fied to accommodate this size sample by taking a sample containing 0.72 mg. of calcium, using one fifth the amount of each reagent, and diluting to a volume of 10 ml. Standards are made in the range of 2 to 30 y of magnesium. EXPERIMENTAL
The spectrum of the magnesium dye lake in the presence of 3.6 mg. of calcium, 2 ml. of EDTA, and 50 y of magnesium was obtained from 500 to 600 mp us. a blank containing no magnesium. A plateau was obtained from 546 to 550 mp a t the maximum; therefore, a wave length of 548 mp was used throughout the study. Although the reagents have been added separately in previous procedures (3, 5 ) , satisfactory results a e r e obtained by adding a mixture of all the reagents except the sodium hydroxide. Stored in the refrigerator, this mixed reagent was stable for a t least a week. By using standards simultaneously with each group of samples, effects of temperature variation or time of standing were eliminated. Samples made up one day can be read the next if desired, as the magnesium dye lake is stable for over 24 hours if made with poly(viny1 alcohol) solution which has not been stored longer than 1 month. Magnesium is complexed by EDTA only a little less firmly than calcium ( I ) . It is necessary, therefore, to keep the calcium concentration approximately the same in all solutions. This is easily done, as calcium is usually determined
on samples requiring analysis for magnesium. If too little calcium is present in any solution, some of the magnesium will be complexed by the EDTA causing low results. However, a deficiency of calcium up to about 5y0 causes little difference in the readings. With 50 y of magnesium, the same absorbance was obtained with 3.6 to 3.45 mg. of calcium in the solution. With 3.24 mg. of calcium, however, the absorbance was lower by 39y0‘,. Because the calcium content of most samples of the ash of bone, dentin, and enamel is fairly constant, usually little error will be caused by assuming the calcium to be 36% or more in the case of normal tissues. A complete ashing of samples is essential, because small amounts of carbon or carbon compounds, by absorption or complesing of the magnesium, caused low results. Samples were usually ashed overnight at 600” C. with the furnace door ajar t o allow access of oxygen. Interfering Metals and Their Removal. Most samples of mineralized tissue contain no trace metals in amounts large enough to interfere. However, if present, they must be removed. Incisors of rats kept in galvanized iron cages may pick up considerable amounts of zinc in tooth enamel due to gnawing on the wire mesh. This may be removed by extraction with diphenylthiocarbazone (O.lyoin chloroform) at pH 5 . The escess dithizone is removed by extracting nith several portions of chloroform.
Small amounts of chloroform left in the aqueous layer are removed by boiling the solution with a few drops of hydrochloric acid, added to keep the calcium and magnesium phosphates in solution. Iron present in rat incisor enamel is removed by the same procedure. Reproducibility and Recovery. Ten samples containing 5 y of magnesium were made up as in Procedure B. Absorbance compared to the blank gave a mean of 0.061 with R standard deviation of &0.003. Knoii n amounts of magncsium were added to two different bone samples (using Procedure A) with an average recovery of 101.1 f 2.3% in one case and 99.7 f 0.2y0 in the othrr (Table I). Comparison with Gravimetric Method. Table I1 shows the results of analyses of several mineralized tissues by Procedure A, Procedure B, and the gravimetric method. LITERATURE CITED
(1) Bjerrum, J., Schwarzenbach, G., Sillen, L. G. (com ilers), “Stability Constants of Metal-!on Complexes,” Part I,
“Organic Ligands,” Special Publ. 6, Chemical Society, London, 1957. ( 2 ) Hillebrand, W. F., Lundell, G. E. F., Bright, H. A., Hoffman, J. I., “Applied Inorganic Analysis,” 2nd ed., pp 626, 638, Wiley, New York, 1953. (3) Kenyon, 0. A., Oplinger, G., AXAL CHEM.27, 1125 (1955). (4) Mann, C. K., Yoe, J. H., Zbid., 28, 203 (1956). (5) Young, H. Y., Gill, R. F., Ibid., 23,751 (1951). RECEIVEDfor review hIarch 16, 1959. Accepted July 30, 1959.
Estimation of Small Amounts of Fluoride in Body Fluids HARRY W. LINDE’ Reseorch loborotories, Ohio Chemical and Surgical Equipment Co., Division of Air Reduction Co., Inc., Murray Hill,
b Submicrogram amounts of fluoride ion may b e estimated directly in body fluids or water by measuring their inhibitory effect on the enzymatic hydrolysis of ethyl butyrate. In blood or urine, the sensitivity is about 0.1 y per ml. and in water about 0.1 y per 100 ml. The reproducibility is about 0.1 to 0.2 y. The procedure is useful for determining fluoride ion in the presence of compounds containing fluorine in nonlabile organic combination, as it does not require sample ashing, which might decompose these compounds. Substances which interfere with enzyme activity or which a r e excessively buffered will interfere with the method. 2092
ANALYTICAL CHEMISTRY
T
current use of fluorinated organic compounds in medicine makes it desirable to have a method to determine the possible formation of fluoride ion within the body from these compounds. The method recently described by Singer and Armstrong (9) extends the sensitivity and accuracy of the older methods for determination of fluorine in biological material (2) to the point where they would be suitable for the desired analyses, Their procedure, however, involves alkaline ashing of the specimen prior to analysis with the attendant hazard of the production of fluoride ion by the rupture of carbon-fluorine bonds. A method was sought which would not reHE
N. 1.
move fluorine from organic combination, which could be used directly in the presence of biological materials, and which would be sensitive a t the normal level of fluoride in the fluids of the body, roughly 0.1 to 0.5 y per gram (6). Of the methods reviewed by MeKenna (7), Busch, Carter, and MeKenna ( J ) , and Ehing, Horton, and the techniques based on the Willard (4, ability of fluoride ion to slow the enzyme-catalyzed hydrolysis of esters (1,6, 8, 10) appeared to be most suitable. The procedure based on the liver
* Present addreas, Department of Anesthesiology, Hospital of the University of Pennsylvania, Philadelphia 4, Pa.
Table 1.
Specimen Control
A B
\
C
\
... S L O O D
D
--URINE
E "U
,
53 PER C E N T
Analysis
F- Added,
of Blood and Urine Samples
Y
Acid Produced4 Blood
Per Cent Hydrolysis
Fluoride, y/Ml.
0.5 2.5 4.0 0 0 0 0 0 0 0 0 0 0 0 0
4.38 3.38 2.93 5.85 5.71 6.09 6.33 4.56 5.13 6.50 6.53 5.60 5.57 5.76 5.86
76.0 58.5 50.7 101.1 98.9 105.1 109.4 79.0 88.8 112.3 113.0 97.0 96.5 99.7 101.2
0.5 2.5 4.0
Control Control b
b
0.42 0.19 b b
0.10 0.10 0.08 b
Urine
IO0 HYDROLYSIS
Figure 1 . Calibration curves fluoride in blood and urine
0.5 2.5 Control 4.0
for
lipase-catalyzed hydrolysis of aliphatic esters (1,6,10) was thought best. The method described here, which is based on the work of Loevenhart, Peirce, and Amberg (1, 6), involves incubating a mixture of the sample, ethyl butyrate, and water with the enzyme, liver lipase. lit the end of the incubation period, the butyric acid liberated by the enzymatic hydrolysis of ethyl butyrate is titrated potentiometrically with dilute alkali. Six samples may be analyzed in duplicate in about 3 hours of an analyst's time and a total elapsed time of about 7 hours. The method should be applicable to fluorides in the presence of other substances which do not interfere with the enzyme or the ester and which are not excessively acidic. basic, or buffered. REAGENTS
LIVER LIPASE. Place 50 grams of fresh beef liver (frozen liver may be used) in a Waring Blendor with 200 ml. of distilled water and homogenize for about 5 minutes, Filter the liver suspension through 5 thicknesses of clean cheesecloth into a 500-ml. bottle, add 5 ml. of toluene as a preservative, and dilute to 500 ml. Incubate a t 35" to 40' C. for 24 hours to coagulate the solid matter and store the suspension in a refrigerator. To use, filter the amount necessary through medium porosity filter paper (Whatman No. 40). The unfiltered suspension, when stored in a refrigerator, retains enzymatic activity for a t least one month. ETHYL +BUTYRATE. Boiling point 119-121" C., Eastman No. 374. SODIUMFLUORIDE SOLUTION.The solution contains 0.5 y of fluoride ion per ml. Prepare a stock fluoride solution containing 0.1 mg. of fluoride ion per ml. (0.221 gram of sodium fluoride per lo00 ml.). Dilute 5.0 ml. of this stock solution to 1000 ml. immediately before each use.
A
B C D E a I,
0 0 0 0 0 0 0 0 0 0 0
3.91 3.83 4.34 3.58 3,63 2.95 3.24 3.83 4.27 3.75 3.85
Control
98.1 96.3 109.0 89.5 91.2 74.2 81.5 96.3 107.1 94.2 97.2
0.06 b
0.09 0.08 0.23 0.15 0.06 b
0.07 0.06
As milliliters of 0.054N sodium hydroxide. Less fluoride than control sample.
PROCEDURE
To a series of 250-ml. glass-stoppered Erlenmeyer flasks (two flasks for each sample to be analyzed), add 100 ml. of distilled water. To three other flasks add 1.0, 5.0, and 10.0 ml. of a 0.5 y per ml. solution of fluoride and distilled water to each to make 100 ml. Add 1.0 ml. of the control sample of blood or urine to each of the flasks containing only water, and a 1-ml. aliquot of each sample to each of two flasks. Add 0.10 ml. of ethyl butyrate to all flasks, stoppering the flasks immediately after this addition. A t 4minute intervals, add 1.0 ml. of liver lipase to the flasks, mix, and incubate in a thermostat a t 25" f 0.5" C. for 5 hours. Following the incubation, remove the flasks in order one a t a time, immediately add 10 ml. of 95% ethyl alcohol, and titrate potentiometrically to pH 7.7 with 0.05N alkali. To correct for acidity in the sample or reagents, add 100 ml. of water, 10 ml. of ethyl alcohol, 1.0 ml. of the control sample, 0.10 ml. of ethyl butyrate, and 1.0 ml. of liver lipase to a flask; mix, and then titrate immediately to pH 7.7 with alkali. If the samples are excessively acidic or basic, they should be neutralized prior to incubation. The amount of fluoride ion may be calculated as follows:
Per cent hydrolysis
=
-x 100
A - B C - D
where
A
acid present in standard or unknown after hydrolysis. B = acid present in standard or unknown prior to hydrolysis. C = acid present in control after hydrolysis. D = acid present in control prior to hydrolysis. =
The per cent hydrolysis is inversely proportional to the logarithm of the fluoride ion concentration. Standards are prepared by the addition of known amounts of fluoride to portions of the control sample. Alternatively, the amount of acid formed with known additions of fluoride to the control sample may be plotted directly against the logarithm of the added fluoride ion concentration in these samples. Then the line may be extrapolated to the amount of acid produced by the control sample itself to give an estimate of the fluoride in the control sample. RESULTS
To test this procedure, six blood (heparinized) and six urine specimens were taken on four consecutive days VOL. 31, NO. 12, DECEMBER 1959
2093
from a subject who had no known exposure to fluorine compounds over the period of study. Using the specimens taken first as controls, the other five blood and urine specimens were analyzed in duplicate as described above. The results are tabulated in Table I and the standard curves are plotted in Figure 1. About 50 other samples in which there had been exposure to a fluorinated organic compound were tested, with similar reproducibility. The specific data are not av~.ilablefor publication. The standard deviation of all the blood analyses (excluding the control) taken as a group was 0.18 y of fluoride, and that of all the urine analyses, 0.07 y . The standard deviation of duplicate blood analyses was 0.09 y and that of duplicate urine analyses, 0.02 y. As the body stores considerably larger amounts of fluoride in the bones and teeth than are present in the body fluids, the fluoride concentrations in the blood and urine might change over a period of days in the absence of any exposure to fluorine. The true standard deviation of the method, as applied to body fluids, probably lies somewhere in the range covered by the above figures. The actual calculation of the standard deviations was complicated because it was impossible t o assign numerical values to the coneentration of fluoride ion in all cases. Therefore, the calculations were made using the values for per cent hydrolysis and then converting these values to amounts of fluoride ion by means of the standard curves. DISCUSSION
The enzymatic method for the determination of fluoride ion lacks the reproducibility of the method described by Singer and Armstrong (9), although it is somewhat more sensitive for blood analysis. This modest increase in sensitivity is of no great advantage except where limited amounts of blood are available, as from infants or small laboratory animals. The main virtue of the enzymatic method is that it does not involve a n alkaline ashing step, thus avoiding the production of fluoride ion from the decomposition of organic fluorine compounds. This technique will permit determination of fluoride ion in the presence of organic compounds containing
2094
ANALYTICAL CHEMISTRY
nonlabile fluorine atoms, and in this respect, supplements the method of Singer and Armstrong. In addition, the enzymatic method is exceedingly sensitive in simple aqueous Solutions, being able to estimate fluoride ion concentrations of about 0.001 p.p.m. The reagents needed for the enzymatic method are easily prepared and require no elaborate purification before use. A high degree of technical skill is not necessary to carry out the anal3 ses. The procedure given here is similar to that of Loevenhart et a / . ( I , @ , evcept for the addition of ethyl alcohol as suggested by SchlamoFFitz (8) and the shortened incubation timp. Ethyl alcohol is added before the titration to inhibit further enzymatic hydrolysis, as hydrolysis is still proceeding a t the end of the incubation period. Its rate is increased as the pH is increased toward neutrality during the titration (1.6). Both blood and urine contain buffers (bicarbonate, phosphates, organic acids, etc.) and substances which either hydrolyze ethyl butyrate or aid in the enzymatic hydrolysis of this ester. To minimize these effects, a large dilution was used in preparing the samples for analysis as brief trials in more concentrated solutions gave poor results. The incubation period was chosen for convenience. Very long incubation periods of 24 hours or longer tend to lessen the difference between sample and control (1, 6 ) , and short periods of to 1 hour tend to magnify errors, because the amount of acid produced is small. I n dilute neutral aqueous solutions, ethyl butyrate hydrolyzes only very slowly. In 11 days at 25” C. the hydrolysis of a mixture of 0.1 ml. of ethyl butyrate in 5 ml. of water produced only about 0.005 meq. of the acid. The addition of ester-splitting enzymes such as liver lipase speeds this hydrolysis remarkably. The addition of 1 ml. of lipase solution to 2 ml. of water and 0.1 ml. of ethyl butyrate produced 0.06 meq. of acid in only 1 hour. ?’he acidity formed tends to slow this reaction-Le., the enzymatic activity is p H sensitivebut does not stop it. The addition of microgram quantities of fluoride inhibits the hydrolysis markedly. For example, a solution with no fluoride produced about 0.1 meq. of butyric acid in 5 hours while one with 3 y of fluoride produced only about 0.01 meq. in the same period. The extent of the enzymatic hydrol-
ysis is a function of many factors other than the concentration of fluoride ion. Because of the number of potential variables (time, incubation temperature, etc.), they Fere not exhaustively investigated. Rather, a suitable set of conditions was chosen empirically and rigidly adhered to. Khile it is not necessary to duplicate exactly the time, temperature, and amounts of reagents given here, the conditions chosen should be followed strictly. A set of standards, made up in the presence of a fluoride-free control sample, should be carried through with each analysis of a group of samples. I n the eases of blood and urine, standards must be prepared in solutions containing appropriate blood or urine controls as substances present in both of these fluids increase the hydrolysis of ethyl butyrate. ACKNOWLEDGMENT
The blood and urine specimens were obtained through the kindness of Max S. Sadove and Reuben C. Balagot of the University of Illinois Research and Educational Hospitals, Chicago, Ill. LITERATURE CITED
(1) Amberg, S., Loevenhart, A. S.>J . Biol.
Chem. 4, 149 (1908). ( 2 ) Association of Official Agricultural
Chemists, Washington, D. C., “Official Methods of Analysis,” 8th ed., p. 415. (3) Busch, G. W., Carter, R. C., McKenna, F. E., “Analytical Chemistry of the Manhattan Project,” C. J. Rodden, ed., National Nuclear Energy Series, Division VIII, Vol. I, p. 226, McGrawHill, Kew York, 1950. ( 4 ) Elving, P. J., Horton, C. A., WilI:Lrd, H. H., “Fluorine Chemistry,” J. H. Simons, ed., p. 53, Academic Press, New York, 1954. ( 5 ) Gettler, A. O., Ellerbrook, L., Am. J . M e d . Sci. 197, 625 (1939). (6) Loevenhart, A. S., Peirce, G., J . Biol. Chem. 2 , 397 (1907). ( 7 ) McKenna. F. E.. Nucleonics 8. 24 f1951): 9.40.51 (1951). (8) Schlamowih, ilk, U. S . Atomic Energy Comm. (Manhattan Project) Report M1718, May 25, 1945, cited in National Nuclear Energy Series, Division VI, “Pharmacology and Toxicology of Uranium Compounds,” C. Voegtlin, H. C. Hodge, eds., Vol. 1, p. 183, McGraw-Hill, New York, 1949. (9) Singer, L., Armstrong, W.D., ANAL. CHEM.31, 105 (1959). (10) Stetter, H., Chem. Ber. 81,532 (1948). \
,
-
\ - - - - , I
I
RECEIVED for rsview December 6. 1058. Accepted July 30, 1959.
