1983
Anal. Chem. 1980, 52, 1983-1984
Identification of Chrysotile Asbestos by Microdiffraction D. R. Beaman' and H. M. Baker Dow Chemical Company, Midland, Michigan 48640
If the proposed Environmental Protection Agency guidelines ( 1 )concerning the asbestos content of ambient water are used to set limitations for waste water effluents, it is imperative that the accuracy of the analysis methods based on transmission electron microscopy (2-5) be improved. The problem is most severe when analyzing an effluent with high solids and low chrysotile content. In such cases the detection limits are high, each detected chrysotile fiber may correspond to 1 X lo6 to 1 X lo7 fibers/L, and only a few fibers are observed in an analysis requiring 8-12 h. Under such circumstances the interim method proposed by the EPA ( 5 ) is subject to significant error because the chrysotile fibers are identified only on the basis of their morphology and selected-area electron diffraction patterns (SAED). This paper discusses some of the problems associated with the present methodology (2-5) and describes a means of significantly improving the results. Microdiffraction has been found to provide a far better means of fiber identification than selected area electron diffraction when carried out in transmission electron microscopes having relatively clean vacuum systems.
RESULTS AND DISCUSSION While chrysotile morphology is diitinctive, it is not entirely unique and is often compromised or obscured by other solids in the sample ( 2 ) . A group of ten investigators counted chrysotile fibers by using a series of electron photomicrographs with the results shown in Table I (6). The samples were all 50% NaOH and clean when compared with a typical effluent sample. The broad range in counted fibers illustrates the difficulties with morphological identification. Millette et al. have shown that chrysotile and the clay minerals halloysite and polygorskite (7) have similar morphologies. Selected area electron diffraction of chrysotile fibrils is even less reliable. In earlier work ( 2 , 8, 9) it was found that less than 15% of the chrysotile fibrils in a clean water standard provided positive SAED patterns, and the percentage providing positive SAED patterns only reached 50% when there were 4 fibrils in a fiber. Other investigators have encountered similar difficulties. Ampian (10)reported for amphibole-type asbestos that positive identification using SAED was only forthcoming from carefully indexed patterns yielding accurate lattice parameters. Ross (11) found SAED patterns of asbestos minerals difficult to obtain and interpret and that 200 keV was required for distinct patterns. Biles and Emerson (12) reported that most chrysotile fibers in beer did not give identifiable patterns. On the other hand, Samudra (13),using a camera length of 20 cm, reported that 99% of the chrysotile fibers in the size range of 2Ck120 nm (diameter) provided good patterns. Feldman (14) found in four samples of the same reference solution (chrysotile in filtered distilled water) that 68% (58-84% range) of all fibers were diffracting but that only 47% (3646% range) of the fibers with lengths below 0.4 pm were diffracting. In a recent ASTM study of a filtered water sample spiked with chrysotile, the percentage of chrysotile fibers identified on the basis of SAED varied from 5 to 70% and averaged 30% for ten independent laboratories. The results would be far worse for an effluent sample in which many fibers would be coated or in close proximity to other crystalline solids. There are several reasons for the difficulties and variations encountered with SAED identification. A major problem is the lack of a consistent classification scheme. Some inves-
Table I. Counting of Chrysotile Fibers in a Series of Electron Micrographs (6) no. of no. of fibers photosample micrographs rangea mean
A B C a
+la
re1 std devn, %
16 22
7-73
24
19
79
62-121
87
21
18
43-66
54
18 8
15
Minimum and maximum number of fibers counted by
1 0 different investigators.
tigators require fully indexed diffraction patterns (10, 15) for positive identification. This is impractical in an analysis which requires 8-12 h even when identification is based on visual inspection of the diffraction pattern on the fluorescent screen; it is also usually unnecessary in the case of chrysotile where the SAED pattern is distinctive. While there are some patterns that are similar to chrysotile (7, 9), any uncertainties can generally be resolved by obtaining an elemental spectrum with an energy dispersive spectrometer (2,16). The chrysotile pattern is streaked in alternate layer lines, has a characteristic layer line spacing, and exhibits distinctive reflections, e.g., in the second and fourth rows from center. If the positive identification of a fiber is based on the presence of all these features, only a small fraction of chrysotile fibers would be counted (2,8). The problem is that many patterns are initially incomplete and others will fade within 30 s to such an extent as to be unidentifiable. This is an electron beam induced degradation due to dehydroxylization (10) and carbon contamination. Under such conditions an analyst, discouraged by poor diffraction patterns, may relax the criteria for positive SAED identification and classify as positive all fibers providing any indication of crystallinity. In a chrysotile standard where only 10% of the fibrils gave positive SAED patterns, 4(t70% gave indications of crystallinity ( 2 , 8). In the extreme case the analyst tires of poor patterns and begins classifying fibers on the basis of morphology only. Relying only on crystallinity and/ or morphology leads to highly inaccurate results and, in the case of an effluent, can result in the detection of large quantities of asbestos where none is present. These problems, coupled with differences in instruments ( 2 ) and the presence of fiber coatings or interfering crystalline solids, make SAED identification relatively unreliable. The quality of the analyses can be improved by subjecting individual fibers to elemental analysis with an energy dispersive spectrometer (EDS). In modern instruments with large solid angles (30 mm2 detector surface and 15 mm specimen-to-crystal distance) the Mg/Si ratio of a fibril can be acquired in under 30 s. The ambiguities associated with each individual mode can be minimized by basing the fiber identification on the combination of morphology, SAED, and EDS. By use of this approach interlaboratory reproducibility has been better than 20% (8). The diffraction portion of the identification can be improved by using the microdiffraction mode rather than SAED. Microdiffraction was performed on chrysotile standards in the JEOL lOOCX and the Philips EM400T analytical transmission electron microscopes. The former was a diffusion pumped system subjected to rigorous cleaning procedures and the latter had an ion pumped column. Twelve fibrils that did not
0003-2700/80/0352-1983$01.00/0C' 1980 American Chemical Society
1984
Anal. Chem. 1980, 52, 1984-1987
the analysis area rather than the field limiting aperture used in SAED. In SAED the field limiting aperture and the spherical aberation of the objective lens limit the minimum analysis diameter to 0.5-1 pm. In microdiffraction the beam diameter at the sample defines the diffraction analysis diameter which can be as small as 40 nm.
ACKNOWLEDGMENT We wish to thank T. Huber of JEOL, USA, and J. Fahy of Philips Electronics Instruments for making the JEOL lOOCX and Philips EM400T instruments available for this work. We also gratefully acknowledge assistance with the experimental measurements by I. Piscopo of Philips and T. Yoshioka of JEOL.
LITERATURE CITED Figure 1. Microdiffraction pattern from a chrysotile fibril obtained in a JEOL lOOCX instrument at 100 kV
provide good SAED patterns were successively examined in the microdiffraction mode, and a positive pattern was obtained on each (see Figure 1). In another experiment 40 consecutive fibrils gave positive microdiffraction patterns. The patterns were stable and did not fade within the time required to establish diffraction conditions. In two experiments on pattern stability the pattern did not fade in 20 min. This is a significant improvement over SAED and will allow electron diffraction to be effectively utilized in the routine analysis of chrysotile asbestos in water and air samples. In relatively clean water samples, such as tap water and many lakes, an accurate analysis should be possible by using the EPA interim method (5) with SAED replaced by microdiffraction. In relatively unclean waters, such as waste-water effluents and many river samples where a few fibers lead to a large concentration and the fibers may be compromised by coatings and other interfering materials, the identification should be based on morphology, the Mg/Si ratio determined by EDS and the microdiffraction pattern. When this approach is used in conjunction with a sample preparation technique where fiber losses are minimized (carbon-coated Nuclepore) ( 3 , 9),improved accuracy and interlaboratory reproducibility should be possible. Microdiffraction has been referred to as microbeam diffraction (17, 18), focused beam Riecke technique (19), or focused aperture microdiffraction (20). The image and diffraction patterns are formed in the normal manner, but the sample is illuminated with a fine beam of electrons. The fine parallel beam of electrons is formed by using a strongly excited first condenser lens and a small (typically 20 pm) second condenser aperture. The second condenser is focused to provide the minimum spot diameter. The diffraction pattern is formed on the back focal plane of the objective lens, magnified (intermediate lens), and projected (projection lens) as in conventional diffraction. Microdiffraction differs from SAED in that the small second condenser aperture defines
"Asbestos, Ambient Water Quality Criteria". Criteria and Standards Division, Office of Water Planning and Standards, U.S. Environmental Protection Agency: Washington, DC, 1979;Document No. 297-917. Beaman, D. R.; Flle, D. M. Anal. Chem. 1978, 48, 101-110. Cook, P. M.; Rubin, I. 6.; Magglore, C. J.; Nicholson, W. J. Proceedings of Internatlonal Conference on Envronmental Senslng and Assessment; IEEE: Las Vegas. 1976;Sectlon 34-1. Mlllette, J. R. I n "Electron Microscopy of Microflbers"; Asher. I.M., McGtath, P. P., Eds.; Proceedings of the First FDA Office of Science, Summer Symposium, U.S. Government Prlntlng Office: Washington DC, 1976;pp 85-92. Anderson, C. H.; Long, J. M. "Interim Method for Determining Asbestos in Water"; US. Environmental Protection Agency: Washlngton E€, Jan 1980;EPA-600/4-80-005. Summary Report (111) of the Asbestos Methods Task Force, New Orleans, Chlorine Instttute Products Analysis and Speclflcations Committee, Analytical Procedures Subcommlttee, H. Bohmer, Chalrman. Feb 4,
1980. Mlllette, J. R.; Twyman, J. D.; Hansen, E. C.; Clark, P. J.; Pansing, M. F. I n "Scanning Electron Microscopy/l979/1"; Joharl, 0. Ed.; Scanning Electron Microscopy, Inc.: AMF O'Hare, IL, 1979,pp 579-586. Beaman, D. R.; Walker, H. J. NBS Spec. Pub/. ( U . S . )1078, No. 506,
249-269. Beaman, D. R.; Walker, H. J. I n "Electron Mlcroscopy of Microfibers"; Asher, I.M., McGrath, P. P., Eds.; Proceedings of the Flrst FDA Office of Science, Summer Symposium, U.S. Government Printing Offlce: Washlngton DC, 1976;pp 98-105. Arnplan, S. G. I n "Electron Microscopy of Microfibers"; Asher, I.M., McGrath, P. P., Eds.; Proceedlngs of the First FDA Office of Science, Summer Symposium; U S . Government Printing Office: Washington, DC, 1976,pp i2-27. Ross, M. I n "Electron Microscopy of Microfibers"; Asher, I.M., McGrath, P. P., Eds.; Proceedings of the First FDA Offlce of Sclence, Summer Svmooslum: U.S. Government Printing Office: Washlnaton. - DC. 1976, pi, 34-35. Biles, 6.; Emerson, T. R. Nature (London) 1966, 219,93-94. Samudra, A. V. I n "Scanning Electron Microscopy, 1977/1"; Joharl, O., Ed.; Scanning Electron Microscopy, Inc.: AMF O'Hare, IL, 1977, pp
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385-392. Feldman, R . S. USEPA, Cincinnatl, OH, private communicatlon, 1979. Lee. R. J.; Lalb, J. S.; Fisher, R. M. NBS Spec. Pub/. ( U . S . )1978, No.
506,367-402. Beaman, D. R. I n "Environmental Pollutants"; Toribara, T. Y., Coleman, J. R., Dahneke, B. E.,Feldman, I., Eds., Plenum Press: New York, 1977, pp 255-294. Sherman, E. S.:Thomas, E. L. J . Mater. Sci. 1979, 14, 1109-1 113. JEOL News 1077, 15E (No. I), 14-24. Thompson, M. N. Phi/@s Nectfon Optics Bull. 1977, EM110, 31-39. Warren, J. B. I n "Introduction to Analytical Electron Microscopy"; Hren, Joy, D. C., Eds.; Plenum Press: New York, 1979, J. J., Goldstein, J. I., pp 369-385.
RECEIVED for review April 11, 1980. Accepted July 14,1980.
Recovery of Naphthalene during Evaporative Concentration Cecil E. Higgins" and Michael R. Guerin Analytical Chemistry Division, Oak Ridge National Laboratory, Oak Ridge, Tennessee 37830
The analysis of trace organics usually requires concentrating organic extracts to small volumes prior to instrumental analysis. The use of a concentration apparatus employing a nitrogen blanket and reduced pressure is desirable because 0003-2700/80/0352-1984$01 .OO/O
the inert atmosphere and low temperature inherent with the use of such a system helps to ensure stable composition. Unfortunately, diaromatic compounds such as the naphthalenes and biphenyls are frequently almost completely lost 0 1980 American Chemical Society
