Anal. Chem. 1982, 54, 2571-2576
2571
Immobilized Enzyme System for the Conversion of Benzo[ a ]p:yrene to Fluorescent Metabolites Sandra N. Walters, Arthur P. D’SIIva, and Velmer A. Fassel“ Ames Laboratory, Iowa State Universify, Anies, Iowa 5001 7
The enzyme aryl hydrocarbon hydroxylase (AHH) immobilized on a polyacrylamide suipport poly(acryiamide-co-N-acryioxysucclnlmide) has been utilized for the emzymatlc conversion of benzo[a]pyrene (B[a]P) to 3-hydroxybenro[a]pyrene (3OH-B[a]P), a fluorescent metabolite. Because the procedure mimlcs reactlons occurring in mammalian tlssues, the measurement of the fluorescence of 3-OH-B[e]P offers promise as a means of assesslng the potentlal mutagenicity of B[a]P occurring In complex mlxtures.
The ubiquitous carcinogen benzo[a]pyrene (B[a]P) is often implicated in respiratory carcinogenesis and is the target compound most frequently determined in assessing the carcinogenic potential of samples suspected to cause lung cancer. However, it has been generally recognized that the carcinogenicity of a sample cannot be unequivocally correlated with either the B[a]P content or the fraction that contains all the polynuclear aromatic hydrocarbons (PAHs), since the ultimate carcinogenicityof samplles is largely controlled by synergistic effects from sample concomitants. It ha!$ been observed that cigarette smoke tar contains substancesthat enhance or inhibit carcinogenic activity ( l ) .Even for the single class of compounds, the alkylated and multialkylated naphthalenes, which are found in cigarette smoke tar,both activating and inhibiting effects on the carcinogenicity of B[a]P have been observed (2). The presence of particulates, asbestos, or hematite have also been shown to increase the mutagenic effect of B[a]P (2). The above studies indicate that the mutagenicity of a sample cannot be established by the determination of B[a]P by chemical methods alone. Thus, there is! a necessity for the development of analytical methods, preferably biochemical, which mimic in vitro the in vivo effects that mediate the mutagenicity of B[a]P, especially in the presence of concomitants. Ideally, the total sample should ble directly subjected to such tests. Most PAHs, including B[a]P, are biologically inert and require metabolic activation to exert their mutagenic and carcinogenic effects (3). The first and major enzyme in the main metabolic pathway of PAHs is argl hydrocarbon hydroxylase (AHH), a multienzyme complex present in most mammalian tissues. A variety of foreign compounds, including many PAHs, are known to increase AHH metabolic activity through the process of enzyme induction (3,4). B[a]P, and other PAHs, are metabolized by AHH tal dihydrodiols, phenols, quinones, and epoxides (5). Several of these metabolites, usually the reactive epoxides, are more carcinogenic than the parent compound ( 3 , 4 , 6 ) .One of the principal metabolites produced from B[a]P by AHH is 3-hydr1oxybenzo[a]pyrene (3-OH-B[a]P),a moderately intense fluorophor with characteristic green emission a t 525 nm on excitation at 430 nm. Because the fluorescence of the precursol. molecule B[a]P is usually measured a t 403 nm on excitation at 380 nm, the metabolite, 3-OH-B[a]I), can be readily measured without interference from its parent compound. However, interferences from AHH metabolites of unknown constituents present in environmental samples may occur because the fluorescence properties of the other imetabolites may be similar to those 0003-2700/82/0354-2571$01.25/0
of 3-OH-B[a]P. Fluorescence measurements of the metabolic product 3-OH-B[a]P are routinely performed to measure the enzyme activity and concentration of AHH (7,8). This determination of AHH enzyme activity present in an organism, e.g., rat, provides a sensitive bioassay for many carcinogens based on the increase in AHH activity due to enzyme induction following treatment of the organism with a carcinogen. It has been reported that 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD), an extremely toxic compound, can be detected at mol levels by such an assay (9). In vivo treatment by carcinogens such as 3-methylcholanthrene,or pesticides such as N-nitrosocarbonyl (IO), increase and alter the metabolism of B[a]P by their effects on AHH. Other compounds, such as the food additive propyl gallate (II), can act directly, in vitro, on the enzyme AHH to alter its metabolism of B[a]P. Thus, measurement of the metabolic conversion by AHH of a carcinogen, such as B[a]P, present in a mixture may provide an assessment of the ultimate mutagenicity and carcinogenicity of the sample. Conventional AHH assays are usually performed in liquid media under carefully controlled experimental conditions because the enzyme preparations utilized in such assays are sensitive to high temperature, changes in pH, microbial attack, and hydrolysis. Because of such effects, reuse of expensive enzyme preparations is rarely possible. In contrast, enzyme stabilization by immobilization on solid supports has been found to circumvent such limitations and has led to the wider utilization of the unique properties of enzymes in industry and analytical chemistry (12, 13). Specifically, a drug detoxifier system incorporatingimmobilized liver microsomes has been reported (14). In this paper we report on the use of an immobilized enzyme assay for B[a]P, which is different from the conventional AHH assay. In our assay, the production of the B[a]P metabolite, 3-OH-B[a]P, by an immobilized AHH enzyme preparation is measured by its fluorescence. The conversion of B[a]P to 3-OH-B[a]P by our immobilized enzyme system has been characterized. In addition, the B[a]P metabolites produced from a mixture of six PAHs and from a carbon black extract by immobilized AHH have been measured. This method may be useful for assessing the carcinogenic potential of real samples such as carbon black, which is widely distributed in the environment due to its use in such diverse products as automobile tires and photocopy processes (15, 16).
EXPERIMENTAL SECTION Reagents and Chemicals. AU chemicals were of reagent grade. The 3-OH-B[a]Pwas provided by David Longfellow of the National Cancer Institute. Carbon lampblack, lot 702959,was from Fisher Scientific Co. (Pittsburgh, PA). Sprague-Dawleyderived rats were purchased from SASCO (Omaha, NE). Microsomal Preparation. Male rats, each weighing approximately 100 g, were given daily intraperitoneal injections of 3-methylcholanthrene(25 mg/kg body weight) in corn oil for 4 days prior to preparation of liver microsomes according to the method of Kinoshita et al. (17).The rats were killed by decapitation and each liver (4-5 g) was quickly removed and homogenized in 30 mL of 0.25 M sucrose and 0.05 M tris(hydroxymethy1)aminomethane (Tris) buffer, pH 7.5,with a Potter-Elvehjem homogenizer. The liver homogenate was centrifuged at 0 1982 American Chemical Society
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ANALYTICAL CHEMISTRY, VOL. 54, NO. 14, DECEMBER 1982
lOOOOg for 20 min. The resulting supernatent was then centrifuged at 1OOOOOg for 60 min. The final pellet consisting of the liver microsomal fraction was solubilized with a Dounce homogenizer in 0.25 M sucrose, 0.05 M Tris buffer, pH 7.5, and was stored at -40 "C. The protein concentration was determined by the method of Lowry et al. (18). AHH-PAN Preparation. Poly(acry1amide-co-N-acryloxysuccinimide)(PAN) was prepared by the method of Pollak et al. (19). AHH-PAN batches of 20 plates were prepared by solubilizing 0.8 g of PAN (with 450 pM ester/g of PAN) in 3.2 mL of 0.03 M MgC12,0.3 M Hepes buffer, pH 7.5, containing 1 mg of benzo[e]pyrene (B[e]P) and 1mg of NADPH to protect the active site of the enzyme AHH during immobilization. While the mixture was stirred, 0.3 mL of 0.5 M triethylenetetramine (TET) (distilled, aqueous) was added to begin cross-linking and 1 min later 0.4 mL of the microsomal preparation containing 6-12 mg of protein was added. The resulting solution was quickly pipetted in 0.2 mL aliquots into petri plates (60 mm diameter) and spread into a uniform thin layer using a bent glass rod. After addition of TET, the gel hardened in 10 min and polymerization was complete within 60 min. Each petri plate of AHH-PAN was washed two times with pH 7.5 buffer containing 50 mM Hepes, 10 mM MgC12,and 10 mM lysine, to remove unreacted ester groups, and then washed two times with pH 7.5 buffer containing 50 mM Hepes and 10 mM MgC12and was stored at 4 "C. AHH-PAN Assay for Determination of B[a]P. The AHH-PAN assay method for B[a]P determination was based on the enzyme activity assay for AHH by Dehnen et al. (8). Unless otherwiseindicated, 1.8mL of incubation solution containing 0.24 pmol of NADPH, 3 pmol of MgC12,and 1mmol of Tris at pH 8.2 was added to each AHH-PAN petri plate containing 300 to 600 pg of immobilized microsomal protein. The metabolic reaction was initiated by the addition of B[a]P (in methanol) to the incubation solution in an AHH-PAN petri plate. The conversion of B[a]P to its metabolites by AHH-PAN occurred at 37 "C in an incubator during a 40-min period. The reaction was stopped by the addition of 0.2 mL of 1M NaOH with 10% (w:v) Triton X-100, which denatures the enzyme, and 1% EDTA, which chelates the cofactor Mg2+.The metabolite 3-OH-B[a]Pfluoresces in the anionic form and its solubility, and thus fluorescence, is enhanced by the detergent. This final metabolite solution was transferred to test tubes, centrifuged to ensure removal of turbidity, and the fluorescence spectra were then recorded. The term "incubation solution" refers to the buffered solution containing NADPH, Mg2+and the substrate B[a]P which is reacted with AHH-PAN during the incubation period. The "final metabolite solution" refers to the solution which is measured spectroscopically and is comprised of the incubation solution containing the metabolites produced by AHH-PAN with the addition of 1M NaOH with Triton X-100 and EDTA. Carbon Black Extract Preparation. A Soxhlet extraction of 22.3 g of carbon black with approximately 300 mL of chlorobenzene (distilled) was performed for 63 h (20). The chlorobenzene was removed with a rotary evaporation apparatus and the sample was dissolved in 200 mL of methanol. AHH-PAN Assay for B[a]P Metabolites in Carbon Black Extract. This procedure was based upon the AHH enzyme assay method of Nebert and Gelboin (7). Samples of carbon black extract (CBE) were added to AHH-PAN plates with incubation solution containingNADPH, Mg2+,and buffer as described above. After the 40-min incubation period, the solutions were transferred from AHH-PAN plates to test tubes. The metabolites were extracted into 4 mL of hexane-acetone (33, v:v). Three milliliters of the organic phase was transferred to a new test tube and the metabolites were then extracted into 1.2 mL of 1 M NaOH. The fluorescence spectra of the metabolites present in the NaOH solution were recorded. Sample blanks were prepared by adding carbon lampblack samples to AHH-PAN plates prepared from heat-inactivated microsomes and otherwise following the same assay procedure. For comparison purposes the actual concentration of B[a]P in the carbon black extract was determined independently by laser-excited Shpol'skii spectrometry (LESS) with deuterated B[a]P as an internal standard (21). Fluorescence Spectrometry. The fluorescencespectra were measured with a fluorometer assembled in this laboratory. The components and experimental conditions are indicated in Table
Table I. Fluorescence Spectrometer Components excitation source
200-W mercury lamp enclosed in a monochromator illumination housing (Oriel Model 7340) powered by an appropriate power supply (Oriel Model 8600) excitation monochromator 0.25 m, f/3.6 scanning monochromator, 2 mm slits (Jarrell-Ash Model 82-420) filters Corning 666 at the excitation side of the sample and a Schott KV-470 at the emission side of the sample emission monochromator 0.220 m focal length, f / 4 scanning monochromator with holographic grating, 0.26 mm slits (Spex . - Model 1670) photometer Spex (Model DPC2) photometer with a cooled photomultiplier (HTV-R955 PMT) and an appropriate strip chart recorder (Houston Instruments Model Model 4900)
WAVELENGTH (nm)
Figure 1. Fluorescence spectra of (A) 98 ng of 3-OH-B[a]P. (B) Metabolites produced from 532 ng of B[a]P by free rat liver microsomes (75 pg of protein). (C) Metabolites produced from 532 ny of B[a]P by AHH-PAN (450 pg of protein). The lower spectral line indicates background fluorescence of the sample.
I. The fluorescence intensity was measured as the peak height at 525 nm and was corrected for background fluorescence. The excitation wavelength was 430 nm.
RESULTS AND DISCUSSION B[a]P Metabolite Spectra. The fluorescence spectra of 3-OH-B[a]P and the metabolites produced from nanogram amounts of B[a]P by both free and immobilized microsomes containing AHH are shown in Figure 1. The similarity in the fluorescence spectra suggests that 3-OH-B[a]P was the major metabolite being determined in the assay, although the metabolite 9-hydroxybenzo[a]pyrene (9-OH-B[a]P) may make a small contribution to the fluorescence spectra. The major metabolite produced by rat liver AHH from B[a]P is 3-OHB[a]P (35-60%), while 9-OH-B[a]P, which has a similar spectrum to 3-OH-B[a]P, constitutes only 3-12% of the metabolites formed (22,23). Because the analytical calibration curve for each assay was derived from fluorescence spectra of B[a]P metabolites produced by AHH-PAN rather than from spectra of 3-OH-B[a]P standards, the assay in principle provides a measure of B[a]P metabolism and may not provide a quantitative determination of the B[a]P precursor in the sample. The background fluorescence spectrum shown in
ANALYTICAL CHEMISTRY, VOL. 54, NO. 14, DECEMBER 1982
Figure 1was obtained from a B[a]P-free incubation solution with AHH-PAN processed in the same way as the other samples. The Raman peak of water at 520 nm in the background spectrum impoises a limit on the ultimate powers of detection achievable. Bemuse a good correlation between peak height and area was obiserved, the former was measured at 525 nm for all of the data reported below. Properties of the Metabolite 3-OH-B[a]P. The fluorescence intensity of the metabolite 3-OH-B[a]P was observed to be a linear function of concentration between 30 and 240 ng/mL, as has been reported previously (8). The final solution cointaining 3-OH-B[a]P produced from 133 ng of B[a]P by the ,4HH-PAN assay was stored at room temperature under constant (24 h) fluoreiscent room lighting. Over 70% of the initial 3-OH-B[a]P fluorescence remained after 2 weeks, and 4OC%remained after 33 days. Photodegradation of 3-OH-B[a]Pdoes not present a serious problem in this assay since it iei a slow process ;and the effects are negligible over the time period required for the assay. The absolute amouint of 3-OH-B[a]P recovered from AHH-PAN plates was determined by adding known amounts of 3-OH-B[a]P in incubation solutions to AHH-PAN plates prepared from heat-inactivated microsome13(300 pg of protein/ plate). Enzyme inactivation was necessary to prevent metabolism of 3-OH-B[a]P by AHH, although the loss of 3-OHB[a]P due to further metabolism by AHH was found to be less than 10% in other experiments with 3-OH-B[a]P as the substrate or analyte in the AHH-PAN away. To determine the recovery from the plate, we compared the fluorescence spectra of the final incubation solution resulting from an AHH-PAN assay (with inactive AHH) of known amounts of 3-OH-B[a]P to the spectra of reference solutions containing the same known amounts of 3-OH-B[a]P. The mean recovery of 3-OH-B[a]P was 30%, and varied with 3-OH-B[a]P concentration, being greater at higher concentrations. Most of the loss was probably due to nonspecific binding of 3-OH-B[alp to the lipid membrane and proteins of the microsomes (24). This nonspecific binding is saturable, which would account for the greater recovery at higher 3-OH-B[a]P concentrations. The recovery of 3-OH-B[a]P is similar for free microsomes and AHH-PAN, although some nonspecific binding of 3-OH-B[a]P 'to the PAN matrix itself may occur. Analytical Calibrations. As noted earlier the fluorescence intensity of 3-OH-B[a]F' produced by the! AHH-PAN assay was a linear function of B[a]P substrate concentration. Even though variations in enzyme activity between AHH-PAN batches occurred,the analytical calibration curve of metabolite fluorescence vs. B[a]P concentration with pure B[a]P as the substrate for a given AHII-PAN batch was1 found to be linear. In these and subsequent experiments, the difference between duplicate samples was usually less than 10% of the mean, although differences as high as 20% or 25% occasionally occurred. Each of the experiments characterizing the AHHPAN assay method was performed in duplicate at least three times for different AHH-PAN batches. Similar results were obtained even though the variability between batches was sometimes greater than desired. Reaction Rate. The apparent catalytic and kinetic p r o p erties of AHH were chaniged by immobilization, as has been extensively documented for other immobilized enzyme systems (25). As shown in Figure 2, the metabolism of B[a]P, within the concentration range of 20-300 ng/mL normally used in this assay, was complete in 30-40 min. Complete metabolism of B[a]P to 3-OH-B[a]P by free microsomes occurred in less than 10 min under the same assay conditions. In another experiment, the AHH enzyme activity was determined with limiting amounts of microsomal protein and excess amounts of B[a]P (4.4 mM), NADPH (0.4 mM), and
b
2573
1 0 -
e b
-
-
1
05
I I5
I O
INCUBATION
TIME
1 2 0
lh)
Flgure 2. Effect of Incubation time on B[a]P conversion to 3-OKB[a]P by AHH-PAN for 110 ng of B[a]P and 600 pg of microsomal pro-
teinlpiate.
7.0
8.0 PH
Flgure 3. Effect of incubation pH on B[a]P metabolism by AHH-PAN for 110 ng of B[a]P and 300 pg of microsomal protein/plate.
Mg2+ (2.25 mM) during a 10-min incubation period. The amount of microsomal protein was 10-60 pg for free microsomes and 300 pg for AHH-PAN. The specific activity of the enzyme AHH was 101 pmol of 3-OH-B[a]P produced min-' mg-l of protein for free microsomes with a standard deviation (SD) = 18, and was 24 pmol m i d mg-' for immobilized microsomes with SD = 4.5. A 25% yield of enzyme activity upon immobilization is acceptable for a multienzyme complex such as AHH. Determination of the kinetic constants of AHH in free microsomes is difficult, since Michaelis-Menten type kinetics may not necessarily occur due to nonspecific binding of B[a]P to nonenzymatic sites, low B[a]P solubility in water, rapid depletion of B[a]P, and further metabolism of 3-OH-B[a]P (24). Immobilization of AHH caused a decrease in the apparent reaction rate and a decrease in the apparent affinity for the substrate, which is presumably due to steric distortion or loss of AHH upon immobilization (25) and diffusion of substrate and metabolic products through the PAN matrix (26).
Effect of pH. The optimal pH for the conversion of B[a]P to 3-OH-B[a]P by AHH in free microsomes has been reported to be 7.5 (7). This value was shifted to 8.2 upon immobilization, as shown in Figure 3. A change in the pH profile of enzyme catalytic activity after immobilization commonly occurs (25,27)and is thought to result from an alteration in the microenvironment caused by the support, e.g., presence of charged groups, or an alteration in enzyme conformation
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ANALYTICAL CHEMISTRY, VOL. 54, NO. 14, DECEMBER 1982
Table 11. Effect of TCPO and Norharman on Conversion of B[a]P to 3-OH-B[a]Pa r el 3-OH-B[a]P enhancement fluorescence factor
sample untreated t TCPO t norharman t TCPO and norharman
u IO
30
20
40
~
I
l
I
i
I
I
I
' 01
160
-
I
,
00
c
1.00
1.43 1.47 2.87
a t B[a]P (266 ng) was metabolized by AHH-PAN (300 fig of proteinlplate) during a 45-min incubation period with additions of 2 mM TCPO or 0.6 mM Norharman.
50
c
l
5.78 8.28 8.52 16.61
l
I
l
I
l
I
,
-
m at 4* C
20
40
60
80
100
% H20
2
6
IO
14
STORAGE
18
TIME
22
26
30
34
Flgure 6. AHH-PAN metabolism of 300 ng of B[a]P in incubation solutions containlng varying propottbns of water and methanol/giyceroi (1: l), with appropriate amounts of NADPH and MgCi2. The solution was buffered to an apparent pH of 8.2 with Tris. The plates each contained 300 pg of microsomal protein.
idoys)
Figure 5. Enzyme activity of AHH-PAN wRh storage at (0)0 "C, (I) increase the AHH enzyme activity. Altering AHH Metabolism. As mentioned previously, 4 "C, or (A) 23 "C.
upon immobilization which changes its catalytic properties. Incubation Temperatures. The AHH activity of free microsomes is readily destroyed by heat with a reported half-life of 30 min at 37 "C and 5 min at 45 "C (7). Thermal stability is often increased by immobilization (28). Our observations (see Figure 4) of AHH-PAN indicate no significant loss in enzyme activity at incubation temperatures between 23 OC and 37 "C. Thus incubationsat room temperature could be performed with little loss in activity. Approximately4-fold losses in activity were observed at 4 "C, presumably caused by a reduction in reaction rate, and at 54 "C, presumably from heat deactivation. AHH-PAN Stability. Enzyme stabilization by immobilization does not always occur; destabilization or no change in enzyme stability is just as likely (28). Fortunately, enzyme stabilization did occur with the AHH-PAN system as shown in Figure 5 for three batches of AHH-PAN, each stored for various time periods at one of the indicated temperatures. In this figure, the enzyme activity is expressed as a percentage of the AHH activity, measured by 3-OH-B[a]P fluorescence, on the day of immobilizatiion (day 0), with mean values of duplicate runs indicated. It is seen that most of the enzyme activity was retained through 2 weeks with storage at 0 "C or 4 "C. Enzyme activity was rapidly lost during storage at room temperature. The initial increase in enzyme activity after storage at 0 "C is probably not significant, although freezing may disrupt the PAN structure and free some of the microsomes that were inactivated by immobilization and thus
B[a]P is metabolized by AHH to compounds other than 3OH-B[a]P. Increases in the proportion of 3-OH-B[a]Prelative to the total metabolites produced from B[a]P by AHH should increase the detection limits of the assay. Treatment of the rats with 3-methylcholanthrene increased the AHH enzyme activity in the liver microsomal fraction 20-fold while epoxide hydrase enzyme activity was increased by less than 50% (4). TCPO (1,2-epoxy-3,3,3-trichloropropane), an epoxide hydrase inhibitor (29),was found to increase the amount of 3-OH-B[a]P produced in the AHH-PAN assay system (Table 11). The increased 3-OH-B[a]P production was presumably due to a decrease in the formation of dihydrodiols from the epoxide that is catalyzed by epoxide hydrase and a concomitant increase in the formation of phenols. Norharman (@carboline, SH-pyrido[3,4-b]indole),which has been reported to increase the proportion of 3-OH-B[a]P produced from B[a]P by noncompetitivebinding to the cytochrome P-450 portion of AHH (30),also increased the amount of 3-OH-B[a]Pproduced from B[a]P in our assay system. As shown in Table 11, the effects of these two compounds on 3-OH-B[a]P production from B[a]P by AHH-PAN were additive, which was expected since each compound acts on a separate enzyme. The additive effects are described by a rather complex mathematical expression based on enzyme kinetics rather than simple addition. AHH-PAN Assay in Organic Solvents. Because many PAH compounds have a low solubility in aqueous solutions, the ability to perform these assays for PAHs occurring in organic solutions would increase the applicability. The possibility of using an organic-aqueous incubation solution is
ANALYTICAL CHEMISTRY, VOL. 54, NO. 14, DECEMBER 1982
L 5 500
fi00
500
WAVELENGTH
600
500
2575
60(
(nm)
Flgure 7. Fluorescence slpectra of the AHH-IPAN metabolites of (A) 175 ng of B[a]P, (B) 180 ng of B[a]P in a mixture of benzo[e]pyrene (200 ng), 3-methylcholanthrene (160 ng), fluoranthene (200 ng), and anthracene (200 ng), and (C) background fluorescence.
illustrated in Figure 6. The proportion of H 2 0 may be decreased to less than 4056 before any appreciable loss of AHH activity occurs. This retention of enzymat,icactivity in organic solvents is not surprising for a membrane enzyme which actually requires phospholipid for activity ((31).The structural integrity of PAN is evidently maintained with this solvent mixture, even though the disruption of polyacrylamide structure in many orgamic solvents has been reported (32). Control experiments demonstrated that the fluorescence of known amounts of 3-OIH-B[a]P standards did not vary with the proportion of H 2 0 present in the solution. Assay of PAH Mixture. Figure 7 shows the fluorescence spectra of AHH metabolites produced by the AHH-PAN assay of a sample containing only B[a]P and a sample containing B[a]P plus four other PAHs. The observed fluorescent peaks are in quantitative concordance with the concentration of B[a]P, suggesting that the metabolite spectra of the other PAHs did not interfere riignificantly and that the other PAHs did not have a measurable effect on enzyme activity. Only the fluoranthene metabolites produced a weak but quantitatively insignificant fluorescence under the experimental conditions. Assay of CBE. The fluorescence spectra of metabolites produced by AHH-PAN from pure B[a]F' and from CBE are shown in Figure 8. The AHH metabolite fluorescence of CBE was equivalent to 95 Mg/mL of B[a]P in CBE. The B[a]P concentration of CBE was equivalent to 9.2 pg/mL when determined independently by LESS. The LESS determination of B[a]P was perforimed three times at 3-4 CBE dilutions with two different refereince solutions of B(alp. The increased fluorescence in the finall metabolite solution was not caused by a component of the CBE itself as shown by spectrum E. T o investigate the cause of the increased metabolite fluorescence of CBE, WE! performed a standard additions experiment. A series of four reference samples containing individually 625, 1250, 25100,and 3100 ng of B[a]P were metabolized by AHH-PAN, The plot of the resulting metabolite fluorescence vs. the B[a1P concentration produced the expected linear response when the data were treated via a least-squares regression. Another series of samples containing the same amounts of B[a]P were spiked with identical amounts of CBE, which by LESS analysis contained 92 ng of B[a]P. These samples presumably contained 717, 1342, 2592, and 3192 ng of B[a]P, respectively, and were also metabolized by AHH-PAN. The plot of the resulting metabolite fluorescence vs. B[a]P concentration showed a linear response, with a slope which was the same as the unspiked reference samples (2.41 X for the spiked samples and 2.26 X for the unspiked samples). However, the lines represent a spike addition of 890 ng of B[a]P rather than 92 ng, indicating a considerable enhancement of fluorescence signal. Because
WAVELENGTH.
(091
Figure 8. Fluorescence spectra of (A) 3-OH-B[a]P (325 ng), (B) metabolites of B[a]P (3100 ng) produced by AHH-PAN, (C) sample blank resulting from 0 substrate and AHH-PAN, (D) metabolites of CBE (230 ng of B[a]P) produced by AHH-PAN, and (E) CBE sample blank resulting from an Incubation of CBE (230 ng of B[a]P) wlth AHH-PAN prepared from heat-inactivated mlcrosomes.
CBE constituents causing enzyme activation of AHH would be expected to cause a significant increase in the slope of the fluorescence v8. B[a]P concentration curve of the spiked samples, the observation of parallel lines for the addition experiment strongly suggests that interfering metabolites formed from non-B[a]P components of the extract caused the enhancement. This deduction is supported by the observation that a change of the excitation wavelength from 430 nm to 460 nm, which is an excitation maximum of 3-OH-B[a]P, caused a 10% increase in metabolite fluorescence from pure B[a]P samples and a 30% decrease in metabolite fluorescence from CBE samples. Since the final metabolic solution was extracted into hexane/acetone and then into 1M NaOH, the interference is probably caused by a phenolic metabolite. However, effects of sample concomitants on enzyme activity of AHH or other microsomal enzymes by mechanisms such as increasing the proportion of 3-OH-B[a]P produced from B[a]P (as was caused by Norharman and TCPO) may also have contributed to the increased CBE metabolite fluorescence observed. Interferences caused by other fluorescent metabolites produced from environmental samples may therefore be a limitation of this method as described. The fluorescence interferences may be overcome by application of other spectrometric techniques, such as synchronous spectrometry or Shpol'skii spectrometry. This drawback could be exploited by using the total metabolite fluorescence produced by AHH from an environmentalsample as a measure of the compounds present that are metabolized and are potentially carcinogenic. Such a measurement would require validation by correlating the AHH total metabolite fluorescence to other carcinogen tests. The immobilized AHH enzyme assa37 is a fairly simple and rapid technique that is potentially useful for assessing environmental samples for their carcinogenic potential.
LITERATURE CITED (1) Schmeltz, I.; Tosk, J.; Hllfrlch, J.; Hlrota, N.; Hoffman, D.; Wynder, E. L. "Carcinogenesis"; Jones, P. W., Freudenthal, R. I., Eds.; Raven Press: New York, 1978; p 47. (2) kakowlcz, J. R.; Hylden, J. L.; Englund, F.; Hldmark, A,; McNamura, M. Polynuclear Aromatic Hydrocarbons"; Jones, P. W., Leber, P., Eds.; Ann Arbor Science Publishers: Ann Arbor, MI, 1979; p 835. (3) Gelboln, H. V. Rev. Can. Biol. 1972, 37, 39-60. (4) Thorgelrsson, S. S.; Nebert, D. W. Adv. Cancer Res. 1977, 25, 14% 193. (5) Selklrk, J. K.; Croy, R. G.; Whltlock, J. P., Jr.; Gelboln, H. V. Cancer Res. 1975, 35, 3651-3655. (6) DePlerre, Joseph W.; Ernster, L. Biochim. Blophys. Acta 1077, 149-1 86. (7) Nebert, D. W.; Gelboin, H. V. J. Blol. Chem. 1966, 253, 6242-6249.
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Anal. Chem. 1982, 54, 2576-2582
(8) Dehnen, W.; Tominges, R.; ROOS, J. Anal. Biochem. 1973, 53. 373-383. (9) Niva, A.; Kumakl, K.; Nebert, D. W. Mol. Pharmacol. 1975, 1 1 , 399-408. (10) Beraud, M.; Galliard, S.; Derache, R. Chem.-Biol. Interact. 1980, 31, 103-112 .- - . .-. (11) Yang, C., S.; Strickhart, F. S. Biochem. Pharmacol. 1974, 23, 3129-3138. (12) Ngo, T. T. Int. J . Biochem. 1979, 1 1 , 459-485. (13) Wlseman, A. J . Chem. Techno/.Biotechnol. 1980, 30, 521-529. (14) Cohen, W.; Barlcos, W. H.; Kastl, P. R.; Chambers, R. P. "Methods in Enzvmoioav"; Mosbach. K., Ed.; Academic Press: New York, 1976; Vol:44, pip 319-328. (15) Lee, M. L.; Hltes, R. A. Anal. Chem. 1976. 48, 1890-1693. (16) Rosenkranz, H. S.; McCoy, E. C.; Sanders, D. R.; Butler, M.; Kiriazides, D. K.; Mermeistein, R. Science 1980, 209, 2039-2043. (17) Kinoshita, N.; Shears, B.; Gelboln, H. V. Cancer Res. 1973, 33, 1937-1944. (18) Lowry. 0. H.; Rosebrough, N. J.; Farr, A. L.; Randall, R. J. J . Biol. Chem. 1961. 193,285-275. (19) Poilak, A.; Blumenfeld, H.; Wax, M.; Baughn, R. L.; Whitesides 0. M. J . Am. Chem. SOC.1980, 102,8324-8336. (20) Giammarise, A. T.; Evans, D. L.; Butler, M. A,; Murphy, C. 6.; Kiriazides, D. K.; Marsh, D.; Mermelsteln, R. "Improved Methodology for Carbon Black Extraction", Sixth International Symposium on Polynuclear Aromatlc Hydrocarbons, Columbus, OH, Oct 1981. (21) Yang, Y.; D'Silva, A. P.; Fassel, V. A. Anal. Chem. 1981, 53, 894-899. (22) Yang, S. K.; Selkirk, J. K.; Plotkin, E. V.; Gelboin, H. V. Cancer Res. 1975, 35,3842-3850.
(23) Golan. M. D., Bucker, M.; Schmassmann, H. U.; Raphael, D.; Jung, R.; Blndel, U.; Brase, H. 0.; Tegtmeyer, F.; Frledberg, T.; Lorenz, J.; Stasiecki, P.; Oesch, F. Drug Metab. Dlspos. 1980, 8 , 121-126. (24) Camps, J.; Razzouk, C.; Roberfrold, M. B. Chem.-Biol. Interact. 1977, 16, 23-38. (25) Berezin, I . V.; Kllbanov, A. M.; Martinek, K. Russ. Chem. Rev. 1975, 44 (l), 9-25. (26) Thomas, D.; Brown, G. "Methods in Enzymology"; Mosback, Klaus, Ed.; Academlc Press: New York, 1976; Vol. 44, pp 901-929. (27) Goldman, R.; Goldstein, L.; Katchalski. E. I n "Biochemical Aspects of Reactions on Solid Supports"; Stark, G. R., Ed.; Academic Press: New York, 1971; pp 1-78. (28) Klibanov, A. M. Anal. Biochem. 1979, 93, 1-25. (29) Oesch, F.; Jerlna, D. M.; Day, J. W.; Rice, J. M. Chem.-Biol. Interact. 1973, 6, 189-202. (30) Tadahlko. F.; Matsuyama, A.; Nagao, M.; Suglmura, T. Chem.-Blol. Interact. 1980, 32, 1-12. (31) Levin, W.; Ryan, D.; West, S.; Ayh, L. J. Biol. Chem. 1974, 249, 1747-1754. (32) Ingalls, R. G.; Squlres, R. 0.; Butler, L. G. Biotechnol. Bioeng. 1975, 17, 1627-1837.
RECEIVED for review June 7, 1982. Accepted September 20, 1982. This research was suported by the U.S. Department of Energy, Contract No. W-7405-Eng-82,Office of Health and Environmental Research, Physical and Technological Studies, Budget Code GK-01-02-04-3.
Determination of Transferable Hydrogen in Coal Liquids by Mass Spectrometry J. T. Swansiger," H. T. Best, D. A. Danner, and T. L. Youngless Gulf Science and Technology, Pittsburgh, Pennsylvania 15230
Two low-resolution mass spectral (LRMS) group type analyses have been callbrated to quantltate transferable hydrogen and to more accurately characterlre the aromatlc and hydroaromatlc specles In coal llqulds. The analysls for llght coal llqulds (50-300 "C) reports 20 aromatlc and hydroaromatic types lncludlng phenols and dlhydroxybenrenes, whlle the analysls for heavy coal llqulds (300-500 "C) reports 30 aromatlc and hydroaromatic species. The LRMS transferable hydrogen values were shown to be In good agreement wlth 13C NMR values for a serles of coal llquld dlstlllates and process oils.
Hydrogen donors are of fundamental importance in coal liquefaction. It has been shown that hydrogen donors in coal liquefaction are not necessarily the classical hydroaromatics and that coal free radicals can abstract hydrogen from many sources including naphthenes, alkyl aromatics, and dissolved hydrogen (1,2). However, hydroaromatics are of most interest due to their comparatively high reactivity as hydrogen donors. The 13C NMR technique of Seshadri et al. (3), which determines transferable hydrogen by using the hydroaromatic region of the spectrum, was used as the reference technique. The LRMS technique uses the percent transferable hydrogen for each hydroaromatic type to calculate the total transferable hydrogen. Both LRMS analyses are based on our previous work, which described a 17-component group type analysis for coal liquids ( 4 ) . However, attempts to use the early calibration matrix to calculate transferable hydrogen indicated that the LRMS data were consistently high with respect to 13C NMR. Since we considered 13CNMR to be more struc0003-2700/82/0354-2576$0 1.25/0
turally sensitive than the mass spectral group type analysis, agreement with 13C NMR data for transferable hydrogen should be indicative that appropriate calibration compounds and characteristic ions have been selected for the LRMS analyses. An advantage of the group type analyses is the detailed compositional data reported for aromatic and hydroaromatic types. By fitting the LRMS data to 13C NMR data, we feel that we have resolved most of the significant series overlaps and improved structural accuracy of the LRMS analyses.
EXPERIMENTAL SECTION Sixteen narrow boiling fractions of a coal liquid product, covering a boiling range of 42-482 "C, were characterized by GC/MS, LRMS, and 13C NMR. The coal liquid was obtained from SRC-I1 processing of Powhatan No. 5 mine coal. The material fractionated was a blend of debutanizer bottoms and process solvent. Additional physical, chemical, and thermodynamic properties for these fractions are contained in a comprehensive report by Gray (5). The various characterization data for these fractions were used to support selection of calibration components in the LRMS group type analyses. Low-Resolution Mass Spectrometry. The LRMS calibration data and analyses were obtained on a CEC 21-103C mass spectrometer which has been updated by using Nuclide Corp solidstate electronics. The Nuclide components include the highvoltage power supply (HV-21), exponential scan generator (ESG-l),trap current regulator (ER-12/103), ion chamber temperature regulator (ST-3), electrometer amplifier/preamplifier (EA-lO/EAH-300),source divider (SC-103),and magnet control with temperature compensated Hall device (MR-13/103). The existing diffusion pump was replaced by a Leybold-Hereaus220 L/s turbomolecular pump for improved source pumping. The only original parts of the spectrometer still in use are the magnet coils, analyzer, and ion source. The spectral data were acquired 0 1982 American Chemlcal Societv
