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Individual and simultaneous enzymatic determination of ethanol and acetaldehyde in wines by flow injection analysis
Analytica Chimica Acta, 185 (1986) 57-64 Elsevier Science Publishers B.V., Amsterdam -Printed
in The Netherlands
INDIVIDUAL AND SIMULTANEOUS ENZYMATIC DETERMINATION OF ETHANOL AND ACETALDEHYDE IN WINES BY FLOW INJECTION ANALYSIS
F. LAZARO,
M. D. LUQUE DE CASTRO and M. VALCARCEL*
Department of Analytical Chemistry, Faculty of Sciences, Cbrdoba (Spain)
University of Cbrdoba,
(Received 12th December 1985)
SUMMARY The individual and simultaneous enzymatic determination of ethanol and acetaldehyde in wine by flow injection analysis is described. Individual determinations of 0.002-0.016% (v/v) ethanol or 1.0-8.0 rg ml-’ acetaldehyde with r.s.d. 0.7% and 0.5%, respectively, are done with a single-beam spectrophotometer, based on the use of alcohol dehydrogenase and aldehyde dehydrogenase. A diode-array detector and dual reagent injections are used for the simultaneous determination of the two compounds. The errors are <3.5% and <2.0% for ethanol and acetaldehyde, respectively, when the method is applied to wine samples.
As a long-established commercial product of great variety, wine has been studied very thoroughly. Standard methods for the determination of oenological parameters are regulated by national governments [l] and international organizations alike [ 21. Official methods of analysis tend to be slow and so uneconomic. Owing to the inherent complexity of wines, separation techniques (distillation, ion-exchange, precipitation, etc.) are usually needed to isolate the analyte [3] . The use of flow injection analysis (f.i.a.) should allow fast determination of oenologic parameters. Enzymatic methods for the spectrophotometric determination of ethanol or acetaldehyde or both simultaneously, are suggested in this paper. The use of separation techniques prior to the determinations is unnecessary because of the high selectivity of the reactions involved and the high concentrations of these analytes in wine, compared with other alcohols and aldehydes, which necessitates considerable dilution of the samples. The determination of ethanol is based on its oxidation by NAD+ in the presence of alcohol dehydrogenase (ADH) at pH 9 [4] . Semicarbazide is used as trapping agent for the acetaldehyde formed in the reaction. CH3CH20H + NAD +&= Acetaldehyde
CH$HO
is determined
0003-2670/86/$03.50
+ NADH + H+
by oxidation
with NAD’ in the presence of
0 1986 Elsevier Science Publishers B.V.
58
aldehyde dehydrogenase, AlDH, at pH 9 [5] : CH$HO
+ NAD’ + OH- AID \ CH$OO-
+ NADH + H+
The NADH is monitored spectrophotometrically in both cases. The merging-zones mode of f.i.a. and a single-beam spectrophotometer are used for the individual determination of ethanol and acetaldehyde. The problem of the simultaneous determination of both analytes posed by their appreciably different concentrations in the samples is solved by using a diode-array detector [6]. The reversed mode of f.i.a. (in which reagent is injected into a stream of sample) is most suitable in this case. EXPERIMENTAL
Apparatus
and reagents
A Pye-Unicam SP6-500 single-beam spectrophotometer equipped with a Hellma 178.12-QS flow-cell (inner volume 18 ~1) was used for the individual determinations. A Hewlett-Packard (HP) 8451A diode-array spectrophotometer furnished with a HP-9121 floppy disc unit, a HP-98155A keyboard and a HP-7470A plotter were used in the simultaneous determinations. Two Ismatec Mini-S-840 peristaltic pumps, a Tecator type-1 manifold and a home-made dual variable-volume injection valve were used in both types of determination. Absolute ethanol standards (purity 99.5%) and distilled acetaldehyde (assayed by iodimetric back-titration [ 71) were used. Alcohol dehydrogenase (E.C. 1.1.1.1; ca. 400 U mg-’ protein), aldehyde dehydrogenase (E.C. 1.2.1.5; ca. 26.5 U mg-’ protein) and the &nicotinamide adenine dinucleotide (NAD+) were supplied by Boehringer Mannheim. The following buffer solutions were prepared. For buffer Bl, a mixture of 33.00 g 1-i NhP20,, 1.57 g 1-l glycine, 8.76 g 1-l NaCl and 8.0 g 1-l semicarbazide hydrochloride was adjusted to pH 9.0 with sodium hydroxide. For buffer B2, a mixture of 33.00 g 1-l N%P,O, and 4.9 g 1-l KC1 was adjusted to pH 9.0 with hydrochloric acid. For buffer B3, a 33.00 g 1-l Na4P20, solution was adjusted to pH 9.0 with hydrochloric acid. Sample solution Sl was wine, diluted as required with buffer Bl (itself diluted 1 + 5) for ethanol and B2 for acetaldehyde determination. Sample solution 52 was wine diluted 200-fold with diluted (1 + 5) buffer B3. Enzyme solution El was ADH (130 U ml-‘) and NAD’ (4.0 g 1-l) in buffer Bl for the individual determination of ethanol. Enzyme solution E2 was AlDH (8.0 U ml-‘) and NAD’ (4.0 g 1-l) buffer in B2. Configurations
The individual determinations were done with a symmetric merging zones manifold (Fig. 1A) in which the enzyme (El or E2) and sample (Sl) were injected into pyrophosphate buffer streams (Bl or B2) which later merged, and yielded NADH which was monitored at 340 nm. The reversed flow-
59
B
1
I
El iE21
E2
El
DAD
52 DV
W
Fig. 1. Configurations used: (A) for the individual determination; (B) for the simultaneous determinations, Sl, S2, El, E2, Bl and B2 are described in the text. DAD and DV denote the diode-array spectrophotometer and dual injection valve (composed of valves V, and V,), respectively. W is waste.
injection scheme for the simultaneous determination of ethanol and acetaldehyde involved two serial injection valves which simultaneously injected separate portions of El and E2 into the sample stream (S2) (Fig. 1B). The reactant zones reached the detector at different times. RESULTS
AND DISCUSSIONS
Enzymatic determination of ethanol The optimization of the flow-injection and chemical variables for greatest sensitivity yielded the values listed in Table 1. The glycine, pyrophosphate and sodium chloride concentrations did not affect the analytical signal; conversely, the semicarbazide concentration exerted a significant influence through its trapping effect. The concentrations of ADH and NAD+ had the most significant effect on the peak height, which increased with increasing concentration of both until it became almost constant at 130 U ml-’ and 4.0 g l-l, respectively. The sample pH could be varied between wide limits (4.0-9.5) without change in peak height because of the capacity of the buffer added to the enzyme, Bl; the pH of the buffer could also range between 7 and 10 without noticeable decrease in the analytical signal. Nevertheless, a pH of 9.0 was
60 TABLE 1 Optimum values of the variables Variable
Determination EthanoP
ADH (II ml-‘) AIDH (U ml-‘) NAD+ (g 1-l) NaJ’,G, (g l-‘) NaC!l (g 1-l) KGl (g 1-l) Glycine (g 1-l) Semicarbazide (g 1-l) Buffer pH Sample pH Flow rate (ml mine1) Reactor length R, (cm) Reactor length R, (cm) Reactor length R, (cm) Loop V, volume (~1) Loop V, volume (~1)
130 4.0 33.0 8.8 1.6 8.0 9.0 4.0-9.5 0.78 6.0 6.0 193 30.0 60.0
Acetaldehydes
8.0 4.0 33.0 14.9 9.0 3.3-9.5 0.62 6.0 6.0 140 60.0 90.0
Both analytes simultaneouslyb 130 8.0 4.0 33.0 8.8 14.9 1.6 9.0 4.0-9.5 1.10 262 136 30.0 90.0
*Manifold A, Fig. 1. bManifold B, Fig. 1.
chosen for Bl as this was the most suitable for the determination of acetaldehyde (see below). Temperatures of G25”C were satisfactory. The lengths of reactors R0 and R1 were the minimum necessary to connect the two injection valves with the manifold. The length of Rz and the flow rate were chosen so as to obtain the maximum signal. The volume of sample was always smaller than that of the enzyme solution in order to keep the former surrounded by ADH/NAD+ solution. The sampling frequency achieved for ethanol determinations was 55 h-l. The determination of ethanol in wine poses a problem deriving from the high concentration of the analyte, and requires much dilution (up to 1:5000). This was implemented by one of three procedures. The manual dilution of the sample prior to introduction into the flow system is the commonest procedure, but is somewhat inaccurate in that it often requires the measurement of a small sample volume. Nevertheless, it allows a wide linear calibration range, and good sensitivity and relative standard deviation (Table 2). The results obtained for ethanol in different types of wine and brandy from Jerez are compared to those obtained by the EEC method [8] in Table 3. The flow dilution technique involves utilizing asymmetric merging zones (chasing zones) in which the “head” of the enzyme sector merges with the tail of the sample. The greater the distance between the heads of the two sectors, the greater the dilution. In this case R,/R, was 8. This procedure afforded a linear calibration range and a precision comparable to that of the manual
61 TABLE 2 Calibration parameters for the determination
of ethanol
Dilution method
Parameter
Linear range (% v/v) Slope (%-’ ) Correlation coefficienta R.s.d. (%)b
Manual
Flow
Photometric
0.004-0.016 20.207
0.06-0.26 1.224
0.002-0.016 14.024
0.9955 1.1
a5 points. bll determinations ethanol (flow).
0.9988 1.1
of 0.008%
0.9944 0.7
ethanol (manual and photometric)
and 0.16%
TABLE 3 Determination
of ethanol in wine and brandy
SamDle
Wine 1 Wine 2 Wine 3 Wine 4 Wine 5 Wine 6 Wine 7 Wine 8 Brandy 1 Brandy 2
Ethanol concentrationa (% v/v) 14.37 15.17 15.26 15.35 17.45 19.90 20.10 15.46 39.95 66.85
Average error
Errors Manual dilution
Flow dilution
+7.2 -4.5 +l.O -9.1
t 22.6 +13.5 +13.3
t4.3
+ll.O +19.9 -0.1 -2.2 -4.4 -6.9 9.7
-0.5 4.3
t2.3 -2.1 -6.7 3.9
t8.3
Photometric dilution +14.8 t1.7 -5.1 -3.4 +0.7 -2.8 -0.8 +4.4 -1.9 -2.6 3.5
aEEC official method.
dilution procedure; however, the errors incurred in its application to real samples were considerably larger (see Table 3). The photometric “dilution” method is based on measuring absorbances at wavelengths away from the absorption maximum [6] and was applied in conjunction with manual dilution to diminish the sensitivity of the method further, thus slightly increasing the linear calibration range (to 0.002-0.016%) and hence decreasing the extent of manual dilution of the sample required. This photometric procedure has the greatest accuracy (average error 3.5%) for the analysis of real samples, and the greatest precision (r.s.d. = 0.7%). Enzymatic determination of acetaldehyde The results obtained in the optimization of variables for acetaldehyde also appear in Table 1. As in the ethanol method, the pyrophosphate concentra-
62
tion did not affect the analytical signal, but the signal did increase with increasing potassium chloride concentration, reaching a maximum at 14.9 g 1-i. Increasing the NAD+ and enzyme concentrations increased the peak height, until it became constant above 4.0 g 1-l and 8.0 U ml-‘, respectively. Changing the NAD+ concentration had a less significant effect than changing the AlDH concentration. The optimum pH range was 8-10, while the sample pH could range between 3.5 and 9.5 without appreciably altering the signal. Acetaldehyde present in wine is found free and bound to sulphur dioxide; therefore, to determine total acetaldehyde, the pH of the carrier solution was adjusted to 9.0 and a 20% (v/v) concentration of this buffer was added to the samples to obtain a final pH of 9.0-9.5, to ensure that all acetaldehyde present was unbound. The other variables (flow injection and temperature) were optimized as for ethanol, except that the sample and enzyme volumes used were somewhat larger so as to increase sensitivity. The sampling frequency attained was 50 h-i. The calibration graph was linear for 1.0-8.0 pg ml-’ acetaldehyde (correlation coefficient 0.9988, 5 points) with a slope of 0.0335 ml pg-‘. The relative standard deviation (11 determinations of 5 pg ml-‘) was 0.5%. Owing to its volatility, acetaldehyde was determined in wine by the standard addition method. The values shown in Table 4 were obtained for different sherries. The recovery and mean deviation were 102% and 2.0%, respectively. Simultaneous determination of ethanol and acetaldehyde The most serious problem encountered in the simultaneous determination of these compounds is the different dilution required for each (average dilution 1:1500 and 1:80 for ethanol and acetaldehyde, respectively). Attempts were made to bring the determination ranges of both species closer by not using semicarbazide in solution Bl, in order to decrease the extent of alcohol oxidation, and by measuring the absorbance at other than the wavelength of maximum absorbance, but the results were unacceptable because the signal from acetaldehyde was too small. TABLE 4 Determination Wine
of acetaldehyde Concentration found (pg ml-‘) 178 131 145 473 362 143 148 334
Recovery of standard additions (%) 2.0 fig ml-’ 98 96 99
112 100 102 106 103
5.0 fig ml-’ 103 107 103 106 91 103 104 99
63 TABLE 5 Simultaneous determination of ethanol and acetaldehyde Sample
1 2 3 4 5
Ethanol
Acetaldehyde
Found (% v/v)
Recovery (%) 0.04%
13.4 12.0 13.3 12.7 13.4
105 97 101 101 100
0.08%
Found (rg ml-‘)
Recovery (%) 0.4 rg ml”
0.8 rg ml-l
93 100 105 100 94
56 102 135 221 288
90 104 105 104 102
107 99 101 104 98
Finally, a simple flow configuration (Fig. 1B) was chosen in which solutions of different composition (each with a different enzyme) were injected into a stream of diluted (1:200-1:300) sample, The indicator reaction for acetaldehyde developed in reactor RI, and in R,, and R1 for ethanol. The two reacting sectors reached the diode-array spectrophotometer at different times. Absorbance measurements based on the sum of the absorbances in the 336, 338, 340, 342 and 344~nm channels were used for acetaldehyde measurements, which changed the range of the method for acetaldehyde from 1.0-8.0 pg ml-’ to 0.3-2.0 pg ml-‘. Semicarbazide was not used in the assay for ethanol and the NADH was monitored at 360 nm, in order to decrease the sensitivity. Also, as the calibration graph for ethanol decreased in sensitivity at higher concentrations, this less sensitive range was used for measurement of the ethanol concentrations in the samples after dilution as described above. Table 5 shows the recoveries obtained on addition to the wine samples of 0.4 and 0.8 pg ml-’ acetaldehyde and 0.04% and 0.08% ethanol. The average recoveries are 101.4% and 98.8%, with absolute average deviations of 4.7% and 3.3% for acetaldehyde and ethanol, respectively. Conclusions Comparison of the flow-injection methods with the official EEC methods shows that the flow-injection technique offers a number of advantages. These include lower sample and reagent consumption, less sample manipulation (no prior separation of the analytes is necessary), lower determination limits, high analysis rate (50-55 samples per hour for one analyte), and the simultaneous determination of both analytes. The CAICyT is thanked for financial support (Grant No. 2012-83). The authors gratefully acknowledge Gonzalez Byass for providing samples of wine.
64 REFERENCES 1 France, Mini&e de l’Agriculture, ArrBte du 24 juin 1963 relatif aux Methodes Officielles d’AnaIyses des Vins et des Mofits, Journal Officiel No. 63-154,1037 (1963) 4561. 2 Recueil des Methodes Intemationales d’Analyse des Vins, Office International de la Vigne et du Vin, Paris, 1962-1973. 3 J. Ribereau-Gayon, E. Peynaud, P. Sudran and P. Ribereau-Gayon, Traite d’oenologie. Sciences et Techniques du Vin, Tome I, Dunod, Paris, 1976. 4 E. Bemt and I. Gutmann, in H. U. Bergmeyer (Ed.), Methods of Enzymatic Analysis, 2nd edn., Vol. 3, Verlag Chemie, Weinheim and Academic Press, New York, 1974, p. 1499. 5 F. Lundquist, in H. U. Bergmeyer (Ed.), Methods of Enzymatic Analysis, 2nd edn., Vol. 3, Verlag Chemie, Weinheim, and Academic Press, New York, 1974, p. 1509. 6 F. LazPo, A. Rios, M. D. Luque de Castro and M. VaicBrcel, Anal. Chim. Acta, 179 (1986) 279. 7 Official Methods of Analysis, 12th edn., Association of Official Analytical Chemists, Washington, DC, 1975, p. 603. 8 Journal Officiel des CommunauMs Europeennes, L. 113,14 Mai 1982, p. 16.
in The Netherlands
INDIVIDUAL AND SIMULTANEOUS ENZYMATIC DETERMINATION OF ETHANOL AND ACETALDEHYDE IN WINES BY FLOW INJECTION ANALYSIS
F. LAZARO,
M. D. LUQUE DE CASTRO and M. VALCARCEL*
Department of Analytical Chemistry, Faculty of Sciences, Cbrdoba (Spain)
University of Cbrdoba,
(Received 12th December 1985)
SUMMARY The individual and simultaneous enzymatic determination of ethanol and acetaldehyde in wine by flow injection analysis is described. Individual determinations of 0.002-0.016% (v/v) ethanol or 1.0-8.0 rg ml-’ acetaldehyde with r.s.d. 0.7% and 0.5%, respectively, are done with a single-beam spectrophotometer, based on the use of alcohol dehydrogenase and aldehyde dehydrogenase. A diode-array detector and dual reagent injections are used for the simultaneous determination of the two compounds. The errors are <3.5% and <2.0% for ethanol and acetaldehyde, respectively, when the method is applied to wine samples.
As a long-established commercial product of great variety, wine has been studied very thoroughly. Standard methods for the determination of oenological parameters are regulated by national governments [l] and international organizations alike [ 21. Official methods of analysis tend to be slow and so uneconomic. Owing to the inherent complexity of wines, separation techniques (distillation, ion-exchange, precipitation, etc.) are usually needed to isolate the analyte [3] . The use of flow injection analysis (f.i.a.) should allow fast determination of oenologic parameters. Enzymatic methods for the spectrophotometric determination of ethanol or acetaldehyde or both simultaneously, are suggested in this paper. The use of separation techniques prior to the determinations is unnecessary because of the high selectivity of the reactions involved and the high concentrations of these analytes in wine, compared with other alcohols and aldehydes, which necessitates considerable dilution of the samples. The determination of ethanol is based on its oxidation by NAD+ in the presence of alcohol dehydrogenase (ADH) at pH 9 [4] . Semicarbazide is used as trapping agent for the acetaldehyde formed in the reaction. CH3CH20H + NAD +&= Acetaldehyde
CH$HO
is determined
0003-2670/86/$03.50
+ NADH + H+
by oxidation
with NAD’ in the presence of
0 1986 Elsevier Science Publishers B.V.
58
aldehyde dehydrogenase, AlDH, at pH 9 [5] : CH$HO
+ NAD’ + OH- AID \ CH$OO-
+ NADH + H+
The NADH is monitored spectrophotometrically in both cases. The merging-zones mode of f.i.a. and a single-beam spectrophotometer are used for the individual determination of ethanol and acetaldehyde. The problem of the simultaneous determination of both analytes posed by their appreciably different concentrations in the samples is solved by using a diode-array detector [6]. The reversed mode of f.i.a. (in which reagent is injected into a stream of sample) is most suitable in this case. EXPERIMENTAL
Apparatus
and reagents
A Pye-Unicam SP6-500 single-beam spectrophotometer equipped with a Hellma 178.12-QS flow-cell (inner volume 18 ~1) was used for the individual determinations. A Hewlett-Packard (HP) 8451A diode-array spectrophotometer furnished with a HP-9121 floppy disc unit, a HP-98155A keyboard and a HP-7470A plotter were used in the simultaneous determinations. Two Ismatec Mini-S-840 peristaltic pumps, a Tecator type-1 manifold and a home-made dual variable-volume injection valve were used in both types of determination. Absolute ethanol standards (purity 99.5%) and distilled acetaldehyde (assayed by iodimetric back-titration [ 71) were used. Alcohol dehydrogenase (E.C. 1.1.1.1; ca. 400 U mg-’ protein), aldehyde dehydrogenase (E.C. 1.2.1.5; ca. 26.5 U mg-’ protein) and the &nicotinamide adenine dinucleotide (NAD+) were supplied by Boehringer Mannheim. The following buffer solutions were prepared. For buffer Bl, a mixture of 33.00 g 1-i NhP20,, 1.57 g 1-l glycine, 8.76 g 1-l NaCl and 8.0 g 1-l semicarbazide hydrochloride was adjusted to pH 9.0 with sodium hydroxide. For buffer B2, a mixture of 33.00 g 1-l N%P,O, and 4.9 g 1-l KC1 was adjusted to pH 9.0 with hydrochloric acid. For buffer B3, a 33.00 g 1-l Na4P20, solution was adjusted to pH 9.0 with hydrochloric acid. Sample solution Sl was wine, diluted as required with buffer Bl (itself diluted 1 + 5) for ethanol and B2 for acetaldehyde determination. Sample solution 52 was wine diluted 200-fold with diluted (1 + 5) buffer B3. Enzyme solution El was ADH (130 U ml-‘) and NAD’ (4.0 g 1-l) in buffer Bl for the individual determination of ethanol. Enzyme solution E2 was AlDH (8.0 U ml-‘) and NAD’ (4.0 g 1-l) buffer in B2. Configurations
The individual determinations were done with a symmetric merging zones manifold (Fig. 1A) in which the enzyme (El or E2) and sample (Sl) were injected into pyrophosphate buffer streams (Bl or B2) which later merged, and yielded NADH which was monitored at 340 nm. The reversed flow-
59
B
1
I
El iE21
E2
El
DAD
52 DV
W
Fig. 1. Configurations used: (A) for the individual determination; (B) for the simultaneous determinations, Sl, S2, El, E2, Bl and B2 are described in the text. DAD and DV denote the diode-array spectrophotometer and dual injection valve (composed of valves V, and V,), respectively. W is waste.
injection scheme for the simultaneous determination of ethanol and acetaldehyde involved two serial injection valves which simultaneously injected separate portions of El and E2 into the sample stream (S2) (Fig. 1B). The reactant zones reached the detector at different times. RESULTS
AND DISCUSSIONS
Enzymatic determination of ethanol The optimization of the flow-injection and chemical variables for greatest sensitivity yielded the values listed in Table 1. The glycine, pyrophosphate and sodium chloride concentrations did not affect the analytical signal; conversely, the semicarbazide concentration exerted a significant influence through its trapping effect. The concentrations of ADH and NAD+ had the most significant effect on the peak height, which increased with increasing concentration of both until it became almost constant at 130 U ml-’ and 4.0 g l-l, respectively. The sample pH could be varied between wide limits (4.0-9.5) without change in peak height because of the capacity of the buffer added to the enzyme, Bl; the pH of the buffer could also range between 7 and 10 without noticeable decrease in the analytical signal. Nevertheless, a pH of 9.0 was
60 TABLE 1 Optimum values of the variables Variable
Determination EthanoP
ADH (II ml-‘) AIDH (U ml-‘) NAD+ (g 1-l) NaJ’,G, (g l-‘) NaC!l (g 1-l) KGl (g 1-l) Glycine (g 1-l) Semicarbazide (g 1-l) Buffer pH Sample pH Flow rate (ml mine1) Reactor length R, (cm) Reactor length R, (cm) Reactor length R, (cm) Loop V, volume (~1) Loop V, volume (~1)
130 4.0 33.0 8.8 1.6 8.0 9.0 4.0-9.5 0.78 6.0 6.0 193 30.0 60.0
Acetaldehydes
8.0 4.0 33.0 14.9 9.0 3.3-9.5 0.62 6.0 6.0 140 60.0 90.0
Both analytes simultaneouslyb 130 8.0 4.0 33.0 8.8 14.9 1.6 9.0 4.0-9.5 1.10 262 136 30.0 90.0
*Manifold A, Fig. 1. bManifold B, Fig. 1.
chosen for Bl as this was the most suitable for the determination of acetaldehyde (see below). Temperatures of G25”C were satisfactory. The lengths of reactors R0 and R1 were the minimum necessary to connect the two injection valves with the manifold. The length of Rz and the flow rate were chosen so as to obtain the maximum signal. The volume of sample was always smaller than that of the enzyme solution in order to keep the former surrounded by ADH/NAD+ solution. The sampling frequency achieved for ethanol determinations was 55 h-l. The determination of ethanol in wine poses a problem deriving from the high concentration of the analyte, and requires much dilution (up to 1:5000). This was implemented by one of three procedures. The manual dilution of the sample prior to introduction into the flow system is the commonest procedure, but is somewhat inaccurate in that it often requires the measurement of a small sample volume. Nevertheless, it allows a wide linear calibration range, and good sensitivity and relative standard deviation (Table 2). The results obtained for ethanol in different types of wine and brandy from Jerez are compared to those obtained by the EEC method [8] in Table 3. The flow dilution technique involves utilizing asymmetric merging zones (chasing zones) in which the “head” of the enzyme sector merges with the tail of the sample. The greater the distance between the heads of the two sectors, the greater the dilution. In this case R,/R, was 8. This procedure afforded a linear calibration range and a precision comparable to that of the manual
61 TABLE 2 Calibration parameters for the determination
of ethanol
Dilution method
Parameter
Linear range (% v/v) Slope (%-’ ) Correlation coefficienta R.s.d. (%)b
Manual
Flow
Photometric
0.004-0.016 20.207
0.06-0.26 1.224
0.002-0.016 14.024
0.9955 1.1
a5 points. bll determinations ethanol (flow).
0.9988 1.1
of 0.008%
0.9944 0.7
ethanol (manual and photometric)
and 0.16%
TABLE 3 Determination
of ethanol in wine and brandy
SamDle
Wine 1 Wine 2 Wine 3 Wine 4 Wine 5 Wine 6 Wine 7 Wine 8 Brandy 1 Brandy 2
Ethanol concentrationa (% v/v) 14.37 15.17 15.26 15.35 17.45 19.90 20.10 15.46 39.95 66.85
Average error
Errors Manual dilution
Flow dilution
+7.2 -4.5 +l.O -9.1
t 22.6 +13.5 +13.3
t4.3
+ll.O +19.9 -0.1 -2.2 -4.4 -6.9 9.7
-0.5 4.3
t2.3 -2.1 -6.7 3.9
t8.3
Photometric dilution +14.8 t1.7 -5.1 -3.4 +0.7 -2.8 -0.8 +4.4 -1.9 -2.6 3.5
aEEC official method.
dilution procedure; however, the errors incurred in its application to real samples were considerably larger (see Table 3). The photometric “dilution” method is based on measuring absorbances at wavelengths away from the absorption maximum [6] and was applied in conjunction with manual dilution to diminish the sensitivity of the method further, thus slightly increasing the linear calibration range (to 0.002-0.016%) and hence decreasing the extent of manual dilution of the sample required. This photometric procedure has the greatest accuracy (average error 3.5%) for the analysis of real samples, and the greatest precision (r.s.d. = 0.7%). Enzymatic determination of acetaldehyde The results obtained in the optimization of variables for acetaldehyde also appear in Table 1. As in the ethanol method, the pyrophosphate concentra-
62
tion did not affect the analytical signal, but the signal did increase with increasing potassium chloride concentration, reaching a maximum at 14.9 g 1-i. Increasing the NAD+ and enzyme concentrations increased the peak height, until it became constant above 4.0 g 1-l and 8.0 U ml-‘, respectively. Changing the NAD+ concentration had a less significant effect than changing the AlDH concentration. The optimum pH range was 8-10, while the sample pH could range between 3.5 and 9.5 without appreciably altering the signal. Acetaldehyde present in wine is found free and bound to sulphur dioxide; therefore, to determine total acetaldehyde, the pH of the carrier solution was adjusted to 9.0 and a 20% (v/v) concentration of this buffer was added to the samples to obtain a final pH of 9.0-9.5, to ensure that all acetaldehyde present was unbound. The other variables (flow injection and temperature) were optimized as for ethanol, except that the sample and enzyme volumes used were somewhat larger so as to increase sensitivity. The sampling frequency attained was 50 h-i. The calibration graph was linear for 1.0-8.0 pg ml-’ acetaldehyde (correlation coefficient 0.9988, 5 points) with a slope of 0.0335 ml pg-‘. The relative standard deviation (11 determinations of 5 pg ml-‘) was 0.5%. Owing to its volatility, acetaldehyde was determined in wine by the standard addition method. The values shown in Table 4 were obtained for different sherries. The recovery and mean deviation were 102% and 2.0%, respectively. Simultaneous determination of ethanol and acetaldehyde The most serious problem encountered in the simultaneous determination of these compounds is the different dilution required for each (average dilution 1:1500 and 1:80 for ethanol and acetaldehyde, respectively). Attempts were made to bring the determination ranges of both species closer by not using semicarbazide in solution Bl, in order to decrease the extent of alcohol oxidation, and by measuring the absorbance at other than the wavelength of maximum absorbance, but the results were unacceptable because the signal from acetaldehyde was too small. TABLE 4 Determination Wine
of acetaldehyde Concentration found (pg ml-‘) 178 131 145 473 362 143 148 334
Recovery of standard additions (%) 2.0 fig ml-’ 98 96 99
112 100 102 106 103
5.0 fig ml-’ 103 107 103 106 91 103 104 99
63 TABLE 5 Simultaneous determination of ethanol and acetaldehyde Sample
1 2 3 4 5
Ethanol
Acetaldehyde
Found (% v/v)
Recovery (%) 0.04%
13.4 12.0 13.3 12.7 13.4
105 97 101 101 100
0.08%
Found (rg ml-‘)
Recovery (%) 0.4 rg ml”
0.8 rg ml-l
93 100 105 100 94
56 102 135 221 288
90 104 105 104 102
107 99 101 104 98
Finally, a simple flow configuration (Fig. 1B) was chosen in which solutions of different composition (each with a different enzyme) were injected into a stream of diluted (1:200-1:300) sample, The indicator reaction for acetaldehyde developed in reactor RI, and in R,, and R1 for ethanol. The two reacting sectors reached the diode-array spectrophotometer at different times. Absorbance measurements based on the sum of the absorbances in the 336, 338, 340, 342 and 344~nm channels were used for acetaldehyde measurements, which changed the range of the method for acetaldehyde from 1.0-8.0 pg ml-’ to 0.3-2.0 pg ml-‘. Semicarbazide was not used in the assay for ethanol and the NADH was monitored at 360 nm, in order to decrease the sensitivity. Also, as the calibration graph for ethanol decreased in sensitivity at higher concentrations, this less sensitive range was used for measurement of the ethanol concentrations in the samples after dilution as described above. Table 5 shows the recoveries obtained on addition to the wine samples of 0.4 and 0.8 pg ml-’ acetaldehyde and 0.04% and 0.08% ethanol. The average recoveries are 101.4% and 98.8%, with absolute average deviations of 4.7% and 3.3% for acetaldehyde and ethanol, respectively. Conclusions Comparison of the flow-injection methods with the official EEC methods shows that the flow-injection technique offers a number of advantages. These include lower sample and reagent consumption, less sample manipulation (no prior separation of the analytes is necessary), lower determination limits, high analysis rate (50-55 samples per hour for one analyte), and the simultaneous determination of both analytes. The CAICyT is thanked for financial support (Grant No. 2012-83). The authors gratefully acknowledge Gonzalez Byass for providing samples of wine.
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