ANALYTICAL CHEMISTRY, VOL. 51, NO. 8, JULY 1979
1315
Determination of Polycyclic Aromatic Hydrocarbons in Aqueous Samples by Reversed-Phase Liquid Chromatography Kenneth Ogan, Elena Katz, and Walter Savin" Perkin-€/mer Corp., Norwalk, Connecticut 06856
An analytical method is described for the determination of 16 polycyclic aromatic hydrocarbons (PAHs) in aqueous samples. These 16 PAHs are of environmental concern, since 15 of them are included on the EPA Priority Pollutant list. After liquid-liquid extraction, the concentrated samples are analyzed by reversed-phase liquid chromatography with fluorescence detection. These 16 PAHs are almost fully resolved chromatographically, so they can be determined in a single run. The method is applied to the analysis of several environmental water samples and several of these compounds are quantitated at concentrations below 10 ng/L in the original sample.
In 1971 the World Health Organization set an upper limit of 200 ng/L for the total concentration of six PAHs in domestic waters ( I ) . -4dditional PAHs were known to occur in water samples but only these six could be easily separated by the analytical methods in use at t h a t time (2). In 1976. in response to a suit by the Natural Resources Defense Council, the U.S. Environmental Protection Agency (EPA) drew up a list of organic compounds to be monitored and limited in effluent waters in the United States (3). Included on this priority pollutant list are 15 of the PAHs listed in Figure 1. T h e E P A is now in the process of establishing upper concentration limits for these compounds and recommending analytical methods for their determination. Modern liquid chromatography (LC) offers many advantages for t h e analysis of PAHs in environmental samples (4-17). Many of the PAHs suspected of mutagenic or carcinogenic activity contain four to six fused rings. While most analyses based on LC have dealt with only a few members of this PAH category, two recent reports (13,16) do treat much larger sets. Three groups of PAHs within this category are often poorly resolved: (a) benz[a]anthracene and chrysene (four rings; see Figure l),(b) benzo[e]pyrene, benzo[n]pyrene, benzo[b]fluoranthene, and benzo[h]fluoranthene (five rings), and (c) benzo[ghz]perylene and indeno[l,2,3-cd]pyrene (six rings). We have obtained chromatographic separation of all of these PAHs, together with the other PAHs on the E P A Priority Pollutant list. This chromatographic separation, combined with the sensitivity and specificity of fluorescence detection, produces a n excellent means of analyzing the concentrated extracts of aqueous environmental samples for these PAHs a t levels as low as 1 ng/L in the original sample. While we provide some examples of the determination of P A H pollutants in real samples, the full method including sample preparation must still be validated for each particular kind of sample.
EXPERIMENTAL Apparatus. We used a Perkin-Elmer Series 3 liquid chromatograph and a Model 650-1OLC fluorescence detector. The chromatography conditions are given in Table I. Reagents. Chrysene, benzo[ghi]perylene, indeno[l,2,3-cd]pyrene were obtained in pure form from Analabs, Inc. (North Haven, Conn.). The benzo[c]pyrene was obtained from Aldrich Chem. Co. (Milwaukee, Wis.). These standards, which were 0003-2700/79/0351-1315$01.00/0
Table I. Chromatographic Conditions liquid chromatograph column mobile phase
detect or
data collection
Perkin-Elmer Series 3, with dynamic stirrer accessory, Rheodyne Model 7120 injection valve, 1O-bL loop Perkin-Elmer HC-ODs, 0.26 x 2 5 cm 10 + m , C : , packing, Part SO.089-0,16 acetonitrile i n water: T1: 30CL-50%, 15 min T2: 50';-100%, 8 min, linear T3: i O O T - l O O C ; , 2; min 'r purge: 20%, 5 min T equil: 50'6, 1 5 min 0.3 m l ' m i n flow rate room temperature Perkin-Elmer IIodel 6 50-1OLC, spectrofluorimeter. See Figure 2 for excitation and emission wavelengths. Bandpass = 1 2 nm Perkin-Elmer Sigma 10 chromatography data stat ion
Table 11. Detection Limit for Each PAH as Calculated from t h e Signal-to-Noise Ratio of the Individual Peak in Chromatograms
compound naphthalene acenaphthene fluorene phenanthrene anthracene fluoranthene pyrene benz[a]anthracene chrysene benzo[ elpyrene benzo[ blfluoranthene benzo[h]fluoranthene benzo[ alpyrene dibenz [a,h]anthracene benzo [ghilperylene indeno [ 1,2,3-cd]pyrene
detection stock limit, pg, mixture, (comproconcn., mise wavepg/mL lengths) 4.84 0.022 1.46 12.2 14.5 9.68 19.4 14.5 14.5 9.68 1.45 0.097 1.45 2.90 2.90 1.94
30 0.5 12 130
120 40
75 35 70 45 3 0.3
2.5 4 9 8
available as powders, were dissolved in acetonitrile at 500 pg/mL. The benzo[k]fluoranthene and benzo[b]fluoranthene were obtained from Nanogens (Watsonville, Calif.). The other PAH standards were purchased as 100 pg/mL solutions in hexane from Chem Service (West Chester, Pa.). Acetonitrile was used as purchased from Burdick & Jackson Labs (Muskegon, Mich.). Water was filtered, deionized, and purified by a carbon bed system from Continental Water Conditioning Corp. (El Paso, Tex.). A stock mixture of PAH standards was made up from the individual solutions. Since the several PAH standards were prepared in different solvents, the stock mixture was made up in a solvent that was compatible containing 81% hexane, 6% acetonitrile, 11.4% chloroform, and 1.6% toluene. The concentrations of the individual PAHs in the mixture, given in Table 11, were chosen to give comparable fluorescence peaks with the C 1979 American Chemical Society
1316
ANALYTICAL CHEMISTRY, VOL. 51, NO. 8, JULY 1979 COMPOUND
COMPOUND
NAPHTHALENE
NPh
CHRYSENE
ChY
ACENAPHTHENE
Ace
BENZO[e] PYRENE
B [el PY
FLUORENE
FI BEN20 [b] FLUOR A NTHENE
B [b] Ft
BENZO [K] FLUORANTHENE
B[k]Ft
BENZO[a] PYRENE
B[dPy
DIBENZ [a, h] ANTHRACENE
di B [a, h] A
BENZO [ghi]PERYLENE
B [g hi] Per
INDENO [1,2,3 -cd] PY RENE
I[1,2,3 - cd] Py
PHENANTHRENE
Phe
ANTHRACENE
An
FLUORANTH ENE
Ft
PYRENE
PY
BEN2 [a] ANTHRACENE
B [a3 A
Flgure 1. Identification, structures, and abbreviations of PAHs used in this work. All but benzo[e]pyrene are on the U.S. EPA Priority Pollutant
list
8 SOLVENT
PROGRAM
~
'10 ACETONITRILE IN WATER 130% I I I
b
z
@ DETECTOR WAVELENGTH PROGRAM I I I 1 I
I
Xe,=280nm Xem:340
nm
I
.J I
_____-I
I
X,,=305 nm X,,:430 nm
l
1 1
Xe,:305 nm Xe,:500
- - .nm
Figure 2. (A) Solvent program for chromatographic analysis showing
the pump control profile. (6)Detector wavelength program for monitoring the PAHs in Figure 1 wavelength program in Figure 2B. Portions of this mixture of standards were diluted 1:10,1:50,1:100,and 1:200 with acetonitrile for use as standards. Sample Preparation, The sample was collected in 1-L brown glass bottles with caps lined with PTFE (polytetrafluoroethylene). In the laboratory, 1 L of the sample was transferred to a 1-L separatory funnel (with a PTFE stopcock) and was extracted with three successive 60-mL portions of dichloromethane. The sampling bottle was washed with each dichloromethane portion, prior to adding the portion to the separatory funnel. The three dichloromethane extract fractions were combined and dried by passage through a bed of anhydrous MgS04, about 3 cm thick and 5 cm in diameter. The volume of the extract was reduced by solvent evaporation, using a distillation apparatus or a Kuderna-Danish evaporator, When the extract volume was reduced to about 5 mL, 0.5 mL of acetonitrile was added and the remaining dichloromethane was evaporated. Analysis. The chromatographic analysis used the threesegment solvent program shown in Figure 2A, and the detector excitation and emission wavelength program shown in Figure 2B. In order to equilibrate the column for the next run, the mobile phase was changed to 20% acetonitrile for 6 min, followed by 15 min at 50% acetonitrile. Identical equilibration periods between injections must be maintained to obtain reproducible retention times. An equilibration time of 21 min together with the analysis time of 50 min yielded a 71-min analysis. For sample injection, the 1O-pL sample loop was first washed with a t least 30 pL of acetonitrile, followed by air. A minimum of 15 pL of the concentrated extract was then introduced with the syringe in order to ensure complete and reproducible filling of the sample loop.
J
L ~~
1
.-.
~
10
2c
30
43
MINUTES
Figure 3. Chromatogram of 12 PAH standards using isocratic elution
(80% acetonitrile in water at a flow rate of 0.5 mL/min) at room temperature. For peak identification, see Figure 1. P is perylene
Calculation, The response factor, RF, is defined as the ratio of the peak area to the amount of analyte standard injected. Because we used several gain settings on the fluorescence detector to cover the desired dynamic range (about 20-fold), the standard dilutions were selected so that at least one dilution was used in each range. Thus each PAH had an RF for each range. We determined these RF values each day. If analysis of a sample extract yields X , ng injected (peak area times response factor) of the ith PAH, the concentration of this compound in the original aqueous sample is given by C, = X , / ( f V c , )ng/L
where f is the ratio of the injection volume to the total extract volume, V is the volume (in liters) of the original aqueous sample, and tLis the extraction efficiency for that PAH. (We have taken t to be 1.0.)
RESULTS Figure 3 is a n isocratic separation of 12 of the PAHs in Figure 1,demonstrating separation of the members of the 3 subgroups of PAHs described in the introduction. Figure 4 is the chromatogram of the PAHs in Figure 1,using the solvent program and detector wavelength program in Figure 2. The detection limit for each PAH was calculated from the signal-to-noise ratio of the individual peaks in chromatograms such as Figure 4, assuming a minimum detectable signal-
ANALYTICAL CHEMISTRY, VOL. 51, NO. 8,JULY 1979
1317
I
-
.__ I
.~
3
23
30
4c
c
MINUTES
Flgure 4. Chromatogram of 16 PAH standards using the solvent and detector wavelength programs in Figure 2. For peak identification, see Figure 1. The tail on peak 6 is an impurity in the solvent
20
IC
30
40
MINUTES
Figure 6. Chromatogram of the extract of a plastics manufacturing effluent water obtained with the conditions in Figure 2. For peak identification, see Figure 1 , ker=3C5nm i e x = 2 8 0 nm , A e m = 5 0 0nm > Aerr=340 n n kem:430 nm I 1,n t -. z ~
1 -
ke,:Z80 nm
I-
W
+
hem:340 nm
&
z
0 w
r
I 1
1.
r-"
L
I
I
I
0
IO
20
30
40
50
MINUTES I 0
,
10
1 30
215
40
50
MINUTES
Figure 5. Chromatogram of the extract of flyash wash water obtained with the conditions in Figure 2. For peak identification, see Figure 1
to-noise level of 2. These values are tabulated in Table 11. The excitation and emission wavelengths given in Figure 2B are compromise settings; a t least an order of magnitude improvement in detection limit can be achieved for some of these PAHs by using the optimum excitation and emission wavelengths for the individual compounds. T h e linearity between fluorescence response and concentration was verified for these PAHs. Dilutions ranging from 1:lO to 1:200 of the stock mixture of PAH standards were run under the conditions of Figure 2, with the gain of the fluorescence detector set to appropriate values for the various dilutions. The gain settings were used to scale the peak areas relative to the most dilute standard. Plots of the logarithm of scaled peak area vs. the logarithm of the PAH concentration were linear, with slopes not significantly different from 1.0 for each of the PAHs. T h e retention time precision within one day, using the solvent program in Figure 2A, ranges from 0.8% to 1.6%, relative standard deviation, among the first seven compounds (through fluoranthene), and 0.5% to 0.7% for the remaining compounds. Peak area precision within one day ranges from 3% to 8 % , relative standard deviation. Comparisons among analyses from day-to-day produced only a slight reduction of precision. We applied this analytical method to several real environmental samples and three examples are presented in this
Figure 7. Chromatogram of the extract of a river water sample obtained with the conditions in Figure 2. For peak identification, see Figure 1
paper. Figure 5 is the chromatogram of the extract of a fly ash wash water sample, analyzed using the programs in Figure 2. Figure 6 is the chromatogram of the extract of an aqueous effluent from a plastics manufacturing site. The collection and extraction of these samples were done by Battelle Columbus Laboratories. The preparation of these samples was similar to the procedure described under Sample Preparation, but included a sample fractionation step. The dichloromethane extract volume was reduced to 0.5 mL in a Kuderna-Danish evaporator and applied to a 1 X 25 cm column of silica gel with 25 mL of n-pentane followed by 25 mL of 40% CH2C1, in n-pentane. The eluent from this last 25-mL portion was collected, and its volume reduced by solvent evaporation. After 0.5 mL of acetonitrile was added, the remainder of the CH2C12and n-pentane was evaporated. Figure 7 is the chromatogram of the extract of a river water sample. This extract was prepared using the procedure described under Sample Preparation. The PAH compounds in Figures 5 , 6 and 7 were identified by their retention times by comparison with Figure 4. T h e addition of small amounts of PAH standards to a portion of each extract resulted in chromatograms with enhanced peaks a t the predicted times, corroborating their identification. We calculated the individual PAH concentrations in these samples. At least three runs were made for each sample extract. The results in Table I11 are the averages, and the uncertainties represent the range of values found. For simplicity, we have used a value of 100% for the extraction efficiency for these compounds. This introduces negligible
1318
ANALYTICAL CHEMISTRY, VOL. 51, NO.
8,JULY 1979
Table 111. PAHs in Environmental Samples
compound naphthalene fluorene phenanthrene anthracene fluoranthene pyrene benz[a]anthracene chrysene benzo[ elpyrene benzo[ b ] fluorant hene benzo[ Iz]fluoranthene benzo[a]pyrene dibenz[a, h]anthracene benzo [ghilperylene indeno [ 1,2,3-cd1pyrene
sample, ng/L plastics fly-ash wash manuf. river water 1640 i 30 1400 z 20 26 i 1 2 5 4 i 14 320 + 80 93 i 40 1280 * 320 62 i 25 9 0 ? 15 1 2 0 + 20 40+ 8 102 i- 1 4 130 i 10 100 i 20 160 i 1 5 16i 2 29 i 8 61 i 4 1 2 0 t 20 6 i 2.5 5 i 1
40
4 i 2
9 . 3 i 0.8
9.3
I
1.5 i 0.2
2.3
6
1.2
1.9i 0.3 4.5 t 1.5
6.0
0.5 9 6
13 f 2 7 i 3
16 I 1.5 10 :0.3
i
0.3
7.3
i
51
I
22
?-
80
i i
20 40
7 0 + 10 -
i
_+
O a t
t
0.1 0.5
-
error since the literature (18, 19) reports that the extraction efficiencies are greater than 90% for these compounds.
DISCUSSION As described in the introduction, within the set of PAHs in Figure 1 there are three subgroups of individual PAHs which have proved difficult to separate by liquid chromatography. Reversed-phase LC offers the best option for separation of these isomeric fused ring compounds, but CIS bonded-phase columns from different manufacturers exhibit different selectivity toward these compounds. Fox and Staley (6) reported a partial separation of B[a]A and Chy (but not of B[e]Py and B[k]Ft) on a Dupont Zorbax ODS column. Das and Thomas (13) reported B[a]A and Chy unresolved, as well as B[k]Ft, B[a]Py, and perylene unresolved on the Zorbax column. Eisenbeiss e t al. (17) and Hagenmaier et al. (11) reported that a Merck RP-18 column resolves B[b]Ft, B[k]Ft, and B[a]Py almost completely, but that B[ghi]Per and 1[1,2,3-cd]Py are only partially resolved. Krstulovic et al. (9) reported B[a]Py and perylene unresolved on either a Whatman Partisil-10 ODS column or a Waters Bondapak CI8 column. Smillie et al. (16) reported B[a]Py and B[k]Ft unresolved and B[a]A and Chy unresolved on these same two columns and also on a Varian Micropak CH column. The column we have used separates all members of these three subgroups: (a) B[a]A and Chy, (b) B[e]Py, B[a]Py, B[b]Ft, and B[k]Ft, and (c) B[ghi]Per and 1[1,2,3-cd]Py. A more thorough study of the selectivity differences between columns will be published (20). A reduced mobile phase strength is necessary to separate acenaphthene and fluorene. Trials with linear solvent gradients from 50% acetonitrile to 90 or 100% acetonitrile often resulted in little or no separation of diB[a,h]A and B[ghi]Per on the HC-ODS column. Also the resolution of this pair of compounds under the isocratic conditions of Figure 3 was found to vary on different HC-ODS columns, with no resolution observed on some columns. Consequently, we evaluated the effect of different acetonitrile concentrations on the isocratic elution of the 12 PAHs used in Figure 3. Using the k ' (capacity ratio) data from these experiments, we calculated the selectivity factor (Y for each of these PAHs relative to B[a]Py, where k $ is the capacity ratio of B[a]Py. a, = k : / k ; Figure 8 is a typical plot of a, as a function of mobile phase acetonitrile content.
0' 6C
I
70
83
3c
sc
% ACETONITRILE
Figure 8. Selectlvity factor a,r e h t i e to benzo[a]pyrene, as a function of acetonitrile content of the mobile phase (isocratic elution)
The effect of changes in the mobile phase composition on the relative selectivity of diB[a,h]A and B[ghi]Per helps to explain why linear gradients from 40 or 50% acetonitrile can result in loss of resolution between these two compounds, as compared to isocratic separation with 90 to 100% acetonitrile. On the basis of this observation, we chose a solvent program which remains a t 50% acetonitrile long enough to separate acenaphthene and fluorene, and then quickly changes to 100% acetonitrile in order to ensure the separation of diB[a,h]A and B[ghi]Per. The retention times of An, Fth, and Py are strongly affected by the length of time the column has been equilibrated at 50% acetonitrile prior to injection of the sample. A minimum equilibration time of 20 min is required and only by use of exactly the same equilibration time before each run are the retention times reproducible. Accurate reproduction of the equilibration and gradient times from run to run can be readily achieved with the microprocessor LC instruments now available. PAHs are naturally fluorescent and hence are readily monitored with a fluorescence detector, resulting in high sensitivity and specificity for these compounds. The specificity of this detector can be both an advantage and a disadvantage. Interference from most other compounds in the sample is greatly reduced, but as the number of PAHs of analytical interest is increased, the chances of them all having a common excitation and emission wavelength is reduced. We determined the excitation and emission spectra of each compound in Figure 1 by injecting the individual pure compounds. As the fluorescence peak appeared, the LC pumps were stopped and a shut-off valve closed just ahead of the injection valve. This procedure traps the peak from the column in the fluorescence detector cell so that the excitation and emission spectra can be recorded (21). For each compound, the excitation spectrum was rapidly scanned and the approximate wavelength of the largest peak was noted. The excitation monochromator was then set to this wavelength, and then the emission spectrum was recorded. The emission monochromator was then set to the wavelength of the major emission peak. and the excitation spectrum recorded. The wavelengths of peaks in these spectra are listed in Table IV. Italicized values indicate distinct, strong peaks. If the excitation and emission spectra of several compounds overlap, a compromise excitation and emission wavelength pair
ANALYTICAL CHEMISTRY, VOL. 51, NO. 8, JULY 1979
Table IV. Peak Wavelengths in Excitation and Emission Spectra excitation emission peaks peaks hex Ace F1 Phe
250 275 260 29 5
330 328 315 355,365
An
305
Fth
305
385, 405, 425, 505 340. 485
NPh
R[aIA Chy
29 5 29 5
B[eIPy B[ b ] Fth B[k] Fth B[a I PY
305 300
diB[ a.k ] A
300
B[ghi ]Per
305
375, 390, 410 390,410 366, 3 8 5 , 405 395 500 41 5 , 435 41 0 , 430, 460 395, 430, 440 340,420
I [ 1,2,3-cd] Py
305
480, 500
295
29 5 305
__
280 300 26 5 250, 290, 325 250, 350, 3 70 245, 290. 330,350 245, 290, 325 285, 340 270, 315
1319
Table V. Three Groups of PAHs and the Appropriate Compromise Excitation and Emission Wavelengths = 280 nm, A,, naphthalene acenapht hene fluorene phenanthrene
1. A,,
A, 340 340 310 360 405 480 390 410 385
290, 330 280, 330 250,305 270,295, 370, 390 292, 350
395 500 410 410
290. 370, 385 250,300, 360
420
400
500
can be chosen for detection of the compounds. The excitation and emission spectra of several of the PAHs included in this study do not overlap, indicating that a single compromise excitation and emision wavelength pair is not possible. However, from inspection of the spectra of this set of PAHs, we have divided them into two groups. Fortunately, these two groups are distinct chromatographically, all of one group eluting before any of the second group. This meant that only a single change in excitation and emission wavelength was required, occurring between Phe and An, in order to detect all of these compounds. We added a second emission wavelength change between B[ghz]Per and I[ 1,2,3-cd]Py in order to improve the detection limit for 1[1,2,3-cd]Py by a factor of 70. T h e resulting three groups of PAHs and the appropriate compromise excitation and emission wavelengths are listed in Table V. The PAHs are listed in their order of elution. The only PAH on the EPA Priority Pollutant list that is not included in Figure 1 is acenaphthylene. Acenaphthylene elutes between naphthalene and acenaphthene. It is weakly fluorescent, with excitation and emission bands located a t longer wavelengths than most of the other compounds studied. Most commercial samples of acenaphthylene contain small amounts of acenaphthene. Since acenaphthene fluoresces so much more strongly than acenaphthylene, we had difficulty in interpreting the chromatography of these two compounds. Similar problems seem to h m e affected the Sadtler Standard Fluorescence Index, which gives identical fluorescence spectra for these two compounds. The chromatogram in Figure 4 was obtained with the slits of both monochromators in the spectrofluorimeter set a t 12-nm bandpass. The detection limits can be improved by increasing the bandpass. However, impurities in the mobile phase which collect on the coiumn during equilibration at 50% acetonitrile are eluted by the gradient and these cause broad peaks near Fth and Py and a base-line shift just after Py. The magnitude of these extra peaks and base-line changes become prohibitive if the bandpass is greater than 14 nm; thus we used a bandpass of 12 min. T h e addition of PAH standards to the sample extracts served to corroborate the identification of specific peaks. Our
= 340 nm
= 305 nm, A,, = 430 nm anthracene fluoranthene pyrene benzo[a]anthracene chrysene benzo[e]pyrene benzo[ blfluoranthene benzo [ k ] fluoranthene benzo[a]pyrene dibenz[ qhlanthracene benzo [ghilperylene
2. A,,
3. he, = 305 nm, h e , = 500 nm
indeno[ 1,2,3-cd]pyrene quantitative calculations were based on the assumption that these peaks were due solely to the compounds thus identified. If a peak were due in some part to a fluorescent compound which was not, adequately resolved from the specific PAH being quantitated, then the actual concentration of that PAH would be less than we calculated. Background signals from compounds eluting near or with the compounds of analytical interest will also limit the sensitivity of this analysis. However, the selectivity of fluorescence detection, in comparison to UV detection, greatly reduces interferences of this type. In addition, interference from other fluorescent compounds is reduced by using the narrow spectral bandwidth of a monochromator as opposed to the broader spectral bandwidth of a cut-off filter (22). It is important that interference from other compounds eluting close to a compound of analytical interest be reduced as much as possible. Even with the spectrofluorimeter which we used as the detector, there was still some overlap of peaks in Figures 6 and 7 which obscured the true base line in some regions and reduced the accuracy of peak quantitation. However, the resulting uncertainties in these analyses, which are included in Table 111, correspond to very small concentrations of PAHs, 20 ng/L or less for PAHs which elute after phenanthrene. In particular, for the suspected carcinogens, B[a]A, B[b]Ft, B[a]Py, B[ghi]Per, and I[1,2,3-cd]Pyjthe observed uncertainty in quantitation in these samples was less than 8 ng/L. While this is an order of magnitude larger than the detection limits obtained with standards, it is still very low compared to other methods. Finally, it is important to note that the chromatographic analysis we have described is not restricted to the analysis of extracts of aqueous samples. It could be applied just as easily to samples prepared by other means, such as Soxhlet extraction of solids or adsorption trapping (23).
ACKNOWLEDGMENT We are grateful to Paul Strup of Battelle Columbus Laboratories for his advice on sample preparation and for supplying the extracts of two effluent water samples. We thank Ralph Riggins, also of Battelle Laboratories, for alerting us to the confusion between acenaphthene and acenaphthylene. We thank J. G. Atwood and A. Poile for numerous comments and suggestions.
LITERATURE CITED (1) "International Standards for Drinking Water", 3rd ed., World Health Organization. Geneva Switzerland, 1971, p 37. (2) B.Cathrone and M. Fielding, Proc. Anal. Div. Chem. Soc., 15, 155-158 (1978).
ANALYTICAL CHEMISTRY, VOL. 51, NO. 8 , JULY 1979 "Sampling and Analysis Procedures for Screening of Industrial Effluents r i i Pollutants", US. Environmental Protection Agency. Environment Monitoring and Support Laboratory, Cincinnati, Ohio, April 1977. H. Boden. J . Chromatogr. Sci., 14 391-395 (1976). M. Dong, D. C. Locke, and E. Fenand. Anal. Chem. 48, 368-372 (1976). M. A. Fox and S. W. Staley, Anal. Chem., 48, 992-998 (1976). C. Golden and E. Sawicki, Anal. Lett.. 9, 957-973 (1976). H. S. Hertz, W. E. May, S. N. Chesler, and B. H. Gump, Environ. Sci. Techno/., 10, 900-903 (1976). A. M. Krstulovic, D. M. Rosie. and P. R. Brown, Anal. Chem.. 48, 1383- 1386 (1976). D. W. Grant and R. B. Meiris, J . Chromatogr., 142, 339-351 (1977). H. Hagenmaier, R. Feirabend and W. J a w , Z. Wasser-Abwasser Fwscb., IO, 99-104 (1977). S. A. Wise, S. N. Chesler, H. S. Hertz, L. R. Hilpert, and W. E. M a y , Anal. Chem., 49, 2306-2310 (1977). 8 . S. Das and G. H. Thomas, Anal. Chem., 50, 967-973 (1978). A. Radecki, H. Lamparczyk, J. Grzybowski, and J. Halkiewicz, J . Chromatogr., 150, 527-532 (1978). D. E. Seizinger, Trends Flwresc. (Perkin-Elmer Gorp.), 1, 9-10 (1978).
fw P
(16) R. D. Smillie, D. T . Wang, and 0. Meresz, J. Environ. Sci. Heakb, A13, 47-59 (1978). (17) V. F. Eisenbeiss, H. Hein, R. Joster, and G. Naundorf, Chemie-Technik., 6, 227-231 (1977) [English version: Chromatogr. Newslett., 8, 8-12 ( 1978)] . (18) D. W. Ellis, Project Completion Report, University of New Hampshire, "The Analysis of Aromatic Compounds in Water Using Fluorescence and Phosphorescence", NTIS PB 212/268 (1972). (19) J. E. Wilkinson, P. E. Sbup, and P. W. Jones, Thiid Internatinxi Symposium on Potynuclear Aromatic Hydrocarbons, Columbus, Ohio, 25-27 October, 1978. (20) K. O p n , E. Katz. J. G. Atwood, and W. Slavin. manuscript in preparation. (21) W. Slavin, A. T. Rhys Williams, and R. F. Adams, J . Chromatogr., 134, 121-130 (1977). (22) K. Ogan, E. Katz, and T. J. Porro, manuscript in preparation. (23) K. Opn, E. Katz, and W. Sbvin, J . Chromtogr. Sci., 16, 517-522 (1978).
RECEIVED for review March 21,1979. Accepted April 26, 1979.
CORRESPONDENCE ~~
Anodic Oxidation of Cuprous Sulfide and the Preparation of Nonstoichiometric Copper Sulfide Sir: We have studied the anodic oxidation of cuprous sulfide in acidic medium, using a carbon paste electrode. This electrode, developed by Schultz and Kuwana (I) and French ( 2 ) ,is essentially used for the study of sparingly soluble electroactive compounds (3-6). T h e paste consists of a mixture of powdered graphite, the electroactive compound, and a pasting liquid or "binder" which is either a nonconductive liquid, such as Nujol (2-3) or a n electrolyte (4-7). Electrodes are made by packing the paste into a tube, against a glassy carbon contact, and covering the paste with a glass frit (6). T h e carbon paste electrode, a reference electrode, and a counter electrode are immersed in an electrolytic solution. Voltamperometric curves can be obtained, using the electrode in a stationary mode. T h e curves present a low residual current. T h e characteristics of the voltammograms are influenced by the nature of the pasting liquid, the amount of electroactive compound introduced in the electrode, and the potential scan rate. If the binder is an electrolyte, all of the compound can be oxidized or reduced. For low potential scan rates (for instance V/s) and small amounts of electroactive compound, the current-voltage curves are similar to those obtained in thin-layer electrochemistry; for reversible systems, they present symmetric anodic and cathodic peaks (6, 7 ) . T h e maximum of the current occurs at the standard potential of the corresponding redox system. The peak heights are given by the relation lp
n2Pvrno 4MRT
= ____
in which mo denotes the weight of solid electroactive compound introduced in the electrode, M its molecular weight, and u the potential scan rate. For high potential scan rates, or large amounts of electroactive compound, the curves are shifted toward more 0003-2700/79/035 1-1320$01 OO/O
Table I. Anodic Oxidation of Cuprous Sulfide at Different Potential Scan Rates m,, mg
0.81 0.84 0.45 0.61
u,
v.s
'
2.5 x 10-3 10-3 5 x 10''
ti
=
QM/
0.95 1.02
96500m" 1.95 2.02
0.55
2.03
Q,a C
0.72 1.97 Q represents the total area between the current-voltage curve and the residual current; Q is determined b y weighing the corresponding recording paper. positive or negative potentials because of the ohmic drop, but the half sum of the anodic and cathodic peak potentials remains constant and equal to the standard potential of the corresponding redox system (7). The electrochemical behavior of cuprous sulfide has been studied by means of a carbon paste electrode containing a 1 M sulfuric acid solution as the binder. For scan rates higher than 5 X V/s, the current-voltage curve is characterized by one anodic peak A (Figure 1). The area under the curve indicates a two-electron transfer. This peak is related to the oxidation of cuprous sulfide, to cupric sulfide and cupric ions (6) according the reaction: Cu,S
-
CuS
+ Cu2+ + 2 e
With decreasing potential scan rates, the peak A splits gradually into five different peaks AI . . . A, (Figure 2); the total area under the curve remains constant and always corresponds to a two-electron transfer (Table I). For v 5 2.5 x IO4 Vas-', the peaks Al . . . Ab are well defined: the oxidation of cuprous sulfide to cupric sulfide then proceeds via the formation of a succession of stable phases Cuz&3 according to the reactions:
C 1979 American Chemical Society
