Enzymes: Catalysts of Life
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The following article was originally published in the journal for educators Chemia w Szkole (eng. Chemistry in School) (3/2016):
Enzymes are large biomolecules, most commonly proteins, that act as highly specific biological catalysts. They significantly accelerate a wide variety of chemical reactions that together form the biochemical basis of life.
Reactions involving enzymes typically proceed much faster than those occurring without catalytic assistance. For instance, carbonic anhydrase, one of the fastest known enzymes, can catalyze the conversion of carbon dioxide CO2 and water H2O into bicarbonate ions HCO3– at a rate of about 106 reactions per second. This represents an acceleration of nearly 107 times compared to the uncatalyzed reaction [1] [2].
Although the digestive effect of gastric secretions on proteins had been observed outside the living body as early as the 18th century [3], it was not until the 19th and early 20th centuries that researchers such as Louis Pasteur, Wilhelm Kühne, and Eduard Buchner fully elucidated the nature of enzyme activity. Buchner’s work with yeast-derived enzymes earned him the Nobel Prize in 1907 [4].
Enzymes are categorized into six [currently seven, author's note: 2025] primary classes according to the types of reactions they catalyze. Each class is designated by a unique Enzyme Commission (EC) number [5]:
- Oxidoreductases (EC 1) – catalyze oxidation-reduction reactions involving a variety of substrates,
- Transferases (EC 2) – catalyze the transfer of functional groups between molecules,
- Hydrolases (EC 3) – catalyze the cleavage of bonds by the addition of water,
- Lyases (EC 4) – catalyze the breaking of chemical bonds by mechanisms other than hydrolysis or oxidation,
- Isomerases (EC 5) – catalyze structural rearrangements within a molecule,
- Ligases (EC 6) – catalyze the joining of two molecules by forming new covalent bonds, typically coupled with ATP hydrolysis.
- Translocases (EC 7) – catalyse the movement of ions or molecules across membranes or their separation within membranes [added as a class in 2018, author's note: 2025].
From a biological perspective, the role of enzymes is truly fundamental. By participating in nearly all anabolic and catabolic reactions, they shape the overall architecture of metabolism. Enzymes direct the flow of metabolic pathways by selectively catalyzing specific chemical transformations. In doing so, they influence how molecules are processed, how energy is managed, and how essential cellular functions are carried out.
At first glance, it may seem that working with highly specialized substances like enzymes is limited to advanced research laboratories. However, this couldn't be further from the truth. In fact, the presence of various enzymes in biological material can be detected using surprisingly simple techniques. These accessible methods can be both educational and deeply rewarding for anyone curious about exploring the world of biochemistry.
Ureases
Ureases are enzymes from the hydrolase class that catalyze the hydrolysis of urea CO(NH2)2, producing ammonia NH3 and carbon dioxide CO2.
In the active site of naturally occurring ureases, the metal ion present is typically nickel. However, laboratory studies have shown that catalytic activity can also be achieved when this metal is substituted with manganese or cobalt [6]. The molecular mass of the active form of the enzyme is approximately 500 kDa.
This enzyme is found in yeasts, various bacteria (such as Helicobacter pylori), and in certain higher plants, including the jack bean Canavalia ensiformis, a member of the Fabaceae (legume) family. It was from this plant that urease was first isolated in 1926 by James B. Sumner, who went on to demonstrate that the enzyme is a protein [7].
For the purposes of our experiments, convenient sources of urease include readily available seeds from vegetable soybean Glycine max, field pumpkin Cucurbita pepo, or giant pumpkin Cucurbita maxima. The seeds may be dried, but it is essential that they have not been exposed to high temperatures during processing.
Place a small amount of seeds rich in urease, such as soybeans, into a mortar (Photo 1A). Although the seeds are quite firm, grinding them produces a yellowish powder after a short time (Photo 1B). Suspend this powder in 20 to 30 cm3 (0.7 to 1.0 fl oz) of water at room temperature and filter the mixture. It is perfectly acceptable if the resulting extract remains slightly cloudy (Photo 2). The solution can be stored in a refrigerator for several days. Just before conducting the experiment, divide the extract into two equal portions. Leave one portion unchanged, and bring the other to a boil for a few minutes, then allow it to cool to room temperature.
To test for urease activity, prepare an aqueous solution of urea at a concentration of approximately 8%, and add a few drops of an alcoholic solution of bromothymol blue C27H28Br2O5S. This pH indicator turns yellow in acidic environments (pH below 7), blue in alkaline conditions (pH above 7), and green when the pH is close to neutral (pH ≈ 7).
The resulting solution should have a pH close to neutral or slightly acidic. If it appears blue, which indicates an alkaline environment, this can be corrected by adding a small amount of diluted acid such as acetic acid CH3COOH.
Distribute the prepared urea solution into three small beakers. The first (Photo 3A) will serve as the control. To the second, add a few cubic centimeters (about 0.1–0.2 fl oz) of the raw soybean extract (Photo 3B), and to the third, add the extract that was previously boiled (Photo 3C). Leave the beakers undisturbed for a few minutes.
After just a few minutes, a noticeable color change occurs as the solution turns blue. However, this happens only in the sample containing the raw soybean seed extract (Photo 3E). This change results from urease catalyzing the reaction described by the following equation:
The ammonia produced further undergoes hydrolysis in an aqueous environment according to reaction:
The reaction environment becomes alkaline, as evidenced by the visible color change of the solution containing the pH indicator.
If the reaction is allowed to proceed longer, a characteristic pungent odor of ammonia becomes distinctly noticeable.
The crucial role of urease is highlighted by the absence of any reaction in the control sample. Clearly, urease significantly accelerates the hydrolysis of urea compared to the uncatalyzed reaction. In fact, even after weeks of waiting, no signs of reaction would appear in the control sample.
The lack of reaction in the third sample suggests that the elevated temperature used to treat the soybean extract in this case inactivated the enzyme. Furthermore, this loss of urease activity is irreversible, as cooling the boiled extract back to room temperature does not restore its function.
It is important to note that urease can act as a potent toxin under certain conditions. However, this would require direct introduction into the bloodstream. Through the hydrolysis of urea, which is present in small amounts in blood, toxic ammonia would be generated. Fortunately, consuming foods containing urease poses no risk, as the enzyme is simply digested like other proteins.
Peroxidases
Another group of intriguing enzymes are peroxidases, classified within the oxidoreductase family. These enzymes catalyze the oxidation of various substrates by hydrogen peroxide H2O2, following the general scheme (X – substrate, XO – oxidation product):
Peroxidases are found in both animal and plant tissues. A relatively high concentration of these enzymes is present in the root of common horseradish Armoracia rusticana (Photo 4), a widely distributed plant from the Brassicaceae family commonly used as a spice.
Horseradish peroxidase (HRP), isolated from the root of horseradish, is a glycoprotein, meaning a protein covalently linked with oligosaccharides. Its molecular weight is approximately 44 kDa. The enzyme’s coenzyme component is a heme group [9].
The peroxidase activity in horseradish root can be demonstrated through several methods. One particularly striking approach involves the enzymatic oxidation of luminol. To perform this, prepare an alkaline chemiluminescent solution by dissolving 0.3 g of sodium carbonate Na2CO3 and 0.05 g of luminol in 100 cm3 (approximately 3.4 fl oz) of distilled water. If sodium carbonate is unavailable, alkaline conditions can alternatively be achieved by adding a small amount of ammonium hydroxide NH3(aq) or sodium hydroxide NaOH.
Immediately before the experiment, add 5 cm3 (approximately 0.17 fl oz) of 3% pharmaceutical-grade hydrogen peroxide to the solution. Then, introduce a piece of horseradish root, preferably cut from its interior (Photo 5). If horseradish is unavailable, common parsley root Petroselinum crispum, which also contains the enzyme, can serve as a substitute.
After darkening the room, the effect shown in Photo 6 becomes visible. The solution, upon contact with the plant tissue, emits a distinctly visible light characteristic of the luminol oxidation reaction under alkaline conditions.
The experiment can also be performed by applying the solution to a cross-section of the root using a Pasteur pipette (Photo 7A). In this case, the luminescence appears uneven, with some areas of the cross-sectional surface glowing noticeably less than others. While part of this variation may result from mechanical damage during sample preparation, it primarily reflects differences in peroxidase content.
Luminol chemiluminescence is known to be triggered by various activators, such as iron(III), cobalt(II), and copper(II) complex compounds soluble in alkaline environments [10]. However, in this case, the reaction is catalyzed specifically by horseradish peroxidase present in the root, resulting in the emission of blue light.
Because the chemiluminescent reaction of luminol is catalyzed, among others, by hemoglobin, luminol is widely used in forensic science to detect the presence of blood, for example, at crime scenes. Blood stains exposed to luminol emit a distinct glow, which can be observed with the naked eye or captured photographically. However, a challenge arises from the widespread occurrence of peroxidases, which, as demonstrated earlier, also catalyze this reaction. The same issue applies to other methods, such as the catalytic oxidation of leuco dyes, for instance the conversion of colorless leucomalachite green to its colored form in alkaline conditions or the oxidation of non-fluorescent leucofluorescein to fluorescent fluorescein under UV light [11]. One possible way to address this problem is based on the fact that peroxidases, like most enzymes, are highly sensitive to elevated temperatures. Briefly heating the sample to an appropriate temperature effectively abolishes peroxidase activity, so a positive result after such treatment increases confidence in the presence of hemoglobin and thus blood.
The activity of peroxidase can also be demonstrated by its catalytic oxidation of colorless benzidine to colored products using hydrogen peroxide, or by the oxidation of potassium iodide KI to free iodine, which can be easily detected using a starch suspension [12].
Proteases
Proteases, also known as proteolytic enzymes, belong to the hydrolase class and catalyze proteolysis, which is the hydrolysis of peptide bonds. A peptide bond is the amide linkage formed between amino acids that make up proteins and peptides.
Proteases are sometimes referred to as peptidases and can be classified into two main groups:
- exopeptidases – which cleave single amino acids from the ends of peptide or protein chains,
- endopeptidases – which hydrolyze peptide bonds within the interior of the chain.
Proteases are found in both animals and plants. In animals, they play crucial roles such as serving as digestive enzymes, while in plants they often fulfill protective functions [13].
Proteases are present in numerous plants. Notable examples include bromelain found in the fruit of the edible pineapple Ananas comosus, actinidin present in the fruits of kiwifruit species such as Actinidia deliciosa and Actinidia chinensis, as well as papain found in the unripe fruit of the papaya tree Carica papaya.
To empirically demonstrate protease activity, the easiest approach is to use pineapple or kiwi fruit. It is important to use only fresh or frozen fruit, as canned fruit is unsuitable due to heat treatment applied during processing.
I used a pineapple fruit (Photo 8) for my experiment. Only small amounts of the material are required; the remainder can be consumed to benefit the body’s nutrition.
An extract from pineapple fruit can be easily prepared by mashing a small amount of pulp with a little water, followed by filtering the resulting mixture. The liquid obtained displays a yellow coloration (Photo 9) and can be stored under refrigeration for several days.
But how can one demonstrate that bromelains, the protein-degrading enzymes, are truly present in pineapple fruit? A common method involves using gelatin, a mixture of partially hydrolyzed collagen products.
A – gelatin solution in water (control sample), B – gelatin solution in water with pineapple extract (test sample), C – gel formed in the control sample, D – no gel formation in the test sample
Start by preparing a gelatin solution by dissolving 10 g of gelatin in 100 cm³ (approximately 3.4 fl oz) of hot water. Once the solution cools slightly, but before it begins to set, add a small amount of water to one sample (control, Photo 10A) and an equal amount of pineapple extract to the other (test, Photo 10B). Place both samples in a cool location and let them sit for several hours.
After some time, you’ll notice that the control sample solidifies into a firm gel (Photo 10C). In contrast, the sample with pineapple extract remains completely liquid (Photo 10D). Even after a long period, it will not gel. How can this be explained?
As we know, at any temperature above absolute zero, all particles are in motion, with their velocity increasing as the temperature rises. In a hot gelatin solution, long protein chains undergo vigorous, random movements. As the temperature decreases, these movements slow down, and hydrogen bonds begin to form between different regions of the same chain or between separate chains. Although hydrogen bonds are relatively weak, their large number at lower temperatures allows them to play a dominant role in aggregating the long polypeptide chains. Water molecules become trapped within this emerging network, leading to the formation of a gel.
When proteases such as bromelain, which is derived from pineapple, are present in the solution, they catalyze the hydrolytic cleavage of peptide bonds. This results in free amino acids and short peptide fragments that are too small to aggregate and form a gel.
Agar, also known as agar-agar, is a substance with properties similar to gelatin. It is extracted from marine red algae belonging to the Rhodophyta phylum. Unlike gelatin, which consists of proteins and peptides, the primary component of agar is agarose, a polysaccharide composed of galactose derivatives C6H12O6. As a result, agar-based systems are capable of forming a gel even in the presence of high concentrations of proteases (Photo 11). Proteases do not catalyze the breakdown of sugars, so they do not interfere with the solidification of agar solutions.
From this experiment, it becomes clear that it is not possible to prepare a gelatin-based jelly flavored with natural pineapple, kiwi, or papaya juice. The reason lies in the proteolytic enzymes naturally present in these fruits, which prevent gelation. Although applying sufficiently high temperatures deactivates the enzymes, it also alters the flavor, which may be undesirable. Nevertheless, such desserts can still be successfully prepared using agar as the gelling agent.
Amylases
Amylases, also known as amylolytic enzymes, belong to the hydrolase class, much like proteases and ureases. However, instead of catalyzing the breakdown of proteins or urea, they facilitate the hydrolysis of starch and other polysaccharides [14]. In animals, amylases are found in saliva and pancreatic juice. In plants, they are present in various tissues, including fruits and germinating seeds.
Given the availability of raw materials, we will attempt to confirm the amylolytic activity of salivary amylase, formerly known as ptyalin.
To prepare the starting solution, dissolve several grams of potato starch in 100 cm³ (approximately 3.4 fl oz) of hot water, then filter and allow it to cool.
Fill three test tubes with the starch solution. The first tube, without any additives, will serve as the control sample (Photo 12A). To the second tube, add a small amount of saliva diluted fivefold with distilled water (Photo 12B), while to the third, add a similar amount of saliva solution that has been briefly boiled (Photo 12C). The liquid in all tubes is colorless. Leave the solutions at room temperature for about one to two hours. Alternatively, if incubated at approximately 37°C (98.6°F), only a few minutes are needed.
After this period, examine the contents of the tubes. All three solutions appear unchanged, remaining colorless (Photo 12D, E, F). To verify the presence of starch, add a few drops of commercially available iodine solution, either alcoholic or aqueous (with potassium iodide KI), to each tube.
The result may come as a surprise since the characteristic deep blue coloration of the iodine-starch complex appears only in the solution without amylase (Photo 12G) and in the solution containing amylase that was previously heated to boiling temperature (Photo 12I), which inactivated the enzyme. In the solution with active amylase, no blue coloration was observed; instead, the solution remained brown due to dissolved iodine. This clearly indicates that the enzyme effectively broke down the long-chain polysaccharides in starch into shorter fragments that do not form the blue iodine complex.
Salivary amylase hydrolyzes starch into maltose and dextrins, representing an initial stage of polysaccharide digestion that begins in the oral cavity.
Explanation
As you can see, enzymes are found virtually everywhere, both around us and within our bodies, and they are surprisingly easy to detect. You can readily observe their properties firsthand.
Enzymes are primarily protein chains of varying lengths, ranging from just a few dozen amino acids (e.g., 4-oxalocrotonate tautomerase) to over 2,500 amino acids in a single chain (such as animal fatty acid synthase) [15] [16]. The region of the chain that binds and interacts with the substrate, containing amino acids essential for catalysis, is known as the enzyme's active site. In addition to substrates, enzymes can also bind other molecules, such as cofactors, which modulate their catalytic activity.
In living cells, enzymes are synthesized by ribosomes as linear chains of amino acids, which then fold into specific three-dimensional structures. An enzyme’s function is determined by this spatial conformation. Elevated temperatures can cause proteins to denature, resulting in structural alterations that typically eliminate catalytic activity. This effect was demonstrated in the enzyme experiments discussed earlier.
The effect of temperature within the non-denaturing range is also easy to observe. For example, in the case of amylase, reaction rates increase with temperature. Most enzymes operate with optimal efficiency within a narrow temperature and pH range.
Another defining characteristic of enzymes is their high substrate specificity, often far exceeding that of inorganic catalysts. For instance, proteases from pineapple specifically hydrolyzed the proteins in gelatin but had no effect on agar. This may seem surprising, since both proteins and polysaccharides are composed of long polymer chains. Enzymes, however, often exhibit even more refined specificity for their substrates. The mechanisms of enzymatic action are frequently explained using models such as the lock-and-key, three-point interaction, or induced fit theories.
Enzymology is a rapidly evolving discipline that explores the occurrence, structure, function, mechanisms of action, and practical applications of enzymes.
Enzyme activity can also be influenced by additional factors, such as the presence of metal ions. There are many fascinating experiments to explore in this area, and I encourage readers to try them for themselves.
References:
- [1] Lindskog S., Structure and mechanism of carbonic anhydrase, Pharmacology & therapeutics, 1 (74), 1997, pp. 1–20 back
- [2] Stryer L., Biochemia, Wyd. 6, Wydawnictwo Naukowe PWN, Warszawa, 2009 back
- [3] de Réaumur R. A. F., Observations sur la digestion des oiseaux, Histoire de l’academie royale des sciences, 461, 1752, p. 266 back
- [4] Buchner E., Cell-free fermentation, Nobel Lectures, Chemistry 1901-1921, Elsevier Publishing Company, Amsterdam, 1966 back
- [5] Moss G. P., Recommendations of the Nomenclature Committee, w: International Union of Biochemistry and Molecular Biology on the Nomenclature and Classification of Enzymes by the Reactions they Catalyse, dostępne online: http://www.chem.qmul.ac.uk/iubmb/enzyme/ [dostęp: 23.04.2016] back
- [6] Carter E. L., Flugga N., Boer J. L., Mulrooney S. B., Hausinger R. P., Interplay of metal ions and urease, Metallomics, 1 (3), 2009, pp. 207–221 back
- [7] Karplus P. A., Pearson M. A., Hausinger R. P., 70 years of crystalline urease: What have we learned?, Accounts of Chemical Research, 30 (8), 1997, pp. 330–337 back
- [8] Pluciński T., Doświadczenia chemiczne, Wydawnictwo Adamantan, Warszawa, 1997, pp. 97 back
- [9] Veitch N. C., Horseradish peroxidase: a modern view of a classic enzyme, Phytochemistry, 65 (3), 2004, pp. 249–259 back
- [10] Ples M., Widmowy blask - chemiluminescencja katalizowana związkiem miedzi (eng. Ghostly glow: copper-catalyzed chemiluminescence), Chemia w Szkole (eng. Chemistry in School), 2 (2016), Agencja AS Józef Szewczyk, pp. 13-17 back
- [11] Ples M., Na tropie - fluorescencyjne wykrywanie śladów krwi (eng. What the Eye Can’t See – Tracking Blood with Chemiluminescence), Chemia w Szkole (eng. Chemistry in School), 1 (2015), Agencja AS Józef Szewczyk, pp. 25-26 back
- [12] Pluciński T., Doświadczenia chemiczne, Wydawnictwo Adamantan, Warszawa, 1997, p. 92 back
- [13] Hattori M., Hirayama C., Konno K., Nakamura M. et al, Papain protects papaya trees from herbivorous insects: role of cysteine proteases in latex, The Plant Journal, 37 (3), 2004, pp. 370-378 back
- [14] Konturek S., Fizjologia układu trawiennego, Państwowe Zakłady Wydawnictw Lekarskich, Warszawa, 1985 back
- [15] Chen L. H., Kenyon G. L., Curtin F., Harayama S., Bembenek M. E., Hajipour G., Whitman C. P., 4-Oxalocrotonate tautomerase, an enzyme composed of 62 amino acid residues per monomer, The Journal of Biological Chemistry, 25 (267), 1992, pp. 17716–17721 back
- [16] Smith S., The animal fatty acid synthase: one gene, one polypeptide, seven enzymes, The FASEB Journal, 15 (8), 1995, pp. 1248–1259 back
All photographs and illustrations were created by the author.
Addendum
Here is a video showing the experiment with horseradish peroxidase described above.
Marek Ples