The Reaction Will Only Continue as Long as the Substrate is Present

Substrate Concentration

It is precisely thermostated (typically at 37°C) before passing through a flow cell (0.6 ml in Miles et al., 1985), in which heat evolution is detected via thermopiles as a potential difference and amplified.

From: Molecular and Diagnostic Procedures in Mycoplasmology , 1995

Environmental Biotechnology and Safety

M. Reis , ... M. Majone , in Comprehensive Biotechnology (Second Edition), 2011

6.51.3.2.4 Influent substrate concentration

Influent substrate concentration affects the kinetics of substrate consumption and polymer storage. Substrate uptake rates and polymer production rates can be expected to increase with increasing substrate concentrations up to a maximum value (see example in Figure 6 (a)), decreasing again at higher substrate concentrations due to inhibition by the substrate (not shown in the figure). Inhibition by substrate has been demonstrated by several authors, particularly in the batch production stage [15, 33, 36]. This effect can be determinant for the effectiveness of the selection stage as well, because substrate inhibition results in lengthening of the feast phase (for a given cycle length) and, therefore, affects the F/F ratio.

Figure 6. Substrate uptake rate dependence on carbon substrate concentration: (a) depicts the experimentally determined specific substrate uptake rate, q _S, as a function of the substrate concentration – this graph was constructed using data obtained in three tests conducted using a culture selected at 45   Cmmol   VFA   l⁻¹ subjected to 30, 45, and 60   Cmmol   VFA   l⁻¹ of influent substrate concentration [30] – and respective adjusted Monod function; (b) shows simulated curves generated using the Monod function determined in (a) describing the substrate uptake rate dependence on substrate concentration during the feast phases of SBRs operated at influent substrate concentrations of 30, 45, and 60   Cmmol   VFA   l⁻¹ (1.1, 1.6, and 2.1   g-COD   l⁻¹, respectively) [30] compared to that obtained in a continuous system operated at an influent substrate concentration of 120   Cmmol   VFA   l⁻¹ (4.2   g-COD   l⁻¹) and with a residual feast reactor concentration of 25   Cmmol   VFA   l⁻¹ (0.9   g-COD   l⁻¹) [35].

On the other hand, the use of low influent substrate concentrations can also result in the selection of cultures with poor PHA storage performance due to the rate-limiting concentration of carbon substrate. As the competitive advantage of PHA-producing organisms mostly consists in that they take up the substrate at a higher rate than competing organisms, this advantage may be lost if the substrate uptake rate is limited by the influent substrate concentration [30, 35].

Because in continuous feast reactors (see Section 6.51.3.2.3) a constant residual substrate concentration is imposed, it can be suggested that as long as the feast reactor residual substrate concentration is kept above a kinetically limiting value, continuous systems present an advantage in terms of PHA culture selection efficiency. In these systems, the substrate concentration is constant and, therefore, a constant pressure for PHA-storing organisms is also maintained ( Figure 6 (b)), while in SBRs, as substrate concentration declines to reach carbon-limiting concentrations, so will the substrate uptake rate and with it the pressure for PHA-storing organisms can also decline ( Figure 6 (a)). On the other hand, the SBR is more flexible and the influent substrate concentration can, in principle, be controlled by adjusting the length of the feed phase.

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Structured acylglycerides emulsifiers with bioactive fatty acids as food ingredients

Alaina Alessa Esperón-Rojas , ... Hugo Sergio García , in Value-Addition in Food Products and Processing Through Enzyme Technology, 2022

2.4.2 The molar ratio of substrates

Substrate concentration is a key aspect; an increase in esterification and transesterification has been reported by increasing the concentration of FFA. However, when the concentration of FFA increases, changes in the polarity and viscosity of the reaction medium occur. In the case of the esterification of CLA to PC, its incorporation increased to 85.8% by increasing the molar ratio of substrates (MRS) (PC:CLA) from 1:2 to 1:4 ( Baeza-Jiménez et al., 2012).

For the incorporation of CA into PC, 49% of incorporation was reached with 1:6 (PC:CA) ratio. While 43% and 35% incorporation of n-3 PUFAs in PC was obtained when the MRS (PC:n-3 PUFAs) was 1:8 (Kim et al., 2010).

In the synthesis of acylglycerols by esterification of MCFAs to glycerol, the effect of the variation of MRS was evidenced both in the performance of the products and in the preference on one of the chemical species formed (MAG   >   DAG   >   TAG), so it is another important controlling variable to obtain products of interest (Esperón-Rojas et al., 2017).

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Kinetics of Microbial Processes☆

N.S. Panikov , in Reference Module in Earth Systems and Environmental Sciences, 2016

Concentration of Limiting Substrate

Substrate concentration in water or soil, s, stands for amount of some essential nutrient used by microorganisms for growth and maintenance. Normally we cannot assess all potentially available nutrients and focus on one or few individual compounds or class of molecules representing the limiting nutrient substrate, i.e. such nutrient(s) that control growth and activity of microorganisms in question. Typically, growth of chemoorganotrophic microorganisms in soil, subsoil and aquatic habitats is limited by available organic compounds, while photosynthetic microbiota is limited either by light or sources of P, N or Fe.

The absolute rate of substrate consumption by microorganisms depends on its concentration in environment and is measured from the rate of s decline over time (ds/dt), while the specific rate of substrate consumption, q is calculated by normalizing the absolute rate to the instant cell mass concentration, x:

(3) $q=-\frac{1}{x}\times \frac{ds}{dt}=Q\frac{s}{K_s+s}$

Eq. (3) is similar to the Michaelis–Menten equation established in enzymology with parameters Q standing for maximum uptake rate and saturation constant K_s numerically equal to such substrate concentration at which uptake rate q  =   0.5 Q. Specific growth rate of microorganisms, μ is related to q by a mass balance Eq. (4) (see below) and therefore can be derived from [3] as follows:

(3a) $\mu =Yq={\mu }_m\frac{s}{K_s+s}\text{,}$

where μ   =   YQ is the maximal specific growth rate. This expression is known as Monod equation employed by the simplest models of microbial growth. Note that both Eqs. (3) and (3a) are empirical and should be applied with caution to experimental data. The substrate uptake rate depends not only on its concentration but also on other environmental factors (temperature, tonicity, pH, etc.) and on the state of cells, what particular transporters are expressed and take part in nutrients consumption. Even more complex and variable is intracellular machinery responsible for growth (see below the section 'Structured models'). Therefore Eqs. (3) and (3a) could be applied only to relatively simple experiments with s as the only independent variable and under conditions that studied cells preserve the same physiological state (no evident changes in gene expression profile or spontaneous mutations or changes in composition of microbial community) at all tested s.

There are two groups of microbial nutrient substrates: 1) catabolic substrates which are sources of energy, and 2) anabolic or conserved substrates which are sources of biogenic elements forming cellular material. Examples of catabolic substrates are H₂ for hydrogen-oxidizing microorganisms, NH₄ ⁺ and NO₂ ⁻ for nitrifying bacteria, S⁰ for sulfur-oxidizing bacteria, and oxidizable or fermentable organic substances for diverse heterotrophic species. Their consumption is accompanied by oxidation and dissipation of chemical substances into waste products, which are no longer reusable as an energy source (H₂O, NO₃ ⁻, SO₄ ^{2   −}, CO₂, etc.). The anabolic substrates are consumed and incorporated into de novo synthesized cell components, being conserved in biomass; thus, sometimes anabolic substrates are referred to as conserved substrates. Contrary to catabolic substrates, the conserved substrates can be reabsorbed after excretion or cell lysis. They include nearly all non-carbon sources of biogenic elements (N, P, K, Mg, Fe, and trace elements), CO₂ for autotrophs, as well as amino acids and growth factors for respective auxotrophic species.

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ENZYMES | Enzyme-Based Assays

K. Matsumoto , H. Ukeda , in Encyclopedia of Analytical Science (Second Edition), 2005

One-Step Reaction

A substrate S is to be determined. S shall be completely converted into the product P in an enzymatic reaction. If S has a characteristic light absorption in the ultraviolet or visible range different from P, S can be directly determined even in the presence of other absorbing substances in the spectrophotometer cuvette. The absorbance decreases by an amount corresponding to the quantity of S converted. Various measuring techniques can be used as well as absorption photometry, such as fluorimetry, luminometry, calorimetry, and potentiometry. If dissolved oxygen is consumed by the reaction, especially such as oxidase reaction, the oxygen consumed is monitored by a Clark-type oxygen electrode amperometrically. This measuring technique is advantageous for colored or turbid sample solution.

The most widely used enzymatic reactions are those with NAD(P)-dependent dehydrogenases. By monitoring the absorbance at 340 nm, the enzymatic conversion of the substrate can be followed directly in the photometer cuvette without influencing the chemical process. The substrate concentration can be calculated using the molar extinction ( ε ₃₄₀=6.3×10³  l   mol⁻¹  cm⁻¹) of NADH.

All enzymatic reactions involving co-enzymes are two-step reactions. However, if one of the substrates is present in a very high concentration in relation to its Michaelis constant, two-substrate reactions can be treated kinetically as one-substrate reactions. These conditions are also desirable for endpoint determinations with co-enzymes, simply to achieve a high reaction rate. However, if NAD(P)H is the second substrate, the degree to which its concentration can be increased is limited by its high absorbance. On the basis of experience, relatively large quantities of enzyme are used with relatively little substrate, so that the reaction proceeds rapidly to completion. The absorbance should thus be easily readable (neither too low nor too high).

Generally, enzyme reactions are equilibrium reactions. If the conversion is incomplete because of an unfavorable equilibrium, a determination is only possible with shifting the equilibrium by special experimental techniques. The equilibrium can be shifted by the increase of substrate concentration, variation of pH, and by trapping agents. The equilibrium constant may be altered by several orders of magnitude. The use of trapping agents, for example, semicarbazide and hydrazine when ketones and aldehydes are reaction product, is convenient for achieving the purpose. However, this technique is not always useful. Trapping agents can inhibit the catalyzing enzyme, as in the case of 3-hydroxybutyrate dehydrogenase (EC 1.1.1.30). Semicarbazide and hydrazine react slowly with NAD to form a compound that absorbs in the longwave ultraviolet and overlaps the absorption spectrum of NADH. The more elegant way is to use a trapping enzyme reaction, whose equilibrium is far in favor of the reaction products. Total substrate conversion can be achieved by coupling such a trapping reaction with the primary reaction. Moreover, if the trapping reaction itself is catalyzed by an NAD(P)-dependent enzyme, in many cases the measuring signal is doubled, as in the case of the determination of ethanol using alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (AlDH):

$\text{Ethanol}+{\text{NAD}}^{+}\stackrel{\text{ADH}}{\to }\text{acetaldehyde}+\text{NADH}+{\text{H}}^{+}$

$\text{Acetaldehyde}+{\text{NAD}}^{+}\stackrel{\text{AIDH}}{\to }\text{acetic}\text{acid}+\text{NADH}+{\text{H}}^{+}$

The equilibrium of the alcohol dehydrogenase reaction is far in favor of ethanol; however, ethanol conversion can be completed by continuous oxidation of acetaldehyde dehydrogenase reaction.

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FGFs in Development and Reproductive Functions

Xiaokun Li , ... Renshan Ge , in Fibroblast Growth Factors, 2018

2.7 Enzyme Assays

The substrate concentrations used for each enzyme were maximal to ensure that the concentration of substrate was not rate limiting. Control samples of culture medium alone were run in parallel with each enzyme assay. Briefly, reaction mixtures (0.2  mL) were prepared in Leydig cell medium that contained 100   nmol/L substrate (1   μCi). Reactions were initiated by adding to the reaction medium an aliquot of 0.2   ×   106 Leydig cells. Cells were maintained at 34°C in the incubator for 30   min. Reactions were terminated by adding ice-cold ethyl acetate, and steroids were rapidly extracted. The organic layer was dried under nitrogen. The radioactivity was measured using a radiometric scanner (System 200/AC3000, Bioscan, Washington, DC). The activity of 3βHSD was determined by measuring conversion of pregnenolone to progesterone. P450_c17 was determined by measuring conversion of progesterone to androstenedione. The activity of SRD5A was determined by measuring the conversion of T to dihydrotestosterone and 3α-DIOL, 17βHSD3 was determined by measuring conversion of androstenedione to T.

The steroids were separated on TLC plates in chloroform-methanol (97:3) for 3βHSD, SRD5A and 17βHSD3 assays; chloroform-ether (7:1, V/V) for P450_c17 assays.

Activity of P450_scc was determined by measuring the conversion of 22-hydroxycholesterol to testosterone.

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Future directions in alcohol dehydrogenase-catalyzed reactions

Jon D. Stewart , in Future Directions in Biocatalysis, 2007

3.1 Improving the kinetic properties of dehydrogenases

Because substrate concentrations are normally maintained at high levels to maximize volumetric productivity, turnover number and resistance to product inhibition are critical dehydrogenase properties. Compared with other enzyme classes, many dehydrogenases have relatively low k _cat values on the order of 1–10 s⁻¹. ³⁵ Increasing the turnover number allows lower catalyst loading in the reactor, which directly translates to lower biocatalyst cost. It also has the indirect benefit of reducing components such as whole cells and proteins that often complicate downstream processing steps by promoting emulsion formation. Directed evolution strategies offer a logical path toward improving turnover number.

Product inhibition is highly undesirable in bioprocesses since it limits the final product titer and volumetric productivity. Two types of product inhibition should be distinguished. Competitive binding of the alcohol product to the dehydrogenase active site that blocks substrate access is the biochemical definition of product inhibition. Bioprocesses, particularly those utilizing whole cells, may also be subject to a different type of product inhibition more properly referred to as product toxicity. In this case, the alcohol product disrupts a key property of the biocatalyst such as the integrity of the cell membrane or a metabolic process responsible for cofactor regeneration. The distinction is important because classical product inhibition may be addressed by alterations to the dehydrogenase itself; overcoming product toxicity requires a more holistic approach.

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Coupling of Flows of Substrates

Wilfred D. Stein , Thomas Litman , in Channels, Carriers, and Pumps (Second Edition), 2015

5.2.1 The Kinetics of Antiport

Either substrate, S or P, can cross the membrane on an antiporter, but only in exchange for a second substrate molecule, either S for S (self—or homoexchange) or S for P (heteroexchange).

Substrate S moves from side 1 on the antiporter at a rate proportional to its concentration S₁; P moves from side 2 at a rate proportional to P₂. Thus, an exchange of S, coming from side 1, with P, coming from side 2, will occur at a rate proportional to the product of S₁ and P₂. In similar fashion, the exchange of S at side 2 with P at side 1 occurs at a rate proportional to S₂ × P₁. Exchange of S with S or P with P has no effect on the net transport of S and P. Thus, at the steady state, where the net transport of S from side 1 to side 2 is equal to that from side 2 to side 1 we have, for uncharged substrate, that

(5.1) ${\mathrm{S}}_1{\mathrm{P}}_2={\mathrm{S}}_2{\mathrm{P}}_1$

or

(5.2) ${\mathrm{S}}_1/{\mathrm{S}}_2={\mathrm{P}}_1/{\mathrm{P}}_2$

Equation (5.2) is the fundamental rule of antiport: at the steady state, the concentration ratio of the one substrate that uses the antiporter is exactly equal to that of the other. Take the case where sodium and calcium both use the same antiporter (as is the case for the muscle cell membranes depicted in Figure 5.2B). Using the primary transport system for sodium (which we will discuss in Chapter 6), the concentration of sodium (S in Eq. (5.2)) is held about 10-fold lower on the inside of the cell (side 1) than on the outside (side 2). As a result of the presence of the antiporter, the concentration of calcium (P in Eq. (5.2)) will similarly be lower at side 1 than at side 2. The special properties of the antiporter of the muscle (see Section 5.2.4) enable the intracellular calcium to be kept at a concentration gradient of several thousand-fold, inside being low. Being able to control the low level of cytoplasmic Ca²⁺ concentration is an important aspect of muscle function. Initiation of contraction of the muscle and its subsequent relaxation occur as a result of changes in the ambient calcium concentration, brought about by calcium channels and pumps.

Just as the simple carrier systems of Chapter 4 could be electrogenic (recall Section 4.7), so, of course, can the antiporters. Indeed, in the case just considered where sodium is exchanged for calcium ions, a complete cycle of net transport on this antiporter carries three sodium ions across the muscle cell membrane in one direction for every (doubly charged) calcium ion carried in the other. There is a net transfer of one positive charge in a cycle. There is a similar situation in the important antiporter that exchanges ADP and ATP (adenosine diphosphate and adenosine triphosphate, respectively) across the membrane of the mitochondrion. Here, the ADP bears three negative charges at neutral pH, whereas the ATP bears four such charges. A cycle of exchange of ADP with ATP transfers one unit of charge across the membrane, and the transfer rates and steady-state concentration ratios will be determined by the transmembrane potential of the mitochondrial membrane. The fact that this is inside negative ensures that at the steady state ATP is concentrated outside the mitochondrion more than the ADP, i.e., that the mitochondrion exports ATP in exchange for ADP, just as is required for effective functioning of the cell. (For recent information on the molecular structure of the ATP/ADP exchanger—including the controversy of whether its monomeric form is active—and the role of this antiporter in cancer, see the section in "Suggested Readings" under ADP/ATP exchange.)

To correctly account for this transfer of charge by an antiporter, we must modify Eq. (5.2), to include the transmembrane potential Δψ and the charge Z _s on substrate S and charge Z _p on substrate P (see Box 5.1 for an introduction to the topic.). Following through Box 3.1, we obtain:

Box 5.1

Electrogenicity and the Antiporters

The proof of Eq. (5.3a) or (b) follows directly from Eq. (2.3), which we will rewrite in a modified form below. Recall that the chemical potential of a substance S is a measure of its capacity to perform work. We denote U _s,i as the chemical potential of the solute S (where the subscript i is the side of the membrane being referred to, 1 or 2), U° _s as the standard-state chemical potential (i.e., the chemical potential of S at a concentration of 1 molal (i.e., 1   mol of S per 1,000   g of water) at 0°C, and at zero electrical potential), V _s as its partial molal volume (i.e., the increment in volume per mole of solute added when an infinitesimal amount of S is added to the solution), P as the pressure exerted on the solution (in excess of atmospheric pressure), ψ _i as the electrical potential at which S finds itself at side i and, finally, Z _s as the valence of the ion S. We then can write the expression for S at side i

$U_{s,i}=U_s°+RT\ln S_i+V_{s}P+Z_{s}F{\mathrm{\psi }}_i$

To obtain the corresponding equation for the second substrate P, we write P for S throughout in the preceding equation. We now calculate the change in chemical potential on moving n _s moles of substrate S from side 1 of the membrane to side 2, and n _p of substrate P in the opposite direction. The standard-state chemical potentials are, of course, unchanged during such a transfer as are, we assume, the pressure terms. At the steady state, the overall change in chemical potential for the coupled transport of S and P is zero. Summing the two equations formed by substituting appropriately in the modified Eq. (2.3) and equating the sum to zero, we obtain

$n_{s}RT\ln (S_2/S_1)+n_s{Z}_{s}F\mathrm{\Delta }{\mathrm{\psi }}_{2\to 1}+n_{p}RT\ln (P_1/P_2)+n_p{Z}_{p}F\mathrm{\Delta }{\mathrm{\psi }}_{1\to 2}=0$

which simplifies to

$\ln {(S_2/S_1)}^{n_s}=\ln {(P_2/P_1)}^{n_p}–(n_p{Z}_p–n_s{Z}_s)F\mathrm{\Delta }{\mathrm{\psi }}_{1\to 2}/(RT)$

In the case where the number of moles of S and of P moved are the same, n _p = n _s , and we recover Eq. (5.3a).

(5.3a) $\ln \frac{{\mathrm{S}}_2}{{\mathrm{S}}_1}=\ln \frac{{\mathrm{P}}_2}{{\mathrm{P}}_1}+(Z_{\mathrm{s}}-Z_{\mathrm{p}})\frac{F\mathrm{\Delta }\psi }{RT}$

or

(5.3b) $2.303\log \frac{{\mathrm{S}}_2}{{\mathrm{S}}_1}=2.303\log \frac{{\mathrm{P}}_2}{{\mathrm{P}}_1}+(Z_{\mathrm{s}}-Z_{\mathrm{p}})\frac{F\mathrm{\Delta }\psi }{RT}$

Note that Eq. (5.3a) or (b) reduces to Eq. (5.2) when the net charge transferred across the membrane in a cycle (Z _s −Z _p) is zero, since the antiporter is in that case not electrogenic.

Consider the calcium–proton antiporter of the bacterium E. coli. This exchanges one proton for one (doubly charged) calcium and is, therefore, electrogenic. A single charge is transferred across the membrane for each transport cycle. As we saw in Section 2.2, at 25°C a transmembrane potential of 59   mV will give a transmembrane concentration ratio of 10-fold for a univalent ion. According to Eq. (5.3a) or (b), the concentration ratio of calcium (outside/inside) will be greater than that of proton by just this same factor of 10, if it is exchanged for the singly charged proton in the calcium–proton antiport cycle. (The second term on the right-hand side of Eq. (5.3b) is (at a membrane potential of 59 mV) equal to 1, the antilog of 10.) This is one example of the many proton antiporters that occur in bacteria. The full transport equation for an antiport system (corresponding to that given for the uniporters as Eq. (4.3)) is given by Stein, 1986 (see "Suggested Readings") pp. 308, 309.

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Significance of enzyme kinetics in food processing and production

Edmundo Juárez-Enríquez , ... Juan Buenrostro-Figueroa , in Value-Addition in Food Products and Processing Through Enzyme Technology, 2022

4.3 Substrate concentration

The increase in substrate concentration also increases the rate of reaction, up to a certain level. When increasing the substrate concentration using optimum levels of pH, temperature, and enzyme concentration, more substrate molecules interact with the enzyme and more product is formed. The increase in the rate of reaction is nearly proportional to the substrate concentration, but once a time that all enzymes are bonded to the substrate molecules, the addition of more substrate will have no effect on the rate of reaction, but it decreases and finally becomes constant (Fig. 34.2C). Excess of the substrate will be difficult for the substrate–enzyme interaction, due to substrates competing for the active site, which only is available for one specific substrate molecule (Kuddus, 2019; Scanlon et al., 2018).

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Acetaldehyde

Hans-Ulrich Bergmeyer , Frank Lundquist , in Methods of Enzymatic Analysis, 1965

Sensitivity

With constant substrate concentration the rate of the reaction is not linearly proportional to the enzyme concentration ( Fig. 1). With high enzyme concentrations the rate of reaction depends only on the concentration of substrate. It is preferable to use these large amounts of enzyme because this will eliminate the effect of small amounts of inhibitory compounds. As little as 0.1–0.2 μg. acetaldehyde/ml. plasma can be detected quantitatively, while about 1 μg. can still be determined accurately. The measured optical density difference corresponds to about a 50% oxidation of acetaldehyde to acetate.

Fig. 1. Relation between the rate of the reaction and the concentration of yeast aldehyde dehydrogenase. The reaction mixture consisted of 7.4 μg. acetaldehyde, 2 ml. neutralized metaphosphoric acid and 1 ml. DPN-buffer mixture (solution VIII). The points are averages of two estimations. They have been corrected for the reagent blank and the different amounts of enzyme.

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Determination of the Degradation Products Maltose and Glucose

Elli Rauscher , in Methods of Enzymatic Analysis (Second Edition), Volume 2, 1974

Optimum Conditions for Measurements

The optimum substrate concentration is 30–60 mg. of starch per ml. of incubation solution. The optimum pH value is 6.9–7.0 in 20 mM phosphate buffer. Chloride ions are added to a final concentration of 50 mM for activation.

After a fixed reaction time, the enzymes are inactivated with perchloric acid and the reaction is stopped. This prevents the 6-PGDH present as a contaminating activity in α-glucosidase preparations from converting gluconate-6-phosphate further into ribulose-5-phosphate with formation of NADPH.

Since large fragments are initially formed from starch, and a uniform degradation to maltose takes place only then, an preliminary incubation phase of about 20 min. is necessary for the α-amylase reaction. The maltose concentration is therefore determined after 20 min. and after 60 min. The α-amylase activity is found from the difference between the two values in the linear region of the reaction.

Under the conditions described here (incubation at 25°C, measurement at Hg 365 nm), measurements are possible up to an α-amylase activity of about 700 U/l. (25 °C) if the glucose concentration of the sample is in the normal range.

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