and usually will not work again even if the temperature falls.

The graph shows that there Is an optimum temperature for enzyme activity, i.e. warm enough to make the reaction proceed rapidly, but not hot enough to denature the enzyme. For most of our enzymes, this optimum lies in the range of 37-43°C. It Is no coincidence that the body temperature of humans is about 37°C. In fact, endothermic animals (those organisms such as birds and mammals that can maintain a body temperature Independent of that of their surroundings) have a number of specialised features to keep their temperature at the optimum point. Even ectotherms (those animals such as lizards whose body temperature fluctuates with that of their environment) make use of sun and shade to maintain their temperature within this range.

I predict that my graph will look like this: -

I predict that my graph will look like this because the rate of reaction will slowly increase as the concentration of amylase increases. I believe that when the concentration of the amylase will reach a certain gradient the rate of reaction will stay constant from there on. By this I mean that when the concentration of amylase reaches a maximum, for example 100%, the rate of reaction cannot increase any further due to the fact that the number of enzymes will be the same as the number of substrates. I carry on increasing the number of enzymes; only the number of enzymes equal to the number of substrates will react causing a maximum reaction rate. This is known as the V-max .

To explain my predicted graph further I have decided to split it up into sections: -

At point ‘ a ’, I predict that the rate of reaction will increase slowerr than proportional to the concentration of amylase. This is because the amylase would have to work quicker to break down all the starch. At point ‘ b ’, I predict that the rate of reaction will increase in proportional to the concentration of amylase. This is because the concentration of amylase would be at a point where there might be 5 starch molecules to every amylase molecule. At point ‘ c ’, I predict that the rate of reaction will increase quicker than proportional to the concentration of amylase. This is because at this point there might be two starch molecules to every amylase molecule, gradually becoming one starch molecule to one amylase molecule. At point ‘ d ’, I predict that the rate of reaction will stay the same even though the concentration of amylase still increases. This is because at this point there will be one starch molecule to one amylase molecule. Now if I carry on increasing the number of amylase molecules, the number of starch molecules will stay the same, and therefore the number of substrates needed to be broken down will also stay the same. This means that part ‘ d ’ of my predicted graph will not show on my graph of results, because it contains a concentration of more than 100% and my graph will only go up to 100%. Therefore I now predict that my graph will look like this: -

If there were 50 starch molecules and 5 amylase molecules then each amylase molecules would have to break down 10 starch molecules, this would take a long time. But if there were 50 starch molecules and 25 amylase molecules then the reaction would be 5 times quicker that 50 starch and 5 amylase. However if there were 50 starch molecules and 50 amylase molecules then the reaction would be at its quickest because each amylase molecule would have only to break down one starch molecule. Now if there were 50 starch molecules and 100 amylase molecules the reaction time would be as quick as 50 starch molecules and 50 amylase molecules. This is because on 50 amylase molecules are needed to break down the 50 starch molecules, meaning that the other 50-amylase molecules do nothing. This is called the V-Max .

I will change the concentration of amylase by diluting it with water. The variables that I need keep the same are: -

• Starch Concentration
• Temperature
• The number of drops of iodine
• Overall Variable

The independent variable  is the concentration of amylase and the dependent variable  is the amount of time taken for the starch to break down into glucose. To measure the concentration of amylase I will mix the amylase with the amount of water that will give me the correct concentration gradient I would need for that specific result. Then take 1 ml from that concentration and add it to the starch.

Reliability

To make my results more reliable I will repeat each concentration gradient 2 more times so that I have a wide range of results. From this I will calculate their averages and from the averages I will calculate the rate of reaction.

To make my results more accurate, I will measure them will extreme care and will be precise. I will make sure that I measure all the liquids at the lower meniscus and not at the upper meniscus. I will make sure that I start the stopwatch as soon as I pour all the substances into the test tube. Then I will stop the stopwatch as soon as the substances in the test tube turn colourless.

In order to conduct a safe experiment, I will follow all the lab rules of the classroom and will behave in a respective and mature manner. I will not do anything dangerous that would put any fellow classmates at risk. I will wear safety goggles and will clear my work area of any unnecessary equipment.

I will make this experiment safe by following the lab instructions of the classroom.

• Pour 5 ml of starch into the measuring cylinder.
• Use one Pipette to make sure that you have measured exactly 5 ml.
• Pour that into a test tube.
• Put the test tube into the test tube rack.
• Heat water in a kettle.
• Pour 100 ml of boiling water into a beaker to create a water bath.
• Put the thermometer in the water bath.
• Add a sufficient amount of cold water into the water bath so that the temperature of the water bath is at 40°C.

REMEMBER:  DO NOT FORGET TO MAINTAIN THE TEMPERATURE OF THE WATER BATH, THE TEMPERATURE MUST REMAIN AT 40°C AT ALL TIMES!

• Measure 5 ml of amylase into the small beaker.
• Use the other Pipette to make sure that you have measured exactly 5 ml.
• Use the Pipette to take 1 ml of amylase from that.
• Put the test tube into the water bath.
• Take 3 drops of iodine and add them into the test tube of starch at the same time as the amylase. [THE SOLUTION SHOULD TURN BLACK]
• Start timing immediately  using the stopwatch.
• Stop timing when the solution turns clear.
• Record your results in your results table.
• Repeat 1 – 16  two more times.
• Repeat 1 –17 , however this time use 4 ml of amylase and 1 ml of water, next time use 3 ml of amylase and 2 ml of water, now use 2 ml of amylase and 3 ml of water, finally use 1 ml of amylase and add 4 ml of water.
• Calculate the average time by adding up all your results and then dividing the answer by the number of results you have for that particular concentration gradient.
• Calculate the reaction rate by dividing 1 by the average time.
• After recording all your results tidy up the equipment.
• Finally draw a graph of your results. Label the y -axis rate of reaction  and label the x -axis concentration of amylase .

Empty Table of Results

I have decided to use these values on my table based on the results of the preliminary work that I carried out prior to this experiment. I will go up every 20% as I feel that is a sufficient gradient that will give me a reliable graph.

Empty Graph

I have decided to use these values on my graph based on the results of the preliminary work that I carried out prior to this experiment. My preliminary work consisted of short experiments know as ‘the spotting tile experiments’. I had to put drops of amylase and starch along with iodine solution onto a spotting tile. The amylase has to be at a certain temperature. I would then time how long the reaction took and then would record that onto a table of results. I would then divide 1 by the time to obtain the rate of reaction. I obtained 0.018 t -1  as the highest reading for the rate of reaction; hence this is the highest reading on my graph. The highest value on the x -axis, concentration of amylase is 100% as there is no value higher.

This is a diagram of my preliminary work: -

Table of Results

I will now draw another table in which I will show the rate of reaction and the points to be plotted on my graph.

Draw the graph yourself!

Both my graphs have show that my predictions were correct. I had predicted that the points on my graph would slowly increase until they reached a certain point where they would stay the same/stable this is called the V-Max . I had predicted that my graph would look like this:-

This has happened because the rate of reaction slowly increases as the concentration of amylase increases. For example if there are 50 starch molecules and 5 amylase molecules then each amylase molecules would have to break down 10 starch molecules, this would take a long time. But if there were 50 starch molecules and 25 amylase molecules then the reaction would be 5 times quicker that 50 starch and 5 amylase. However if there were 50 starch molecules and 50 amylase molecules then the reaction would be at its quickest because each amylase molecule would have only to break down one starch molecule. Now if there were 50 starch molecules and 100 amylase molecules the reaction time would be as quick as 50 starch molecules and 50 amylase molecules. This is because on 50 amylase molecules are needed to break down the 50 starch molecules, meaning that the other 50-amylase molecules do nothing. This is called the V-Max .

To explain the trend of my graph better, I will split it up into three sections: -

At point ‘ a ’, the rate of reaction is increasing slower than proportion to the concentration of amylase. This is because the amylase would have to work quicker to break down all the starch. At point ‘ b ’, the rate of reaction is increasing in proportion to the concentration of amylase. This is because the concentration of amylase would be at a point where there might be 5 starch molecules to every amylase molecule. At point ‘ c ’, the rate of reaction is increasing quicker than proportional to the concentration of amylase. This is because at this point there might be two starch molecules to every amylase molecule, gradually becoming one starch molecule to one amylase molecule. Hence my graph is a curve.

My results support my original predictions because I had predicted correctly using the scientific knowledge I have gained from various sources and also from the preliminary work I had done prior to this experiment. Evidence of my predictions being correct is shown in my graphs.

This experiment was a fair test because I only changed one variable and left all other variable the same where possible. For example I could not change the room temperature because I do not have control over it. I only changed the concentration of amylase.

To make sure that I measured accurately, I poured the liquids slowly and looked very carefully into the measuring cylinder to see where I was meant to stop pouring. When timing with the stopwatch I tried to stop the stopwatch as soon as I saw the solution go clear, however there area for human error here and what might look clear to me might not look clear to a classmate of mine. To see how much human error myself, could cause, I tested my reactions, the test consisted of starting and stopping a stopwatch, the lower down the time the quicker your reactions. On average I got 0.16 seconds, but the most I got was 0.07 seconds, which is quite quick. So you could estimate that I could be +  0.16 seconds on all the times I have measured because that was my average reaction time.

I believe that the procedures or method I used to carry out this experiment are not the most accurate procedures that could be used to carry out this investigation. Another method you could use would be the use of a computerised system, you could use a computer appliance to drop the iodine and concentration of amylase into the starch and start timing as soon as the iodine and amylase mix with the starch. The results that could be obtained from this experiment would be more reliable because there would be very little area for human error.

I believe that my results are quite reliable because I repeated the experiment twice to give me three sets of results from which I calculated averages. My results in the end followed the trend that I said would occur in my prediction, so this means that my prediction was correct. Most of my results were accurate, however some were not because they were anomalous results and did not follow the trend.

If I could do the experiment again, I would improve it by trying to cut out human error. By this I mean that would be more careful whist measuring and would also be quicker in starting the stopwatch. If I were to do the experiment again I would prefer to use a different method such as the method I suggested earlier, a computerised method rather than using this same method again because the computerised method is more accurate and has less of a margin of human error. However the method I used can produce very inaccurate results if I was not careful in measuring and also has a larger margin for human error.

Another alternative method is to use a colorimeter to measure the colour of the amylase, starch and iodine solution, to see when the solution has turned colourless. By using this method you are able to cut out all human error when it comes to measuring the colour of the solution. I could also use a syringe to measure the volume of the liquids before pouring them into the test tube instead of a measuring cylinder. A syringe has smaller scale divisions, of which the thickness of the line is even very small. A measuring cylinders division are large and the thickness of the line on it is quite thick. This would enable be to be more accurate.

Results Reliability

I believe that my results are reliable and sufficient because all the points are on a straight line, the tests covered enough range to show the full shape of the graph, as a result I am certain that I have got enough results to be sure about my conclusion as being correct, acceptable.

Here is a graph to show the reliability of my results: -

The error bars in red on the y -axis show the percentage error of 10 for the time taken for each concentration gradient.

Document Details

• Word Count 3709
• Page Count 13
• Level AS and A Level
• Subject Science

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Review on the role of polyphenols in preventing and treating type 2 diabetes: evidence from in vitro and in vivo studies.

1. Introduction

2. type 2 diabetes, 3. bioactive compounds, 3.1. phenolic compounds.

• Flavonoids: C 6 -C 3 -C 6 compounds characterized by two aromatic rings (A and B) connected to a central heterocyclic ring (C). Their pattern of hydroxyl and methyl substitutions define further classifications into flavonols (e.g., quercetin), flavones (e.g., luteolion), flavan-3-ols (e.g., catechin), flavanonols (e.g., dihydroquercetin), flavanones (e.g., naringenin), isoflavones (e.g., genistein), and anthocyanins (e.g., cyanidin). The latter subgroup comprises water-soluble pigments containing a sugar moiety. These pigments can be red, purple, or blue depending on the pH. Flavonoids are the phenolic class containing the highest number of compounds (over 8000 identified molecules).
• Hydroxycinnamic acids: Derivatives of cinnamic acid, with a C 6 -C 3 structure. Examples include caffeic, p -coumaric, ferulic, sinapic, and chlorogenic acids. Most phenolic acids found in nature belong to this subgroup.
• Hydroxybenzoic acids: Derivatives of benzoic acid, with a C 6 -C 1 structure, carrying at least one hydroxyl group. Examples include gallic acid and its dimeric form ellagic acid.
• Proanthocyanidins or condensed tannins: Tannins are oligomeric and polymeric phenolics that can be either condensed or hydrolysable. Condensed tannins (also known as proanthocyanidins) are formed by repeating units of catechin or epicatechin. Therefore, they can also be classified as flavan-3-ols, and include procyanidins ([epi]catechin polymers), prodelphinidins ([epi]gallocatechin polymers), and propelargonidins ([epi]afzelechin polymers).
• Hydrolysable tannins: Esters of gallic (gallotannins) and ellagic (ellagitannins) acids.
• Other phenolic groups include: lignans (derived from cinnamic acid derivatives), stilbenes (C 6 -C 2 -C 6 compounds with a 1,2-diphenylethylene functional group, in which resveratrol is the most representative compound), coumarins (C 9 H 6 O 2 compounds arranged in a bicyclic structure with lactone carbonyl groups), and hydroxytyrosol (a phenylethanoid derivative encountered in olive oil).

3.2. Antioxidant Function of Phenolic Compounds

3.3. bioefficiency of phenolic compounds, 4. polyphenols as antidiabetic agents, 4.1. in vitro evidence, 4.1.1. inhibition of α-amylase and α-glucosidase, 4.1.2. suppression of pro-inflammatory cytokines, 4.1.3. mitigation of ldl-cholesterol oxidative damage, 4.2. in vivo evidence, 4.2.1. effect of polyphenols on insulin sensitivity, 4.2.2. effect of polyphenols on dyslipidemia, 4.2.3. reduction of inflammatory processes, 5. conclusions, author contributions, data availability statement, conflicts of interest.

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Shahidi, F.; Danielski, R. Review on the Role of Polyphenols in Preventing and Treating Type 2 Diabetes: Evidence from In Vitro and In Vivo Studies. Nutrients 2024 , 16 , 3159. https://doi.org/10.3390/nu16183159

Shahidi F, Danielski R. Review on the Role of Polyphenols in Preventing and Treating Type 2 Diabetes: Evidence from In Vitro and In Vivo Studies. Nutrients . 2024; 16(18):3159. https://doi.org/10.3390/nu16183159

Shahidi, Fereidoon, and Renan Danielski. 2024. "Review on the Role of Polyphenols in Preventing and Treating Type 2 Diabetes: Evidence from In Vitro and In Vivo Studies" Nutrients 16, no. 18: 3159. https://doi.org/10.3390/nu16183159

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Eco-friendly synthesis and characterization of ZnO and Mg-Ag-doped ZnO nanoparticles using Phoenix dactylifera L. seeds: exploring biological activity and structural properties

• Original Article
• Published: 18 September 2024

Cite this article

• Dalal Doudi 1 , 2 ,
• Nasma Mahboub 1 , 2 ,
• Noura Gheraissa   ORCID: orcid.org/0000-0002-1377-4246 3 ,
• Ibtissam Laib   ORCID: orcid.org/0000-0002-9521-3003 1 ,
• Nezar Cherrada   ORCID: orcid.org/0000-0002-4153-6958 1 , 4 ,
• Ridha Messai 5 , 6 &
• Noureddine Slimani 7  

This study focuses on the green synthesis of zinc oxide (ZnO) and magnesium-silver-doped zinc oxide (Mg-Ag-doped ZnO) nanoparticles (NPs) via biomass conversion of Algerian Ghars date palm ( Phoenix dactylifera L.) seeds. Aqueous extracts of the seeds were utilized as reducing and stabilizing agents in the biogenic synthesis process. Structural, compositional, and morphological analyses, including X-ray diffraction (XRD), Fourier-transform infrared spectroscopy (FTIR), scanning electron microscopy-energy-dispersive X-ray analysis (SEM-EDAX), and ultraviolet–visible spectroscopy (UV–Vis), confirmed the successful formation of pure and Mg-Ag-doped ZnO NPs. The UV–Vis absorption spectra showed a shift from 395.6 nm (pure ZnO) to 373.2 nm (Mg-Ag doped), with corresponding energy values increasing from 3.13 to 3.32 eV, indicating changes in electronic structure due to doping. XRD analysis revealed an increase in average crystallite size from 12.8 nm (ZnO) to 22.0 nm (Mg-Ag ZnO) and a noticeable shift in peak positions, confirming successful doping. Biological evaluations demonstrated that Mg-Ag-doped ZnO NPs exhibited enhanced photocatalytic, antibacterial, antioxidant, and antidiabetic activities compared to undoped ZnO NPs. Notably, Mg-Ag ZnO NPs showed superior antioxidant activity with an IC 50 of 10.78 mg mL⁻ 1 and EC 50 of 0.79 mg mL⁻ 1 , compared to ZnO NPs with an IC 50 of 11.51 mg mL⁻ 1 and EC 50 of 0.84 mg mL⁻ 1 . They also exhibited higher photocatalytic degradation efficiency of methylene blue dye (93% vs. 87% for ZnO) under UV light. Antibacterial studies showed that Mg-Ag ZnO NPs had lower minimum inhibitory concentrations (MIC) and minimum bactericidal concentrations (MBC) than pure ZnO NPs, with a MIC of 0.625 mg mL⁻ 1 and MBC of 0.625 mg mL⁻ 1 for E. coli , compared to 2.5 and 10 mg mL⁻ 1 , respectively, for pure ZnO. Furthermore, Mg-Ag-doped ZnO NPs exhibited significant α-amylase inhibition (48.0% at 0.25 mg mL⁻ 1 ), outperforming pure ZnO NPs (38.9% at the same concentration), and showed competitive inhibition to the reference drug acarbose in antidiabetic tests. These findings highlight the potential of rationally designed biogenic ZnO nanostructures synthesized through biomass conversion of P. dactylifera seeds, especially after strategic doping, for various biomedical and environmental applications. This green synthesis approach, utilizing renewable biomass, offers an eco-friendly and sustainable route for producing ZnO-based nanomaterials with tunable properties.

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Department of Molecular and Cellular Biology, Faculty of Natural and Life Science, University of El Oued, 39000, El Oued, Algeria

Dalal Doudi, Nasma Mahboub, Ibtissam Laib & Nezar Cherrada

Laboratory Biology, Environment, and Health, University of El Oued, 39000, El Oued,, Algeria

Dalal Doudi & Nasma Mahboub

Department of Biology, Faculty of Natural and Life Science, University of El Oued, 39000, El Oued, Algeria

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Laboratory of Biodiversity and Application of Biotechnology in Agriculture, University of El Oued, 39000, El Oued, Algeria

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Laboratory of Sciences and Techniques of the Environment and Valorization, Abdelhamid Benbbadis, Mostaganem University, 27000, Mostaganem, Algeria

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Doudi, D., Mahboub, N., Gheraissa, N. et al. Eco-friendly synthesis and characterization of ZnO and Mg-Ag-doped ZnO nanoparticles using Phoenix dactylifera L. seeds: exploring biological activity and structural properties. Biomass Conv. Bioref. (2024). https://doi.org/10.1007/s13399-024-06115-x

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Received : 18 June 2024

Revised : 14 August 2024

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Published : 18 September 2024

DOI : https://doi.org/10.1007/s13399-024-06115-x

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