monochromator slit width. (Increasingthe lamp current would also reduce UBE.) However, in none of the cases investigated would this produce a significant reduction in the total absorbance noise. (Even if up were reduced to zero, OA would not be reduced by more than 30%in any case.) In addition, increasing the lamp current or slit width would lead to a reduction of the absorbance signal in many cases and consequently have a detrimental effect on the detection limit. Lamp flicker noise can be reduced by using a double beam system to monitor changes in lamp intensity. However, if the lamp duty cycle is unchanged, the double beam system necessitates halving the time of observation of the lamp. This causes up to increase by 4,assuming the light throughput is unchanged. UBE will also increase by 4 because of the halved observation time of the background emission (5). Therefore, neglecting UD and UAR and assuming complete elimination of UL, the noise in the sample channel of the double beam system will be US
= (2Up2
+
UF2
+ 2UBE2)1/2
(4)
where up, U P , and UBE are the standard deviation values for the equivalent single beam system. Assuming the reference beam has the same light throughput and observation time as the sample beam, the noise in the reference channel will be CrR
= 4 UP
(5)
If the two channels are combined by taking the difference in the absorbance signals (or ratio of the %T signals), then the total absorbance noise will be OA
= (4C$
= (ffR2
+
UB2
+ 8s2)1/2 f
2UBE2)’/2
(6)
Comparison with Equation 2 (neglecting UD and UAR) shows that the double beam will have a lower UA for those elements with UL > ( 3 4 U B E ~ ) and ~ / ~a higher U A for all other ele-
+
ments. In either case, the difference would be less than a factor of two with any of the elements studied here. Flame transmission noise should, in theory, be eliminated by a background correction system using a second light source. However, the fluctuation of the flame in both time and space, combined with the problem of optically matching beams from two lamps, means that elimination of flame transmission noise is difficult to achieve. In addition, background correction systems suffer from increased photon noise in the same way as double beam systems. Detection limit comparisons (26) have shown that only for As and Se does background correction produce an improvement. It therefore follows that, since photon noise and lamp flicker noise can often be reduced, the flame provides the major limitation to significantly reduced noise at the detection limit level.
LITERATURE CITED (1) J. T. H. Roos, Spectrochim. Acta, Part 6, 24, 255 (1969). (2) J. T. H. Roos, Spectfochim. Acta, Part 6,25,539 (1970). (3) J. T. H. Roos,,, Spectrochim. Acta, Part 6, 28, 407 (1973). (4) W. J. Price, Analytical Atomic Absorption Spectrometry”, Heyden and Son Ltd., London, 1972, pp 74-77, 103-11 1. (5) J. D. Ingle, Jr., Anal. Chem., 46, 2161 (1974). (6) N. W. Bower and J. D. Ingle, Jr., Anal. Chem., 48, 686 (1976). (7) J. D. Winefordner and T. J. Vickers, Anal. Chem., 36, 1947 (1964). ( 8 ) J. D. Winefordner and C. Veillon, Anal. Chem., 37, 416 (1965). (9) M. L. Parsons, W. J. McCarthy, and J. D. Winefordner, J. Chem. Educ., 44, 214 (1967). (10) J. D. Winefordner, V. Svoboda, and L. H. Cline, CRC Crit Rev. Anal. Chem, August 1970. (11) M. L. Parsons and P. M. McElfresh, Appl. Spectrosc., 26, 472 (1972). (12) L. Erdey, G. Svehla. and L. Koltai, Talanta, IO, 531 (1963). (13) H. Khalifa, G. Svehla, and L. Erdey, Talanta, 12,703 (1965). (14) D. R. Weir and R. P. Kofluk, At. Absorp. Newsl., 6, 24 (1967). (15) 9. Meddings and H. Kaiser, At. Absorp. Newsl., 6 , 28 (1967). (16) W. B. Barnett and J. D. Kerber, At. Absorp. Newsl., 13,56 (1974).
RECEIVEDfor review May 4, 1976. Accepted July 19, 1976. This paper is based on one presented at the Fifth International Conference on Atomic Spectroscopy, Melbourne, Australia, August 25-29, 1975.
Competitive Protein Binding Assay for Biotin Monitored by Chemiluminescence Hartmut R. Schroeder,* Paul 0. Vogelhut, Robert J. Carrico, R. C. Boguslaski, and R. T. Buckler Ames Research Laboratory and Corporate Research, Chemistry Department, Miles Laboratories, Inc., Elkhart, Ind. 465 14
A method for monitoring competitive protein binding reactions by means of chemliumlnescence is described. Biotin Is covalently linked through a bridging group to the amine function of isoluminol to yield the conjugate 6-(3-blotlnyiamido-2hydroxypropylamino)-2,3-dlhydrophthalazlne-l,4-dione. The conjugate emits light when oxidized by elther a hydrogen peroxide-lactoperoxidase system at pH 7.4 or by superoxide anion at pH 8.0. Peak light Intensity is related to conjugate concentration to a lower limit of about 5 nM. Thls intensity increases about 10-fold when the conjugate is incubated with avidin, a blnding protein speclfic for biotin, prior to oxidation. When the conjugate and varying levels of biotin are allowed to compete for a limlted number of avidin binding sites, the peak light Intensity produced In subsequent chemiluminescent reactlonsdecreases wlth increasing levels of biotin. Biotin is measured quantitatively at levels as low as 50 nM.
Competitive protein binding assays are frequently used to determine low levels of important biological substances. Usually, these assays are monitored with radiolabeled ligands. Recently, enzyme cofactors were coupled to ligands and these ligand-cofactor conjugates were determined at low concentrations with enzymic cycling or bioluminescent assays (1,Z). We found that the conjugates were inactive with enzymes when the ligands were bound to their specific proteins. Thus, levels of unbound ligand-cofactor conjugates could be measured without separation from the bound form. In a search for other substances which may serve as labels for ligands, we studied chemiluminescent compounds. Ideally, the compound used as a label would emit light during a simple chemical reaction, and this light could be used for quantitative measurement of the label at nanomolar levels or lower. The aminophthalhydrazides are attractive for investigation because they emit light with relatively high
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
1933
-
0
40 60 CMEMILUMINESCENT COMPOUND (nM)
Figure 2. Peak light intensities produced by lurninol, amino-isoluminol and biotinyl-isoluminol during oxidation by superoxide anion Flgure 1. Reaction
sequence for the synthesis of the biotinyl-isoluminol
conjugate quantum efficiency when they are oxidized. Various oxidation systems have been used: (a) hydrogen peroxide and metal catalyst at alkaline pH (3-5), (b) hydrogen peroxide and horseradish peroxidase at neutral pH (6, 7), and (c) superoxide anion generated during oxidation of hypoxanthine by xanthine oxidase at pH 10 (8,9). Luminol and isoluminol appear to be the best labels for initial work since they are readily available. Attachment of a ligand to these aminophthalhydrazides will undoubtedly influence chemiluminescence efficiency. Introduction of electron-donating substituents on the benzene ring generally increases the light yield (20, I I ) , while alteration of the heterocyclic ring virtually eliminates light emission (12).Even though the light generating ability of isoluminol is less than luminol, we chose to attach the ligand through the amino group of isoluminol. Alkylation a t this position avoids steric problems and enhances chemiluminescence efficiency, whereas amino substitution in luminol decreases the efficiency (13,24). In order to study the feasibility of using isoluminol as a label, it was coupled to biotin through a bridging group. Light emission by biotinyl-isoluminol was measured in the presence and absence of avidin, a binding protein specific for biotin.
EXPERIMENTAL Light Measurements. Chemiluminescent reactions were carried out in 6 X 50 mm test tubes mounted in a DuPont 760 Luminescence Biometer. Each tube contained 150 p1 of phthalhydrazide solution with a composition that varied according to the experiment. A 10-p1 aliquot of the designated oxidant was injected from a 25-pl Hamilton syringe. The peak light intensity occurring during the reaction was measured. In some cases, the progress of reactions was monitored with an oscilloscope connected to the biometer. Total light measurements were made with a custom built instrument. The reaction vessel was placed in a light-tight housing facing the photomultiplier, and light reactions were initiated as described for peak intensity measurements. The output of the photomultiplier (1P21) was measured with a Keithley Electrometer Model 610C. The recorder output was converted into pulses with a Burr Brown VFC12 Voltage to Frequency Converter and counted on a Beckman Universal EPUT Model 7350A for a fixed period of 10 s. Dark current was subtracted from the accumulated counts. Oxidation Systems. H202-Lactoperoxidase. A solution of the chemiluminescent compound in 0.1 M Tris-HC1, pH 7.4,140 p1, was mixed with 10 pl of lactoperoxidase (1mg/ml in the same buffer) and incubated for 2 min a t 25 "C. Then 10 pl of 0.95 mM HzOz in 10 mM Tris-HC1, pH 7.4, was injected to initiate the light yielding reaction. Potassium Superoxide. The chemiluminescent compounds were 1934
Reaction mixtures containing luminoi (A),amino-isoluminol ( 0 )or biotinylisoiuminol (m) in 0.1 M Tris-HCI, pH 8.0, were injected with 10 yl 0.05 M KOn, 0.10 M 18-crown-6 in dimethyiformamide.Concentrations of the chemiluminescent compounds are those in the 150-pl reaction mixtures. Peak light intensities from triplicate reactions were averaged dissolved in 140 pl of 0.1 M Tris-HC1, pH 8.0. Ten pl of dry dimethylformamide (DMF) containing 0.05 M KO2 and 0.10 M 1,4,7,10,13,16-hexaoxacyclooctadecane(18-crown-6) was injected t o initiate chemiluminescence. The 18-crown-6 was added to increase solubility of KO2 in DMF. Specific reaction conditions are described with the results. Reagent Solutions. Isoluminol Derivatives and Luminol. Stock solutions of chemiluminescent compounds were prepared in either dimethylformamide or 0.1 N NaOH or 0.1 M Na2C03 a t p H 10.5. These solutions were diluted in doubly distilled water which had been filtered through charcoal. Aliquots of the dilutions were added to buffered reaction mixtures. Lactoperoxidase. The enzyme (Calbiochem, San Diego, Calif., or Sigma Chemical Co., St. Louis, Mo.) was dissolved a t 1mg/ml in 0.1 M Tris-HC1, pH 7.4. This gave about 20 units lactoperoxidase per ml when assayed by a published procedure (15). Auidin. Lyophilized avidin (Sigma), 1mg, was dissolved in 1ml of 10 mM Tris-HC1, pH 7.4. The binding titer of the solution was not determined. Synthesis of 6-(3-Biotinylamido-2-hydroxypropylamino)2,3-dihydrophthalazine-1,4-dione(Biotinyl-Isolurninol). 4-(3Chloro-2-hydroxypropylamino)-N-methylphthalimide(Figure 1, 1)-Twenty-five grams (0.142 mol) 4-amino-N-methylphthalimide (16) and 20.7 g (0.21 mol) l-chloro-2,3-epoxypropane were added to 150 ml of 2,2,2-trifluoroethanol and the reaction mixture was heated to reflux with stirring for 48 h. Seventy to 80 ml of the solvent was removed by distillation, and a heavy yellow precipitate formed upon cooling to room temperature. This precipitate was digested with ethyl acetate, collected by filtration, and dried to give 29.5 g (77%yield) of 4-(3-chloro-2-hydroxypropylamino)-N-methylphthalimide,mp 136-138.5 "C. Anal. Calcd for C12H13ClN203: C, 53.64; H, 4.88; N, 10.45. Found: C, 53.87; H, 4.85; N, 10.81. 4-[3-(N-Phthalimido)-2-hydroxypropylamino] -N-methylphthalimide (Figure 1, II)-4-(3-Chloro-2-hydroxypropylamino)-Nmethylphthalimide (Figure 1, I), 13.5 g (0.05 mol) and 15.7 g (0.085 mol) of potassium phthalimide (Aldrich Chemical Co., Milwaukee, Wis.) were heated to reflux with stirring in 150 ml dimethylformamide for 24 h. The dimethylformamide was removed in vacuo, and the residue was washed with water and filtered. The yellow solid was dissolved in glacial acetic acid and recrystallized by adding water to give 12.8 g (67% yield) of the bis-imide as fine yellow crystals, mp 247-248.5 "C. Anal. Calcd for C20H17N305: C, 63.32; H, 4.52; N, 11.08. Found: C, 63.16; H, 4.38; N, 10.93. 6-[3-Amino-2-hydroxyropylamino]-2,3-dihydrophthalazine-l,4dione (Figure 1, 111) (amino-isoluminol)-4-[3-(N-Phthalimido)2-hydroxypropylamino]-N-methylphthalimide (Figure 1, 111, 5.0 g (13.2 mmol), 90 ml absolute ethanol, and 35 ml 95% hydrazine (Eastman Chemical Co., Rochester, N.Y.) were refluxed with stirring for 4 h. The solvent was removed in vacuo, and the resulting solid was dried for 24 h under vacuum a t 120 "C. This material was stirred for 1 h with 70 ml of 0.1 N HCl. The insoluble, 2,3-dihydroxyphthalaz-
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
ine-1,4-dione(Figure 1,IV)was removed by filtration,and the filtrate was adjusted to pH 6.5with saturated aqueous sodium bicarbonate. A white precipitate which formed was collected by filtration and dried to give 2.2 g of the product (Figure 1,111)(67% yield). After recrystallization from water, the compound decomposed at 273 "C. Anal. Calcd for CllH14N203: C, 52.79;H, 5.64;N, 22.39.Found: C, 52.73;H, 5.72;N, 22.54. The final steps in the synthesis involved conversion of biotin to a mixed anhydride which was reacted with 111. Biotin (Sigma),0.29 g (1.2mmol), and 0.17 ml triethylamine were dissolved in 20 ml dry dimethylformamideunder anhydrous conditions and cooled to -10 "C. A solution of 0.141 ml of ethylchloroformate in 2.86 ml ether was slowly added, and the reaction was stirred for 30 min. A precipitate formed and was separated by filtration.Next a suspension of 600 mg (2.4 mmol) of 6-(3-amino-2-hydroxypropylamino)-2,3-dihydrophthalazine-1,4-dione(111)in 20 ml dry dimethylformamideand 1 ml dry pyridine was quickly added to the filtrate. This mixture was stirred at -10 "C for 30 min and then at room temperature overnight. AVIDIN ( p g l m l ) During this period, a solution was obtained. The solvent was removed by distillation at 60 "C and 0.10 mm Hg pressure leaving an oily resFigure 3. Effect of avidin on the peak light intensity produced with idue which was stirred with 50 mlO.1 N HC1 for 1 h. A white solid biotinyl-isoluminol. formed which was collected by filtration and washed with 0.1 N HC1 For Curve (A), reaction mixtures containing 84 nM biotinyi-isoluminol and various and then water. After drying in vacuo at room temperature overnight, levels of avidin in 0.1 M Tris-HCI, pH 7.4, were incubated at 25 "C for 15 min. 0.55 g (97% yield) of the amide (Figure 1,V) was obtained, mp 170-3 Then the mixtures were assayed with the H2Op-lactoperoxidase system outline OC(dec).Anal. Calcd for C ~ ~ H ~ ~ N C, SO 52.92; ~ S :H, 5.92;N, 17.64. in the Experimental section. For Curve (B), reaction mixtures containing 42 nM Found: C, 51.69;H, 5.90;N,17.63. biotinyl-isoluminol and various levels of avidin in 0.1 M Tris-HCI, pH 8.0, were RESULTS incubated at 25 "C for 5 min. The oxidation was conducted with superoxide anion as in Figure 2. The reagent concentrations are those in the 150-11 reaction Quantitative Determination of Chemiluminescent mixtures Compounds. Lactoperoxidase, horseradish peroxidase, and
catalase were used to catalyze the oxidation of luminol with H202. Luminol was also oxidized by superoxide anion generated during the xanthine oxidase catalyzed oxidation of hypoxanthine (IO, 11). These oxidation systems were studied in various buffers and at various levels of substrates. Luminol could be detected quantitatively with the best sensitivity and reproducibility with lactoperoxidase and H202 under the reaction conditions outlined in the Experimental section. The peak light intensities increase linearly with luminol concentration, and the lower limit for quantitative measurement was about 0.5 nM. In these reactions, light production reached peak intensity within approximately 0.1 s after addition of H202, and then the signal decayed to background levels within several seconds. Luminol and isoluminol also produced light upon oxidation with Koa. The light reached maximum intensity within 0.1 s after injection of this oxidant. T h e greatest peak light intensity was observed a t pH 10.5, but pH 8 was employed in subsequent studies since this condition was more compatible with protein binding reactions. Peak light intensities obtained with various levels of luminol, amino-isoluminol (Figure 1, 111)and biotinyl-isoluminol (Figure 1,V) are shown in Figure 2, and lower limits for detection of these compounds were approximately 0.5,3, and 5 nM, respectively. Binding Reactions Monitored by Chemiluminescence. Reactions employing 84 nM biotinyl-isoluminol and varying amounts of avidin were assayed with the HzO2-lactoperoxidase system and with a KO2 system. In both cases, peak light intensities increased as avidin levels increased up to 13.7 pg/ml and then declined a t greater concentrations (Figure 3). The total light produced by biotinyl-isoluminol upon oxidation by the HzOz-lactoperoxidase system was also enhanced by avidin to the same extent as the peak light intensity (data not shown). In additional experiments, various levels of biotinyl-isoluminol were incubated with a constant amount of avidin, 13.7 pglml, and assayed with the lactoperoxidase-H202 system. The peak light intensity increased linearly with the concentration of biotinyl-isoluminol up to 150 nM, and the intensities were enhanced by approximately an order of magnitude over identical levels of conjugate oxidized in the absence of avidin (Figure 2 and Figure 4). In control measurements without avidin, a linear response was observed with bio-
BIOTINVL480LUMINOL (nM)
Flgure 4. Peak light intensities produced with various levels of biotinyl-isoluminol in the presence of a constant amount of avidin and in the absence of avidin
Reaction mixtures containing various levels of biotinyl-isoluminol in 140 1 10.1 M Tris-HCI, pH 7.4, and with either 13.7 pg avidin per ml (A)or without avidin (O), were incubated at 25 O C for 5 min. These mixtures were assayed with the H202-lactoperoxidase system described in the Experimental section. Concentrations of reagents are those in 150-pl reaction mixtures. Data points are averages of triplicate assays
tinyl-isoluminol up to 350 nM (Figure 4).Similar results were obtained in three separate experiments. Further investigations were undertaken to determine whether the enhancement of peak light intensity by avidin was due to the specific binding of the biotin residue in biotinylisoluminol. Avidin was allowed to react with biotin and then various amounts of this complex were added to 84 nM biotinyl-isoluminol. The peak light intensities measured with the H~O~-lactoperoxidasesystem decreased slightly as the level of the avidin-biotin complex was increased (Table I). Since biotin is irreversibly bound to avidin under conditions of this experiment (I7), these results demonstrate that the enhancement of peak light intensities seen in Figures 3 and 4 was mediated through the biotin binding sites of avidin. In an additional control measurement, the peak light intensity produced with 64 nM aminc-isoluminol was enhanced slightly by avidin (Table 11); however, the effect was small
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
1935
Table 1. Effect of Biotin-Avidin Complex on Peak Light Intensity Produced with Biotinyl-Isoluminola BiotinylPeak isoluminol, Biotin, Avidin, light * nM PM fig/ml intensity 0
84 84 84 84 84
0
0.67 1.33
4.7 3.9 4.0 2.8 2.9
13.7 27.9 41.1 68.5
2.00 3.33
a A solution containing 10 pM biotin and 0.2 mg avidin in 0.1 M Tris-HC1,pH 7.4, was incubated for 2 h at 25 "C. This solution was used to prepare reaction mixtures (140 pl) which were incubated at 25 O C for 5 min. Then the oxidation reaction was carried out with the H202-lactoperoxidase system outlined in the Experimental section. The reagent concentrations are those in the 150-pl reaction mixture prior to addition of H202.
ElOllN (nu)
Figure 5. Competitive binding reactions monitored with the
H202-
lactoperoxidase system
Table 11. Effect of Avidin and Biotin on Peak Light Intensities Produced by Biotinyl-Isoluminol and AminoIsohminola BiotinylAminoPeak Biotin, isoluminol, isoluminol, Avidin, light FM nM nM fig/ml intensity 0 0 0 0 0 0 0
0 0 0 0 0 84 84
0
4 4 4
84
0
4 1.3
84 84
0 0 64 64 64 0
0 0 0 0 0 0
0 13.7
0 13.7 68.5 0 13.7
0 0 13.7 13.7 13.7
0.8 0.9 4.1 4.9 4.6 1.9 25.3 0.8 2.2 0.9 6.1 10.4
a The binding reactions were prepared in 140 ~ 1 0 .M 1 Tris-HC1, pH 7.4, (avidin was added last) and incubated at 25 "C for 5 min. Then the oxidation reaction was conducted with the HzO2-lactoperoxidase system outline in the Experimental section. Concentrations of reagents are those in the 150-p1reaction mixture prior to addition of H202.
compared to that seen in Figure 3 with the biotinyl-isoluminol. Thus, specific binding of the biotinyl residue to avidin is essential for the enhancement observed with biotinyl-isoluminol. Competitive protein binding reactions were conducted with various levels of biotin, 8 4 nM biotinyl-isoluminol and 3.4 pg avidin per ml. T o ensure that the results obtained with these reactions were due to specific binding of the biotinyl residue to avidin and were not attributable to interference by high levels of protein, less than the optimum level of avidin (Figure 3) was employed. When these reactions were assayed with the HzOz-lactoperoxidase system, the peak light intensity decreased as the biotin concentration increased (Figure 5). In control measurements, biotin had only a slight effect on peak light intensities produced with biotinyl-isoluminol in the absence of avidin (Table 11). In other experiments, competitive protein binding reactions similar to those described above were assayed with the superoxide system. Peak light intensities obtained with the assay were dependent upon the biotin concentrations in the binding reactions (data not shown). Both oxidation systems allowed 1936
* ANALYTICAL CHEMISTRY,
Reactions containing various levels of biotin, 90 nM biotinyl-isoluminol and 3.65 fig avidinlrnl (added last) were prepared in 140 fil 0.1 M Tris-HCI, pH 7.4, and incubated at 25 OC for 5 min. Then oxidation with the HzO2-lactoperoxidase system was conducted as described in the Experimental section. The data points are averages from triplicate assays. Biotin levels on the abcissa are those in the 150-pi reaction prior to oxidation
measurement of biotin to a lower limit of about 50 nM in competitive binding assays.
DISCUSSION We have been investigating methods for monitoring competitive protein binding reactions with ligands labeled with nonradioactive substances (1,2).The cofactors NAD and ATP were coupled to ligands in the earlier studies, and these ligand-cofactor conjugates were detected a t low levels by means of enzyme catalyzed reactions. In some applications these methods may be subject to interference by endogenous cofactors and degrading enzymes found in biological materials. Therefore, we investigated the possibility of employing chemiluminescent labels since they are not found in biological systems and they can be detected a t low levels (Figure 2). In earlier studies, we found that cofactors and substrates conjugated to ligands were inactive with enzymes when the ligand moieties were bound to their specific proteins (1,2). Therefore, we were surprised to find that avidin enhanced the peak light intensity produced with biotinyl-isoluminol on oxidation by HzOz and lactoperoxidase (Figure 3). This enhancement is not due to a unique property of the enzyme catalyzed reaction since similar results were observed when superoxide anion was employed as oxidant (Figure 3). It is likely that the enhancement is due to increased chemiluminescence efficiency since avidin increased total light intensity to the same extent as the peak light intensity. The enhancement appeared to require the specific protein-ligand interaction since avidin had a relatively small effect on light production by amino-isoluminol (Table 11)and biotin decreased the peak light intensity in competitive protein binding reactions (Table I1 and Figure 5 ) . The enhancement of peak light intensities by avidin reached a maximum and then diminished a t higher levels of the protein (Figure 3). When increasing amounts of biotinyl-isoluminol were oxidized in the presence of avidin at the level giving the maximum peak light intensity in Figure 3, the peak light intensities increased in a biphasic manner (Figure 4). The biotinyl-isolumino1:avidin ratio at the break in the curve (Figure 4) is about twice the ratio observed a t the top of the curve in Figure 3. These results indicate that avidin enhances the peak light intensity and also interferes with the oxidation to cause some diminution of the intensity. At low
VOL. 48, NO. 13, NOVEMBER 1976
levels of avidin, the enhancement was observed while at high levels the interference gains importance. This conclusion is supported by the observation that the biotin-avidin complex decreases the peak light intensity produced with biotinylisoluminol (Table I). Since avidin enhanced the light produced by biotinyl-isoluminol, a competitive binding assay for biotin was carried out without separation of free and protein bound conjugate. Although elimination of the separation step simplified the assay, this approach decreases sensitivity because light produced by free biotinyl-isoluminol contributes substantial background to the measurements (Figure 5). In cases where sensitivity is a problem, a heterogeneous assay (one involving separation of protein bound from unbound conjugate) could be employed. Also, sensitivity may be improved by: (a) utilizing total light rather than peak light intensity measurements, (b) choosing more efficient chemiluminescent labels, and (c) employing other conditions and oxidants for the chemiluminescent reactions. The present assay for biotin illustrates the feasibility of monitoring competitive protein binding systems with chemiluminescent labels. The work also demonstrates the following advantages over conventional methods for monitoring these types of assays: (a) the assay can be conducted without separation of free and protein-bound conjugate, (b) the labeled ligands are relatively easy to synthesize and characterize, and (c) the inconvenience of handling radioactive materials is eliminated.
ACKNOWLEDGMENT We thank E. 0. Snoke and W. G. Umbarger for excellent technical assistance.
LITERATURE CITED (1) R. J. Carrico, J. E. Christner, R. C. Boguslaski, and K. K. Yeung, Anal. Biochem., 72, 271-282 (1976). (2) H. R. Schroeder, R. J. Carrico, R. C . Boguslaski, and J. E. Christner, Anal. Biochem., 72, 283-292 (1976). (3) U. lsacsson and G. Wettermark, Anal. Chim. Acta, 68, 339-362 (1974). (4) E. H. White, 0. Zafiriou, H. H. Kagi, and J. H. M. Hill, J. Am. Chem. Soc., 86,940-941 (1964). (5) P. D. Wildes and E. H. White, J. Am. Chem. SOC., 95, 2610-2617 (1973). (6) M. J. Cormier and P. M. Prichard, J. Biol. Chem., 243, 4706-4714 (1968). (7) P. M. Prichard and M. J. Cormier, Biochem. Biophys. Res. Comm., 31, 131-136 (1968). (8) G. M. Oyamburo, C. E. Prego, E. Prodanov, and H. Soto, Biochim. Biophys. Acta, 205, 190-195 (1970). (9) E. K. Hodgson and I. Friedovich, Photochem. Photobiol., 18, 451-455 (1973). (10) R. E. Brundreti and E. H. White, J. Am. Chem. SOC., 96, 7497-7502 (1974). (11) H. D. K. Drew and F. H. Pearman, J. Chem. Soc., 586-592 (1937). (12) H. D. K. Drew and R. F. Garwood, J. Chem. SOC., 1841-1846 (1937). (13) A. Spruit-VanDerBurg, Recueil, 69, 1536-1544 (1950). (14) K-D. Gundermann and M. Drawert, Chem. Ber., 95,2018-2026 (1962). (15) M. Morrison, in "Methods in Enzymology", XVII-A, pp 653-657, Assay 2 (1970). (16) H. D. K. Drew and F. H. Pearman, J. Chem. Soc., 26-37 (1937). (17) N. M. Green, Biochem. J., 89, 599-609 (1963).
RECEIVEDfor review June 30, 1976. Accepted August 18, 1976.
Sub-Nanosecond Time-Resolved Rejection of Fluorescence from Raman Spectra J. M. Harris, R. W. Chrisman, F. E. Lytle," and R. S. Tobias" Department of Chemistry, Purdue University, West Lafayeffe, Ind. 47907
A sub-nanosecond time-resolved Raman spectrometer has been constructed using a mode-locked cavity-dumped argon-ion laser as the excitation source and a sampling oscilloscope as the gated photon counter. To test the performance of the instrument, Raman spectra were obtained for solutions of benzene doped with the fluorophors acridine orange (7,= 4.5 ns) and rubrene (7,= 15.6 ns). The magnitude of the background rejection was determined by comparing the level of fluorescence in a time-resolved experiment to that for continuous excitation using the 992 cm-' scattering peak of the solvent as an Internal standard. The resulting Raman to fluorescence enhancements were 34 and 115 for the fast and slow emitters, respectively. These numerical values are shown to compare quite closely to those predicted by theory.
The lower limit of detection in Raman spectrometry is often determined by residual solution fluorescence. This unwanted signal is, in general, difficult to eliminate because of the huge difference in the cross sections for the scattering and emission processes. Naturally, the problem has been aggravated by the ultra-sensitive instrumentation developed to gather vibrational data from dilute solutions. As a typical example, the spectrometer system used in this study can obtain the high resolution spectrum of an efficient fluorophor when present at nanomolar levels. In addition, the possibility of interference is enhanced by the process of thermally assisted excitation.
Thus, a concentrated impurity having a long wavelength absorption band as far as 2400 cm-l to the blue of the laser frequency can generate a substantial background. The two most common approaches to minimizing this artifact have utilized excited state properties of the emitting impurity. In the first of these, quenching, the solution has added to it a foreign agent capable of depleting the fluorescent state via a highly favorable dark reaction (1).In the second, drenching, the impurity is removed via copious quantities of laser radiation coupled with any photochemical pathway that may be present (2). Both of these schemes are rather hit or miss in their nature and are potentially deleterious to the sample molecules themselves. A third general approach recently receiving attention generates the Raman related signal in a collimated beam. Two examples would be coherent antiStokes Raman spectrometry ( 3 ) and inverse Raman spectrometry ( 4 ) . In these techniques, the vibrational data are spatially separated from the background by virtue of the l l r 2 drop in the fluorescence intensity. Both methods, while highly promising, are in their early stages of development and suffer from a signal intensity dependent upon the electric field strength of more than one laser. The approach used in this work is based on the difference in the temporal behavior between scattered and emitted photons. That is, if the sample were to be irradiated by a short-duration light pulse, the Raman signal would follow the intensity-time profile of the excitation whereas the fluorescence would be extended to longer times. The fact that most
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
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